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Nuclear receptors, like other transcriptional activators, switch on gene transcription by recruiting a complex network of coregulatory proteins. Here, we have identified the arginine methyltransferase PRMT1 as a coactivator for HNF4, an orphan nuclear receptor that regulates the expression of genes involved in diverse metabolic pathways. Remarkably, PRMT1, whose methylation activity on histone H4 strongly correlates with induction of HNF4 target genes in differentiating enterocytes, regulates HNF4 activity through a bipartite mechanism. First, PRMT1 binds and methylates the HNF4 DNA binding domain, thereby enhancing the affinity of HNF4 for its binding site. Second, PRMT1 is recruited to the HNF4 ligand binding domain through a mechanism that involves the p160 family of coactivators and methylates histone H4 at arginine 3. This, together with recruitment of the histone acetyltransferase p300, leads to nucleosomal alterations and subsequent RNA polymerase II preinitiation complex formation.
Control of transcription by RNA polymerase II (Pol II) involves regulated formation of multiprotein complexes at the promoter and enhancer regions of the target genes. Nucleation of these dynamic complexes is largely dictated by site-specific DNA-binding transcription factors (activators and repressors) that are in turn subject to cellular, developmental and environmental cues (Roeder, 2005).
Transcriptional activators, including members of the large nuclear receptor superfamily, recruit a series of coactivators that serve both to penetrate the chromatin in the vicinity of the promoter and to directly facilitate the entry of Pol II and its associated general transcription factors to generate the transcriptionally active preinitiation complex (PIC) (Rochette-Egly, 2005; Roeder, 2005). Coactivators acting at the level of chromatin include both the ATP-dependent chromatin remodelling factors and enzymes that carry out covalent modifications (e.g. acetylation, phosphorylation and methylation) of specific residues in nucleosomal histones.
The hepatocyte nuclear factor (HNF4) is an important regulator of animal physiology (Sladek, 1993; Sladek et al., 1990) and plays a key role in development (Chen et al., 1994) and in hepatocyte and intestine differentiation (Watt et al., 2003). In the adult mammal, HNF4 is expressed in liver, intestine, kidney and pancreas (Sladek, 1993; Sladek et al., 1990) and is responsible for the expression of many genes that control glucose and lipid metabolism (Hayhurst et al., 2001). Relevant to this study, HNF4 also plays a role in the regulation of apolipoprotein AI (apoAI) expression, a major component of HDL (Malik, 2003).
HNF4 displays a typical nuclear receptor domain organization that includes a DNA-binding domain (DBD) with a conserved double zinc finger motif and two activation functions designated AF-1 and AF-2 (Hadzopoulou-Cladaras et al., 1997; Sladek et al., 1990). While the AF-1 domain covers a small N-terminal region, the AF-2 domain (extending from amino acid residues 128 to 366) is more complex and, based on both in vitro (Malik and Karathanasis, 1996) and in vivo (Hadzopoulou-Cladaras et al., 1997) studies, is the major activation domain. Although the putative HNF4 ligand binding domain (LBD) is structurally homologous to those of other receptors (Hadzopoulou-Cladaras et al., 1997; Sladek et al., 1990) and despite sporadic reports of fatty acid associations with HNF4 (Dhe-Paganon et al., 2002; Hertz et al., 1998; Wisely et al., 2002), no definitive ligand has yet been identified.
Several coactivators have been implicated in HNF4 function via the AF-2 domain. These coactivators include the closely related histone acetyl transferases (HATs) CBP and p300 (Dell and Hadzopoulou-Cladaras, 1999; Malik et al., 2002; Wang et al., 1998; Yoshida et al., 1997), members of the p160 family of coactivators (SRC-1 and GRIP-1 (Wang et al., 1998) and the Mediator complex (Maeda et al., 2002; Malik et al., 2002). This diversity of nuclear receptor coactivators suggests that they function in a synergistic fashion. However, the precise sequence of the underlying events remains unclear.
Here we identify PRMT1 as a new coactivator for HNF4 that plays a dual role in modulating HNF4 transcriptional activity: at one level it regulates HNF4 DNA binding activity; at another it methylates histones and acts synergistically with other HNF4 coactivators at the HNF4 target promoters.
