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Molecular Neuroendocrinology, Max Planck Institute of Psychiatry, Kraepelinstrasse 2-10, D-80804 Munich, Germany
Received 28 November 2005/ Returned for modification 6 January 2006/ Accepted 27 April 2006
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ABSTRACT |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Evidence that Zac expression during embryogenesis in mesenchymal and neural stem/progenitor cells is tightly regulated in a spatiotemporal fashion and evidence that Zac is upregulated following seizures and transient focal cerebral ischemia in mice suggest that the protein may have additional roles, e.g., in neural differentiation and plasticity (2, 15, 48-50). Indeed, Zac was recently recloned in a subtractive hybridization screen for genes involved in neuronal cell fate specification (31). Moreover, Zac expression can be influenced by hormonal and epigenetic signals during regeneration, differentiation, and age-related degenerative processes, although its exact role in these conditions requires further studies (9, 13, 46, 57).
In adults, Zac is highly expressed in most steroid-responsive tissues (35, 36, 48, 52). Interestingly, Zac potently coactivates or corepresses the hormone-dependent activity of nuclear receptors, including that of androgen, estrogen, glucocorticoid, and thyroid hormone receptors (20); all of these receptors are key regulators of cell growth, differentiation, homeostasis, and development and act in a cell-specific manner. Furthermore, recent studies disclosed the importance of Zac's imprinting status in the etiology of transient neonatal diabetes mellitus, an uncommon form of childhood diabetes (NCBI entry OMIM *601410 [http://www.ncbi.nlm.nih.gov/entrez/dispomim.cgi?id=601410]) (28, 51).
Our earlier studies revealed that Zac can act as a transcription factor through its monomeric or dimeric binding either to a GC-rich palindromic DNA element or to GC-rich direct and reverse repeat elements, respectively (6, 18, 52). The two closely related members of the Zac family Plag1 and Plagl2 share at their N termini virtually identical DNA-binding domains consisting of canonical zinc fingers of the C2H2 type (52). In line with this observation, Zac and Plag target genes identified so far (18, 54, 58) contain closely related GC-rich sequences in their promoter regions. Zac family members show, however, marked differences in their expression patterns in several tissues, indicating different biological roles. Moreover, proteins of the Zac family strongly diverge in their C termini, which could determine specific interactions in transcriptional regulation. In fact, only Zac has been reisolated in a yeast screen for mammalian proteins that bind to the C-terminal activation domain of the nuclear receptor coactivator SRC-1 (GRIP1) (20). Moreover, Zac binds to the C-terminal domain of the homologous coactivators p300 and CREB-binding protein (CBP) and strongly coactivates or corepresses nuclear receptors and p53 in a context-dependent manner (19, 20, 39).
The transcriptional coactivator proteins p300/CBP exert key roles in cellular differentiation, growth control, and homeostasis (16). In response to diverse physiological cues, these proteins coordinate and integrate multiple signal-dependent events at the transcriptional level by virtue of their histone acetyltransferase (HAT) activity. This allows them to influence chromatin activity by modulating nucleosomal histones, modify transcription factors, and influence DNA recognition, protein-protein interactions, and protein stability (25). The HAT activity of these coactivators is markedly increased upon their interaction with specific transcription factors (24, 43), although the underlying mechanisms of this switch in transcriptional activity are poorly understood. Whereas the classic recruitment model proposes that coactivation simply reflects the capacity of the activator to recruit the coactivator, we demonstrate here that HAT activity is controlled by the coordinated binding of Zac to p300. Furthermore, our experiments reveal a new function of C2H2 zinc fingers in the regulation of HAT activity, suggesting a dynamic role of DNA-binding proteins in the enzymatic control of transcription.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Zac constructs and reporter plasmids were described previously (18). Coactivator cDNAs used were pCMV-p300-HA (11), pCMV-p300(DY-HAT)-myc (21), pcDNA3-CBP, pcDNA3-CBP(HAT–)L(1690)K, C(1691)L (42), pCX.Flag-PCAF (7), and pCR3.1.SRC1a (34). Single or combined deletion mutants of the p300 KIX (aa 566 to 650) and CH3 (aa 1673 to 1966) domains were generated by quick-change site-directed mutagenesis (Stratagene). p300 (10), CBP (29), or control siRNA oligonucleotides were purchased from MWG, Germany. Luciferase values were standardized by cotransfection of a ß-galactosidase expression vector (0.1 µg of pRK7ß-Gal). Results represent means and standard deviations from at least two experiments done in triplicate.