To identify potential coactivators involved in HNF4-dependent activation of its target genes in their natural cellular milieu, we have utilized a model system based on the ability of CaCo-2 cells to differentiate into enterocytes and to express HNF4 upon reaching confluence (Soutoglou and Talianidis, 2002). Accordingly, levels of HNF4 dramatically increased after CaCo-2 cells reached confluence (Fig. 1B) and apoAI and another previously characterized HNF4 target gene α1-antitrypsin (α1-AT) (Soutoglou and Talianidis, 2002) were induced (Fig. 1A), while levels of β-actin remained constant.
In ChIP analyses of cells sampled at various times after reaching confluence (Fig. 1C), HNF4 could be detected at both the apoAI and α1-AT promoters by day 2. Its occupancy reached maximal levels by day 6, in agreement with the corresponding protein levels (Fig. 1B). Importantly, HNF4 occupancy correlated well with an increase in R3 methylation of histone H4 (H4) of promoter associated nucleosomes at both the apoAI and α1-AT genes. Since PRMT1 is the only known protein arginine methyltransferase responsible for this modification (Strahl et al., 2001; Wang et al., 2001) our results establish that this cofactor is implicated in transcriptional regulation by HNF4. No significant change in PRMT1 levels was detected (Fig. 1B).
To assess if methylation events are critical for gene expression during CaCo-2 cell differentiation, cells were treated with the methyltransferase inhibitors AdOx and MTA for 24 hr prior to harvesting at day 4 (Fig. 1D). While a 4–5 fold stimulation of HNF4 target genes was detected in the untreated cells (lane 1 vs. lane 2) the inhibitor treatment abolished the induction of these genes, suggesting that methylation events are critical for the differentiation of CaCo-2 cells. The treatment showed preferential effects on HNF4 target genes as there was no effect on β-actin or GAPDH levels.
The preceding suggests that PRMT1 is recruited to responsive promoters through HNF4 and implies potential interaction between PRMT1 and HNF4. To look for evidence of such an interaction in vivo, we cotransfected 293T cells with expression plasmids for PRMT1 and HNF4, and immunoprecipitated HNF4 from the derived extracts. Western blots indicated that PRMT1 was co-precipitated with HNF4 (Fig. 2A, lane 5). We also performed GST pull-down experiments using purified proteins (Fig. 2B) and found that PRMT1 showed a strong specific interaction with GST fused to full-length HNF4 (residues 1-455; lane 3 vs. lane 2). To identify the HNF4 domain that interacts with PRMT1 we tested various GST-HNF4 fusion derivatives. Progressive deletions of the HNF4 C-terminus confirmed that the interaction was mediated through the N-terminus of HNF4 (residues 1-116, lane 7). More precise mapping revealed that the interaction relies on the DBD of HNF4 (residues 50-103, lane 16). Furthermore, a GST fusion (residues 1-82, lane 14) lacking the second zinc finger of the DBD was not able to interact with PRMT1, indicating that the second zinc finger plays a crucial role in this interaction.
Since PRMT1 displays a broad spectrum with respect to its methylation substrates (Bedford and Richard, 2005), and given the direct interaction of PRMT1 with the DBD of HNF4, we asked if the latter might be a substrate. First, we addressed whether HNF4 is methylated in vivo. We overexpressed epitope-tagged HNF4 and PRMT1 in 293T cells and determined if they could be metabolically labelled following treatment with S-adenosyl-L-[methyl-3H]-methionine (SAM). In these experiments, PRMT1 served both as a negative control (since it had no HNF4) and as a positive control for the efficacy of the in vivo methylation reaction (since it is known to be automethylated). Fluorography of the affinity-purified proteins (Fig. 2C) revealed that both HNF4 (lane 4) and PRMT1 (lane 3) were methylated in vivo. In addition to PRMT1, other unidentified proteins that copurify specifically with PRMT1 were also methylated (Fig. 2C, lane 3). Furthermore, treatment with MTA and AdOx abolished HNF4 methylation in vivo (Fig. 2D). Despite multiple attempts, we were unable to confirm these results using mass spectroscopic methods, perhaps because of the abundance of arginine and lysine residues in HNF4 DBD and the resulting large number of small tryptic peptides that could not be easily scored.