GST pulldown assays and in vitro translations.
The CH1 (amino acids [aa] 302 to 528), KIX (aa 566 to 829), HAT (aa 1197 to 1673), and CH3 (aa 1673 to 1966) domains were amplified by PCR, cloned into the BamHI site of pGEX-2TK (Amersham Biosciences) and pRK7Flag, and the sequence was verified. Glutathione S-transferase (GST) fusion proteins of p300 and Zac were grown in DH5
, purified on glutathione Sepharose 4B (product code 17-0756-01; Amersham Biosciences), and checked by Coomassie blue staining. Detailed information on the generation of p300 constructs used for in vitro translation is available upon request. In vitro translations were done with the TNT kit (Promega). The figures show representative autoradiograms and results are means and standard deviations from at least three independent experiments. Signals were quantified in a liquid scintillation counter (Beckman LS 6000IC).
For in vitro competition experiments, hemagglutinin (HA)-tagged Zac zinc fingers (aa 1 to 208), C terminus (aa 382 to 667), or luciferase was in vitro translated in the presence or absence of [35S]methionine. Relative concentrations were determined by use of anti-HA antibody. Results represent means and standard deviations from four independent experiments.
Immunoprecipitations and immunoblots. Monoclonal anti-Flag (product code F3165; Sigma), anti-HA (product code 12CA5; Roche), anti-p300 (sc-584), anti-CBP (sc-369), anti-SRC (sc-8995), and antiactin (sc-7210) antibodies (all from Santa Cruz), and anti-Flag M2 affinity gel (product code A-2220; Sigma) were used according to the manufacturers' recommendations. Cells were lysed in 1 ml of lysis buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100). Extracts were incubated for 1 h on ice and then centrifuged for 5 min at 18,000 relative centrifugal force. Following a preclearing step, soluble extracts were incubated at 4°C on a rocking platform with either anti-Flag M2-agarose affinity gel (A-2220; Sigma) or anti-p300 (sc-584). Immunoprecipitates were washed three times with 1 x TBS (50 mM Tris-HCl, pH 7.5, 150 mM NaCl), eluted with sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis sample buffer and immunoblotted.
Immunoblots were performed on total cell lysates (5 to 50 µg protein) and peroxidase-linked anti-rabbit or anti-mouse immunoglobulin G (product code A-0545 or A-6782; Sigma).
A p300 cDNA fragment encoding part of the N-terminal region (aa 1 to 347) or part of the C-terminal region (aa 2042 to 2157) was subcloned into pGEX-2TK (Amersham Biosciences). Rabbits were immunized four times with 100 µg of the fusion protein at 10-day intervals, and antisera were collected after 75 days. The specificity of the antibody (1:500 to 1:1,000) was tested by immunoblotting of mock- or p300-transfected 293T cells. Cellular lysates (50 µg) were blotted and probed with preimmune sera, antibody anti-p300-N (1:1,000) or anti-p300-C (1:500), or p300 antibodies preabsorbed on GST-, GST-p300-N-glutathione-Sepharose, or GST-p300-C-glutathione-Sepharose, respectively. Signals were specific for p300-transfected cells. No signals were detected with either the preimmune or preabsorbed p300 sera.
Electrophoretic mobility shift assays.
Nuclear extracts were checked with anti-Flag, anti-HA, and anti-p300-N antibodies for equivalent levels of Zac or Zac
C1 expression relative to p300 and derivatives. Then, 10 µg of nuclear cell extract was used for binding reactions as described previously (18).
In vitro HAT assays. Acetylation reactions were performed essentially as described previously (27). GST-p300-HAT (aa 1197 to 1673) (25 nM) was incubated at 30°C for 30 min with GST proteins (GST-Zac, GST-Zac-ZF, GST-TFIIE, or GST-TFIIF, 1 µM each) as substrates in the presence of 3.5 µl [3H]acetyl coenzyme A ([3H]acetyl-CoA) in 1 x HAT buffer (50 mM Tris-HCl, pH 8.0, 10% glycerol, 1 mM EDTA, 10 mM phenylmethylsulfonyl fluoride) in a total volume of 40 µl. Reactions were fractionated on a 10% SDS-polyacrylamide gel. Gels were treated with 2,5-diphenyloxazol (56) and exposed to Hyperfilm MP (product code RPN6K; Amersham Biosciences) for 1 to 3 days. The experiment was done three times, and all experiments yielded similar results.
In vitro-translated p300 proteins (p300, p300KIX, and p300CH3) were preincubated with an equimolar dose of GST-Zac proteins (GST-Zac, GST-ZacC1, and GST-ZacZF6, 1 nM each) for 10 min at room temperature in 1 x HAT buffer in a total volume of 45 µl. Different amounts of GST fusion proteins were used for Fig. 10C. The reactions for kinetic analysis and for HAT progression curves were initiated by the addition of 300 µM biotinylated N-terminal H4 (aa 2 to 24) peptide (product code 12-372; Upstate Biotechnology) and 0.75 µCi [3H]acetyl-CoA (5.3 Ci mmol–1) (product code TRK688; Amersham Biosciences) in a volume of 5 µl, and the mixtures were incubated for 1 to 15 min at 30°C. Saturating (300 µM) or differing concentrations of core histones (product code H9250; Sigma) or H4 peptide were assayed in parallel for Fig. 10A and B. Saturation analysis of H4 peptide and acetyl-CoA binding reactions were initiated likewise, and the mixtures were incubated for 2 min at 30°C. After the addition of 500 µl 1 x HAT buffer with 30 µl of 50% slurry of streptavidine agarose (product code 69203-3; Novagen), the samples were incubated for 20 min at 4°C on a rotating wheel. The beads were washed twice with radioimmunprecipitation assay buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 1 mM EDTA, 0.1% phenylmethylsulfonyl fluoride) and counted in a liquid scintillation counter. All reactions were conducted in the presence of saturating amounts of the direct repeat (DR) element.
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ChIP assay. Chromatin immunoprecipitation (ChIP) assays were performed essentially as described previously (43). Primers used for PCR were from the direct repeat reporter (18) flanking the Zac response elements (5' primer, 5'-TATCTTATGGTACTGTAACTGAGC-3'; 3' primer, 5'-CTAGAGGATAGAATGGCGCCGGGC-3'). Antibodies against acetyl-histone H4 were from Upstate Biotechnology (product code 06-598). Levels of expression of HA-tagged Zac, Flag-tagged p300, and their derivatives were monitored by immunoblotting with anti-HA and anti-Flag antibodies. In all cases, aliquots of PCR carried out for 25 or 35 cycles were analyzed to ensure that amplification was maintained in the linear range.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Given this interaction in vitro and in vivo, we investigated its relevance for Zac transactivation. Apoptotic cell death is known to cause global nuclear condensation, probably leading to broad transcriptional repression (37), confounding coactivator or transcriptional assays, and to potentially target p300/CBP for degradation (38). Therefore, we performed reporter assays with LLC-PK1 cells, which fairly resist Zac-induced cell death at 24 h following transfection (44, 52). We cotransfected Zac together with a reporter plasmid harboring two DR elements or four palindromic elements (PAL) in the absence or presence of additional amounts of p300 or CBP. Coexpression of wild-type p300 or of a HAT-defective version failed to enhance basal activities of the DR or PAL reporters, which were set to 100% (Fig. 1 E and F). In contrast, Zac-induced DR or PAL reporter activities set to 100% (Fig. 1 E and F) were about twofold induced by coexpression of p300 or CBP, while the HAT-deficient versions were completely ineffective in this respect. Similarly, coexpression of p300 but not of its HAT-deficient version specifically increased Zac-dependent transactivation of the CK14 promoter (Fig. 1G), a previously identified Zac target gene (18). In contrast, coexpression of broadly differing doses of the unrelated coactivators PCAF and SRC-1 (GRIP1) failed to stimulate Zac-dependent transactivation of the DR and PAL reporter plasmids (Fig. 1E and F and data not shown) although both PCAF (data not shown) and SRC-1 readily bound to Zac (20; data not shown). To rule out the possibility that these discrepancies simply resulted from differences in Zac expression in the presence of individual coactivators, we carried out immunoblotting. As shown in Fig. 1H, cotransfection of p300 or p300-HATmt did not influence Zac protein levels and vice versa. This was also the case for CBP, CBP-HATmut, PCAF, and SRC-1. Together, these data show that solely p300 and CBP support Zac transactivation in a strictly HAT-dependent manner.