We confirmed that full-length HNF4 (residues 1-455) is efficiently methylated by PRMT1 in vitro (Fig. 2E, lane 1; compare top and bottom panels). PRMT1 methylated all constructs containing the DBD (including residues 1-116, 1-174, and 50-116; lanes 3, 4 and 5) but failed to methylate a construct containing the LBD (residues 128-380, lane 6) or the first N-terminal 24 residues (lane 2). Finer mapping revealed that residues in the second zinc finger (see Fig. 3E) are needed for methylation, as construct 1-82, which lacked this motif, was not methylated (lane 9). Next we mutated the three arginines located within the second zinc finger. Mutants R100Q (lane 12) or R88Q (lane 14) were efficiently methylated by PRMT1. By contrast, mutant R91Q (lane 13) displayed reduced (by circa 80%) HNF4 methylation efficiency. Similarly, mutant R91W (lane 15) completely abolished methylation within the HNF4 DBD. Although the residual methylation that remains in the R91Q mutant suggests the possible existence of secondary methylation sites, these results clearly establish that the major PRMT1 methylation site within the HNF4 DBD is R91.
To assess the functional consequences of HNF4 methylation we compared the ability of wild type HNF4 and a methylation mutant (R91W) to transactivate a reporter plasmid containing the apoAI enhancer. As shown in Fig. 3A, mutant HNF4 displays a 60% decrease in the transactivation potential relative to the wild-type. To rule out effects on HNF4 subcellular localization we performed immunolocalization experiments in 293T transfected with wild type or mutant (R91W) HNF4 or in HepG2 cells treated with MTA and AdOx (Fig. S1). Neither the mutant nor the treatment changed the localization of HNF4, which remained exclusively nuclear.
To verify if the reduced transactivation by the R91 mutant reflects compromised binding activity we quantified the promoter occupancy by ChIP. Fig. 3B shows that the binding of the R91W mutant to the responsive element of the reporter plasmid is reduced by 25%. However, a quantitative dot blot analysis of the HNF4 in the immunoprecipitated material indicated that the transfected mutant protein accumulates to a circa 4-fold higher level than wild-type (Fig. 3B, inset). Thus, once corrected for this difference, our data reveal that methylation at R91 significantly increases HNF4 binding to cognate sites in vivo.
To further confirm the previous result we purified the overexpressed wild-type and mutant (R91W) HNF4 proteins and tested their ability to bind their cognate element in a gel shift experiment (Fig. 3C). The R91W mutant (lanes 7–10) showed a significant decrease in DNA binding capacity compared to the wild-type (lanes 3–6). To verify that the decrease in binding in the mutant protein is due to absence of eukaryote-specific post-translational modification(s) and not due to gross structural changes, we also checked the DNA binding capacity of corresponding proteins that had been expressed in bacteria in control experiments. These non-modified recombinant proteins bind DNA with similar affinities (compare lanes 12 to 15 and lanes 16 to 19), indicating that the decreased binding observed in the case of the mutant HNF4 expressed in 293T cells is caused by the lack of a postranslational modification.
Next we tested if in vitro methylation of HNF4 by PRMT1 affects its DNA binding. A biotinylated template containing four copies of the apoAI site A was used either directly (Fig. 4B, lanes 1 to 7) or after chromatinization (lanes 8 to 13) to test HNF4 binding affinity in the presence or absence of PRMT1 and SAM. Micrococcal nuclease digestion (Fig. 4A) indicated that our template was able to accommodate two nucleosomes. Under all the conditions tested, HNF4 failed to recruit PRMT1 to the template (Fig. 4B). Accordingly, we could not detect any specific complex between DNA, HNF4 and PRMT1 in gel shift experiments (data not shown). Presence of PRMT1 seemed to impair HNF4 binding to the immobilized probe, probably by steric blockage of the DBD (lane 4 vs. lane 3; lane 10 vs. lane 9). Nonetheless, at least on the chromatinized template, efficient stabilization of HNF4 binding (against interference from PRMT1) was seen in the presence of SAM (lane 13 vs. lane 10). This differential effect may be a reflection of the weaker affinity of HNF4 for chromatin compared to naked DNA. Therefore, methylation of the HNF4 DBD leads to stabilization of binding to chromatin targets.