In a complementary approach to these experiments, we depleted HeLa cells of endogenous p300 or CBP by RNA interference. HeLa cells were chosen at this step because they could be efficiently transfected by electroporation (>70% ß-galactosidase-positive cells following transfection of 100 ng of pRK7-ß-galactorisdase expression vector). p300 or control siRNA oligonucleotides (Fig. 2A, columns 2 and 3) failed to affect basal DR reporter activity, which was set to 100% (column 1). In contrast, Zac-dependent reporter activity (similarly set to 100% [Fig. 2A, column 4]) was diminished by p300 siRNA at the highest dose by fivefold (column 7), whereas control siRNA oligonucleotides of the same dose were largely ineffective (column 8). Immunoblotting evidenced that p300 siRNA reduced p300 protein in a dose-dependent manner, which correlated well with reductions in Zac-dependent transactivation (Fig. 2A, bottom panel). Additionally, to exclude the possibility that p300 or control siRNA affected Zac DNA binding, we carried out electrophoretic mobility shift assays (EMSA). These experiments displayed no differences between samples from either treatment or control conditions, attesting to the specificity of the results (data not shown). Similar to how we tested p300, we further tested the role of endogenous CBP to enhance Zac transactivation. CBP siRNA did not reduce basal DR reporter activity and impaired only weakly Zac-dependent activity at the highest dose tested (Fig. 2B). Immunoblotting revealed, however, efficient reduction of CBP protein levels, comparable to the results obtained for p300. In addition, CBP siRNA did not change Zac DNA binding (data not shown). Therefore, the strong decline in Zac transactivation upon p300 knockdown compared to the weak effect of CBP knockdown indicated a preferential interaction of Zac with endogenous p300 to enhance transactivation.
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Mapping Zac and p300 interactions. To delineate the domains of Zac and p300 interacting with each other, we in vitro translated successive segments of p300, schematically depicted at the top of Fig. 3A. Pulldown assays evidenced that the N-terminal KIX domain (aa 566 to 829), the central HAT domain (aa 1197 to 1673), and the C-terminal CH3 domain (aa 1673 to 1966) bound with similar affinities to GST-Zac protein. No binding occurred in the case of the CH1 domain, bromodomain, and steroid receptor interacting domain. To follow up the contribution of the KIX and CH3 domains to overall binding, we deleted them singly or together (Fig. 3B). Zac binding to singly deleted p300 derivatives persisted (columns 2 and 3), whereas combined deletion reduced binding by 60% (column 4). As expected, absence of either Zac binding site abolished p300 binding (column 5).
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ZF, shortened by the N-terminal zinc finger domain; ZacL and ZacPR, with the N-terminal or C-terminal part of the central transactivation domain deleted; and ZacC, shortened by the C terminus. Interestingly, the absence of the zinc fingers strongly reduced binding to the KIX, HAT, or CH3 domains (Fig. 3C, columns 2, 7, and 12). In contrast, the deletion of the central transactivation domains showed no effect (columns 3, 4, 8, 9, 13, and 14), whereas deletion of the C terminus selectively impeded binding to the KIX domain (compare column 5 with columns 10 and 15). Equivalent amounts of GST fusion proteins or of GST alone were used in these experiments (Fig. 3D).