Increased binding of methylated HNF4 in the experiment of Fig. 4B could be explained through two different mechanisms: (a) methylation of HNF4 could reduce the affinity of PRMT1 for the HNF4 DBD, thus making more HNF4 available for DNA binding; or (b) methylation could enhance HNF4 DNA binding directly. To test the first hypothesis, the binding of GST-HNF4 to PRMT1 (with pre-incubation of the proteins in the absence or presence of SAM) was tested under increasingly stringent conditions. Fig. 4C shows that methylation of HNF4 results in an increase of its affinity for PRMT1. Thus, the increased binding of HNF4 to chromatin following methylation cannot be explained by decreased interaction between HNF4 and PRMT1. The fact that methylation changes the affinity of PRMT1 for its own substrate made us wonder if methylation of histone tails changes the affinity for PRMT1 (Fig. 4D). In contrast to the results obtained for HNF4, methylation of the histone H4 tail was found to result in a reduced affinity for PRMT1. This indicates that methylation can either decrease or increase the affinity for PRMT1 depending on the substrate.
Our results suggest that HNF4 DBD methylation occurs in the absence of DNA binding and that the methylation reaction and DNA binding may be mutually exclusive. We prebound HNF4 to an immobilized DNA template containing four copies of the apoAI site A and performed an in vitro methyltransferase assay using an equivalent amount of free HNF4 as control. Fig. 4E shows that methylation of template-bound HNF4 was dramatically reduced compared to that of free HNF4 proving that DNA binding prevents methylation of the DBD. Thus, methylation of HNF4 can only occur prior to DNA binding.
While our binding data indicate that PRMT1 cannot be directly recruited by HNF4 to a DNA template, ChIP assays indicate that recruitment of this factor is correlated with HNF4 occupancy of responsive promoters and suggest that PRMT1 plays a chromatin coactivator role in HNF4 mediated transcription. Therefore, we designed an in vitro assay system to dissect coactivator function.
Initial pilot experiments allowed us to narrow down the LBD as the major HNF4 transactivation domain. First, transient transfection in CaCo-2 cells (Fig. S2A) using GAL4 fusion derivatives of HNF4 indicated that a fusion containing residues 128-380 (LBD) was 100-fold more active compared to GAL4 DBD alone. By contrast, residues 1-128 displayed no significant transactivation activity. Similarly, in in vitro transactivation assays from a naked DNA template containing five GAL4 sites (Fig. S2B), GAL4-HNF4-LBD was able to stimulate transcription to a level that was comparable to the fold-activation elicited by GAL4-VP16.
We focused then on the possible coactivation of this domain by PRMT1 in an in vitro transcription system based on chromatinized templates reconstituted with recombinant histones (Fig. 5A and 5B). The fact that several reports (Chen et al., 2000; Koh et al., 2001; Lee et al., 2002) suggest synergy between several nuclear receptor coactivators and histone arginine methyltransferases led us to test the ability of PRMT1 to function as an HNF4 coactivator alone or in combination with purified p300 and SRC-1 (Fig. 5C).
To validate our chromatin transcription system we first compared the ability of p300, a well-established coactivator both for HNF4 (Malik et al., 2002) and VP16 (Kundu et al., 2000), to stimulate GAL4-VP16- or GAL4-HNF4 LBD-dependent transcription in the presence of acetyl-CoA (Fig. 5D). In contrast with what is observed with naked DNA, the GAL4-HNF4-LBD alone barely stimulated transcription (lane 4 vs. lane 1). However, further addition of p300 increased transcription by 8-fold (lane 5 vs. lane 4). Similarly, p300 was able to stimulate GAL4-VP16 transcription by about 6.7-fold. The p300 coactivator activity was dependent on the presence of an activator as there was no effect in the absence of GAL4-VP16 or GAL4-HNF4-LBD (lane 6).
In further analysis, we found that similar to p300 (Fig. 6A, lane 5 vs. lane 2), SRC-1 moderately stimulated HNF4-dependent transcription when added alone (lane 3 vs. lane 2). By contrast, PRMT1 had only a weak effect on transcription on its own (lane 4). The simultaneous presence of p300 and SRC-1 led only to a level of stimulation that reflected the sum of the independent activities of these coactivators (lane 7 vs. lane 3 and lane 5). PRMT1 did not show cooperative effects with either SRC-1 (lane 6) or p300 (lane 8). However, a strong synergistic effect that was at least 30-fold over that seen with PRMT1 alone (lane 4) was evident when the three coactivators were added together (lane 9). Note that the total coactivator-dependent induction exceeds 30-fold although this could not be precisely quantitated given the essentially undetectable baseline signal (lane 2). Again, this effect was strictly dependent on the presence of GAL4-HNF4-LBD, as the three coactivators failed to stimulate transcription in the absence of the activator (lane 10).
In control experiments with naked DNA template, in contrast to the results with the chromatin template, addition of p300 and SRC-1 inhibited HNF4-mediated transcription by 40% (Fig 6B, lane 1 vs. lane 2); further addition of PRMT1 reduced transcription to almost undetectable levels (lane 3) suggesting that the coactivation capacity of these factors depends on the presence of chromatin templates.
To demonstrate that PRMT1 coactivates HNF4 through methylation of histone H4 R3 we also reconstituted chromatin templates that contained recombinant H4 in which R3 had been mutated to a glutamine (H4 R3Q) in place of the wild-type (Fig. 5B) (An et al., 2004). We compared this template relative to the wild-type for its ability to be activated in a PRMT1-dependent manner in an in vitro transcription assay in which we reduced the amounts of p300 and SRC-1 to better visualize the PRMT1 effect. As a result, HNF4-dependent transcription was only slightly stimulated by p300 and SRC-1 (Fig. 6C lane 1 vs. lane 2). Whereas the specific stimulation by PRMT1 was in the order of 5-fold when wild type H4 was present in the chromatin (lane 3 vs. lane 2), PRMT1 was not able to stimulate transcription from chromatin containing H4 R3Q (lane 7 vs. lane 8). We also tested the effect of omitting SAM from the transcription reactions, which reduced PRMT1 stimulation of transcription by 50% (lane 4). This confirms that the methyltransferase activity of PRMT1 is required for its role as a coactivator. (The residual activity in the absence of ectopic SAM is likely to arise from SAM in the nuclear extract.) These data convincingly prove that H4 R3 is a major target site through which PRMT1 stimulates HNF4 dependent transcription.
To assess the contribution of histone modifications in the above effects we performed HAT assays using our chromatinized template and the purified coactivators (Fig. 6D). Consistent with previous reports (Xu and Li, 2003), SRC-1 acetylation activity was virtually nonexistent (lanes 1 and 2). Significant HAT activity of p300 was observed and, importantly, found to be dependent on GAL4-HNF4-LBD implying that this cofactor is targeted to chromatin in an activator dependent fashion (lane 3 vs. lane 4) and that the LBD of HNF4 suffices for its recruitment to chromatin. Negligible effects on p300 HAT activity were evident upon addition of SRC-1 or PRMT1 (lanes 5 and 6). However, similar experiments to test for histone methylation with pure components did not yield a detectable signal (data not shown). That H4 R3 methylation does in fact take place during the transcription process and is dependent both on the presence of the specified coactivators (Fig. 6E, lane 1 vs. lane 2) and GAL4-HNF4-LBD (lane 2 vs. lane 3) was confirmed by in vitro ChIP using antibodies against dimethylated H4 R3 after in vitro transcription.
Since the HNF4 LBD failed to interact directly with PRMT1 (Fig. 2B) there remains the question of how PRMT1 is recruited to the promoter by HNF4. It is generally believed that the p160 family of coactivators act as docking sites for recruitment of other coactivators to DNA-bound nuclear receptors (Xu and Li, 2003). To evaluate the possibility that HNF4 LBD, SRC-1 and PRMT1 form a ternary complex that may facilitate the recruitment of PRMT1 to HNF4 responsive elements we conducted a GST pull down experiment. First, we confirmed that SRC-1 independently interacts with the LBD of HNF4 and with PRMT1 (Fig 7A). Next we performed GST pull downs to see if there is facilitated recruitment of PRMT1 to the HNF4 LBD by SRC-1. Under the conditions of this experiment, PRMT1 on its own showed a very weak interaction with HNF4 LBD (Fig 7B, lane 4). However, in the presence of SRC-1 (lane 5), significantly greater amounts of PRMT1 were retained indicating that HNF4, SRC-1 and PRMT1 form a ternary complex that may facilitate the recruitment of PRMT1 to the HNF4 LBD.