In line with these results, the isolated zinc fingers efficiently bound to the HAT and the CH3 domains (Fig. 3E, columns 6 and 10), whereas the central transactivation domain failed to interact with any of the p300 domains (columns 3, 7, and 11). Moreover, both the zinc fingers and the C terminus alone weakly bound to the KIX domain (columns 2 and 4), suggesting that they cooperated in binding. Irrespective of this, Zac and ZacC indistinguishably bound to p300 (as opposed to the behavior of isolated domains), suggesting that overall binding reflected multipoint interactions (data not shown).
Mouse and human Zac proteins share high homology in the N-terminal part of the C terminus (labeled C1) but not in the central and C-terminal parts (labeled C2 and C3, respectively), which are specific to mice (Fig. 1A and 3C, top) (18, 44, 52). Therefore, we determined the region of mouse Zac responsible for binding to the KIX domain. In fact, the C1 region accounted for the interaction with p300 (Fig. 3F, column 2). This suggested a conserved function to this part of human ZAC, which actually bound with affinity similar to that of the mouse one to the KIX domain (data not shown). Our previous work evidenced this region of human ZAC to contain high transactivation potency as opposed to the complete absence of transactivation in the corresponding part in mice (18, 52). As a result, transactivation potency and p300 binding are strictly separated in the case of the mouse C terminus but coexist in the case of the human one. Conclusively, Zac interacts with the p300-KIX, -HAT, and -CH3 domains through the zinc fingers and the C1 region. The zinc fingers are necessary and sufficient for binding to the HAT and CH3 domains, while binding of the C1 region is restricted to the KIX domain, possibly in cooperation with the zinc fingers.
Zinc fingers interact cooperatively and selectively with p300. To establish more firmly whether the decline in binding of the isolated C terminus and the zinc fingers to the KIX domain stemmed from cooperative binding, we conducted in vitro competition experiments. Results shown in Fig. 3E showed that the isolated zinc fingers and the C terminus bound with similar affinities to the KIX domain. Therefore, we competed binding of the C terminus to limiting concentrations of the KIX domain with increasing amounts of the zinc fingers and vice versa (Fig. 4A). Equimolar amounts of the unlabeled zinc fingers caused a twofold decline in self-competition but weakly decreased binding of the C terminus. We obtained similar results for the reverse experiment, in which equimolar amounts of the unlabeled C terminus weakly reduced binding of the zinc fingers yet led to a twofold decline in self-competition. In further support of these findings, a fivefold excess of the zinc fingers only halved binding of the C terminus and vice versa but generated a sixfold decrease in self-competition. Finally, a 15-fold excess of competitor completely abolished binding either of the zinc fingers or of the C terminus to the KIX domain, suggesting that they shared overlapping sites. In contrast, competition of the zinc fingers or of the C terminus with a 15-fold excess of in vitro-translated luciferase protein failed to reduce significantly their binding to the KIX domain, attesting to the specificity of these results. Together, these data support a cooperative binding of the zinc fingers and of the C terminus to the KIX domain at possibly overlapping sites. On the basis of these experiments, we surmised the following model for the interaction between Zac and p300 (Fig. 4B): (i) the zinc fingers and the C1 region cooperatively bind to the KIX domain, and (ii) the zinc fingers additionally interact with the CH3 domain. In this regard, numerous transcription factors are known to bind to p300/CBP at multiple sites, although the functional implications of such multipoint interactions remain poorly understood (16, 17). Therefore, Zac binding to the KIX and CH3 domains raised the question of whether their binding is functionally coordinated or whether the KIX and CH3 domains operate independently of each other to mediate Zac coactivation.
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Zinc fingers and p300 interact in a DNA-bound complex.