Together, our data conclusively demonstrate that PRMT1 is an HNF4 coactivator that functions at the level of the chromatin template through methylation of histone H4 R3 and that its recruitment can be facilitated through the formation of a ternary complex between HNF4, SRC-1 and PRMT1.
The main conclusions of this study include: (i) implication of PRMT1 in transcriptional control by HNF4; (ii) demonstration that nuclear receptor DNA binding can be regulated by PRMT1-mediated methylation of the DBD; and (iii) demonstration of functional synergism between SRC-1, PRMT1, and p300 in mediating HNF4-dependent transcription of chromatin templates. Together, these data describe a new regulatory paradigm whereby a methyltransferase controls nuclear receptor function both by directly modulating its binding to DNA and by effecting H4 methylation on the promoters of target genes.
Data presented here show that DNA binding activity of HNF4 can be regulated by methylation of the receptor. While other sites within HNF4 may also be methylated, a key demonstration here is that residue R91, which is located within the D box of the HNF4 DBD and has been implicated in nuclear receptor dimerization, plays a determinative role in HNF4 function. Although the exact physical basis for this effect is presently unclear, it is known that methylation can alter protein-protein interactions (Bedford and Richard, 2005). Thus, R91 methylation could be critical for HNF4 homodimerization and potentiation of DNA binding.
Our findings, together with other reports, reveal that the HNF4 DBD is a hot spot for post-translation modifications (see Fig. 3E), including phosphorylation (Jiang et al., 1997; Viollet et al., 1997) and acetylation (Soutoglou et al., 2000), that modulate its DNA binding. Given the uncertainty regarding the existence of a bona fide HNF4 ligand, the regulation of HNF4 function through modulation of DNA binding is intriguing. Note that in our in vitro transcription system, HNF4 is able to transactivate in the absence of any added ligand, although bacterially expressed HNF4 LBD contains a mixture of fatty acids that might activate HNF4 constitutively (Dhe-Paganon et al., 2002; Wisely et al., 2002). Therefore, HNF4 might represent a less-evolved form of nuclear receptor, with a potential to be regulated by ligands, yet regulated mainly at the DNA affinity level. As a target of diverse modifications, the HNF4 DBD could act as an integrator of different cellular signals, such as metabolic status (Viollet et al., 1997) or cell differentiation, that would allow the fine regulation of DNA binding. On the other hand, the proximity of the acetylated and methylated residues and the fact that these are modified by coactivators, suggest a possible interplay between the various modifications, analogous to what has been described for histones tails (Jenuwein and Allis, 2001). How general this observation is for other receptors remains to be established but it raises the possibility of the existence of a parallel nuclear receptor code.
Beyond the above-described effects of PRMT1 on HNF4 methylation and DNA binding, our ChIP and in vitro transcription analyses firmly establish that PRMT1-mediated methylation of H4 R3 on HNF4 target genes plays a role in the HNF4-mediated transcription. Our data also demonstrate that this arm of the PRMT1 axis is manifested through the HNF4 LBD and SRC-1. Together, these factors generate a ternary complex, possibly as part of a much larger HNF4-nucleated PIC, which provides the platform from where the histone modification then takes place.
The specific roles for histone methylation in regulation of chromatin structure and transcription remain to be determined since methylation of histones per se does not alter their charge (Stallcup, 2001). Conceivably, methylated arginines in histones may inhibit the binding of repressive proteins or provide binding sites for proteins that contribute to transcriptional activation (Daniel et al., 2005; Jenuwein and Allis, 2001). Cooperation between acetylation and methylation (An et al., 2004; Wang et al., 2001) and a preferred order of function that consists of PRMT1 function first and of p300 second (An et al., 2004; Huang et al., 2005) has also been suggested. In agreement with this, we find that after methylation of H4, PRMT1 loses affinity for the methylated histone tail. This transient interaction with the substrate could, in turn, facilitate the interaction of this histone tail with other coactivators like p300.
Indeed, in our in vitro transcription system, PRMT1 function clearly depends on the simultaneous presence of SRC-1 and p300, suggesting a close functional linkage between acetylation and methylation of histones. Yet, since we used nuclear extract in our in vitro system as a source of transcription factors, we cannot rule out the possibility that other proteins play a role in the observed synergy. For example, facilitated recruitment and function of p300 by SRC-1 has been suggested as a mechanism of p300 and SRC-1 synergy (Liu et al., 2001) but we found that the LBD of HNF4 suffices to recruit p300 to chromatin and efficiently acetylate histones, and that SRC-1 has marginal effects on acetylation by p300. A possible explanation is that the synergy between p300 and SRC-1 observed in in vitro transcription may not be caused by these factors alone but through the participation of other coactivators likely recruited by SRC-1. In the same way, it should be noted that whereas H4 R3 methylation takes place during the transcription process and is critical for PRMT1 HNF4-dependent activation, we were unable to detect efficient methylation of H4 in chromatin methylation assays reconstituted from pure components (data not shown). It is therefore likely that additional factors in the extract are responsible for conferring the methylation requirement, at least in the case of HNF4. Our future work will utilize a transcription system that has been reconstituted from homogenous preparations of Pol II, GTFs, and various coactivators (Malik and Roeder, 2003) to address these additional factor dependencies.
Our data disclose two different but interdependent PRMT1 coactivator mechanisms that may well be operating sequentially (Fig.7C). In the model that emerges, PRMT1 first binds directly to the DBD of unengaged HNF4 and methylates R91, increasing its DNA binding capacity. DNA-bound HNF4 recruits PRMT1 through the LBD, via a mechanism that involves SRC-1, resulting in methylation of histone H4 R3 at target promoters. Function of other coactivators like p300 that acetylate histones may thereby be facilitated, making the promoter more accessible to Mediator, RNA polymerase II, and GTFs ultimately leading to the transcription of HNF4 target genes. Thus by virtue of its participation in two of the key steps in the HNF4 activation pathway, PRMT1 may be a device for coupling global regulation of HNF4 function to its gene-specific effects.
Mammalian expression plasmid for PRMT1 has been previously described (An et al., 2004). GAL4-HNF4 fusion proteins for mammalian expression were generated by amplifying HNF4 residues 1 to 128 (GAL4-HNF4 [1-128]) or residues 128 to 380 (GAL4-HNF4 [128-380]) and cloning in pBIND (Promega).
Expression of His-tagged PRMT1 in bacteria was previously described (An et al., 2004). FLAG-tagged p300 and SRC-1 for in vitro transcription were expressed in Sf9 cells from baculovirus vectors. GAL4-HNF4-LBD for bacterial expression containing the GAL4 DBD (residues 1-94) fused to HNF4 LBD (residues 128-380) was generated using overlap extension and cloned into 6His-pET11d. Recombinant protein was purified over Ni-NTA agarose.
The HNF4 GST derivatives GST-HNF4 (1-455), GST-HNF4 (1-389), GST-HNF4 (1-345) were previously described (Malik et al., 2002). Additional HNF4 GST derivatives were generated by PCR amplification from GST-HNF4 (1-455) and cloned into pGEX4T2 (Pharmacia). Mutations in the DBD were introduced by overlap extension. For expression of GST-H4 the H4 tail (residues 1-36) was cloned into the vector pGEX4T2. Recombinant proteins were expressed and purified using glutathione-Sepharose.
The HNF4 responsive luciferase reporter plasmids ABC.Luc and Ax2.Luc were constructed by cloning the apoAI enhancer (Malik, 2003) spanning sites A, B and C (−250/−110) or two copies of the apoAI site A upstream of the apoAI core promoter (−41/+7), respectively, in the vector pGL3 Basic (Promega).
Expression plasmid for FLAG-tagged HNF4 was constructed by amplification of the full-length HNF4 cDNA (Sladek et al., 1990) and subcloning into the vector pFH-IRESneo (Malik and Roeder, 2003). Mutant HNF4 R91W was generated by PCR and similarly subcloned.
Following amplification from the FLAG-HNF4 R91W, the HNF4 R91W mutant was subcloned into 6His-pET11d. His-tagged HNF4 and HNF4 R91W were expressed in bacteria and purified as previously described (Malik and Karathanasis, 1996) with an additional S-sepharose step.
CaCo-2 cells (ATCC) were maintained in DMEM containing 20% fetal bovine serum; medium was changed every other day. For time course experiments, cells were plated to allow confluence to be reached in two days. ChIP assays were performed essentially as previously described (Ma et al., 2003). The recovered DNA was analyzed using primers that covered the apoAI promoter from −323 to −97 and the α1-AT promoter from −338 to +14. In transiently transfected 293T cells the promoter region of the Ax2.Luc plasmid was amplified by PCR using α-32P-dCTP. Resolved products were quantified using a phosphorimager.
CaCo-2 cells were transfected using Lipofectamine 2000 (Invitrogen); 293T cells were transfected using FuGene 6 (Roche). For normalization, a renilla expressing plasmid was included. Luciferase activity was measured after 24 hr with the Promega Dual-Luciferase Reporter Assay kit.
Bacterial vectors for core histone expression and histone purification were as described (Loyola et al., 2004). Mutant histone H4 R3Q was a gift from Jaehoon Kim (An et al., 2004). Expression and purification of recombinant ACF and NAP-1 and procedures for chromatin assembly and microccocal nuclease analysis were as described (An and Roeder, 2004). Plasmid pG5ML array has been described elsewhere (An and Roeder, 2004). In vitro transcription from chromatin templates was performed as described (An and Roeder, 2004) with the following modifications: 50 ng of chromatinized template was preincubated with the indicated transcriptional activators for 30 min followed by addition of the indicated coactivators. After 20 min HeLa nuclear extract was added to allow PIC formation for 15 min. Finally, labelled nucleotides were added and the reactions were allowed to proceed for 45 min. Products were visualized by autoradiography. In vitro transcription from naked DNA has been described elsewhere (Malik and Karathanasis, 1996).
Cells (293T) were transfected with FLAG-tagged HNF4 and PRMT1 expression plasmids and whole cell extracts were immunoprecipitated with anti-HNF4 antibodies (Santa Cruz). Immunocomplexes were washed with BC buffer (Malik and Roeder, 2003) containing 150 mM KCl and 0.1% NP40 and analyzed by western blot using anti-FLAG M2 antibodies (Sigma) or anti-PRMT1 antibodies (Abcam).
GST pull-down assays were performed and washed with BC buffer containing the indicated salt concentration and 0.1 % NP40. Beads were boiled and analyzed by western blot using anti-PRMT1 or anti-FLAG M2 antibodies.
Cells (293T) were transfected with FLAG-tagged HNF4 or PRMT1 expression plasmids and incubated with [3H]SAM (Amersham) in methionine-depleted DMEM (Mediatech) with dialyzed fetal serum (HyClone) at 37ºC for 4 hr. When specified, cells where preincubated with 1 mM each of MTA and AdOx for 1 hr. Whole cell extracts were used for immunoprecipitation with anti-FLAG M2 agarose. Immunocomplexes were resolved by SDS-PAGE, stained with Coomassie and analyzed by fluorography.
A 450 bp biotinylated PCR product containing four HNF4 cognate sites (apoAI site A) and the ML core promoter element was amplified from plasmid pAx4MLΔ53 (Malik and Karathanasis, 1996). It was reconstituted into chromatin using purified recombinant histones as described above. Chromatin and naked DNA were adsorbed to Dynabeads (Dynal) as previously described (Vermeulen and Stunnenberg, 2004). Beads containing 400 ng of DNA or chromatin were incubated with 200 ng of His-tagged HNF4, 1 μg of His-tagged PRMT1, 50 μM SAM in 100 μl of binding buffer (70 mM KCl, 10 mM Hepes-KOH pH 7.8, 5% glycerol, 2 mM MgCl2, 5 mM DTT, 0.25 mg/ml BSA and 0.5 mM PMSF), washed with binding buffer containing 100 mM KCl, boiled in SDS sample buffer, resolved by SDS-PAGE and analyzed by western blot using anti-His antibodies.
Reactions and electrophoresis conditions for apoAI site A interactions were essentially as reported previously (Malik and Karathanasis, 1996).
We are indebted to Dr. R.G. Roeder for support, encouragement and critical reading of the manuscript. We acknowledge T. Jones and A. Nock for technical help. We especially thank J.H. Kim for valuable reagents and V. Palhan, M. Jishage and other members of the Roeder lab for helpful discussions. This work was supported by NIH grant DK060764 to S.M. and in part through The Rockefeller University’s Women & Science Fellowship to M.J.B.