To test whether DNA-bound Zac stably complexes with p300 and, if so, whether ZF6 and ZF7 actually participate in p300 binding, we firstly established EMSA with extracts prepared from PA-TU cells expressing ectopic Zac protein tagged with the Flag epitope. Control experiments confirmed that DNA complex formation was Zac dependent and sequence specific (data not shown). The addition of Flag antibody to the reaction mix supershifted Zac complexes on the DR and PAL elements in a dose-dependent manner (Fig. 5B, lane 2; data not shown). While we detected no significant differences in the affinity of Zac-DNA complexes in the absence or presence of ectopic p300, prolonged electrophoresis disclosed a tiny but consistent decrease in mobility (data not shown). Based on this initial evidence, we raised polyclonal p300 antibodies directed against N- or C-terminal regions of p300 as outlined in the legend to Fig. 5A. Both antibodies failed to affect Zac-DNA complexes in the absence of ectopic p300 (Fig. 5B, lanes 3 and 4). However, following cotransfection of p300, they caused dose-dependent supershifts (lanes 7 and 8), while the respective preimmune sera were ineffective (lanes 9 and 10). EMSA with the PAL element showed similar results under these conditions (data not shown). Supershifted complexes migrated at largely reduced mobility and demonstrated that nearly all of Zac was complexed with p300. These findings begged the question of whether ZF6 and ZF7 in a DNA-bound state participated in p300 binding or whether, alternatively, this function solely reflected the presence of the C1 region. To evaluate the contribution of the C1 region to Zac-p300-DNA complex formation, we compared Zac and ZacC1 in EMSA. Both Zac proteins were strongly supershifted on the DR and PAL elements in the presence of p300 (Fig. 5C; data not shown). Together with results from Fig. 4C, showing that ZF2, ZF6, and ZF7 underlie p300 binding, Zac interaction with p300 despite the absence of the C1 region strongly suggests that DNA-bound ZF6 and ZF7 participate in p300 binding.
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C1 into LLC-PK1 cells at low doses to account for rate-limiting concentrations of endogenous p300. Immunoblots of adjusted amounts of the respective expression vectors demonstrated equal levels of Zac proteins (Fig. 6A). Strikingly, we measured a dramatic difference in transactivation efficiency between these two Zac proteins on the DR and PAL elements (Fig. 6B). These differences were strongest at low doses of Zac (>100-fold) and steadily declined at higher ones, to almost disappear for maximal transactivation. We further cotransfected Zac or ZacC1 into p300-negative PA-TU cells and measured a largely blunted twofold difference in transactivation efficiency (Fig. 6C). Interestingly, cotransfection of a given amount of p300 strongly restored Zac but not ZacC1 activity at low concentrations on the DR and PAL DNA elements (Fig. 6C). Together, these data suggested a role for cooperative binding of the C1 region and of the zinc fingers to the KIX domain to mediate coactivation via p300. If so, high doses of ZacC1 could overcome impaired binding of the zinc fingers to the KIX domain, restoring similar levels of maximal transactivation. Moreover, the almost complete loss in transactivation at low doses of ZacC1 indicates that coordinated binding to the KIX and CH3 domains controls Zac coactivation by p300.
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C1 in growth inhibition of LLC-PK1 cells, while the difference between Zac and ZacC1 in growth inhibition of PA-TU cells was only twofold (Fig. 6D). Thus, coactivation by p300 appears to correlate well with Zac antiproliferation in vivo. KIX and CH3 domains interdependently regulate Zac coactivation. We conducted in vivo competition experiments for p300 binding to infer the function of the KIX and CH3 domains in Zac coactivation (diagrammed in Fig. 7B). We cotransfected Zac with adjusted amounts of the isolated CH1, KIX, and CH3 domains into LLC-PK1 cells (Fig. 7A). Interestingly, increasing amounts of either the KIX or CH3 domain severely impaired Zac transactivation on the DR and PAL elements (Fig. 7C and D). Overexpression of the KIX domain in conjunction with the CH3 domain, which most likely disrupted the interaction between Zac and p300, was only slightly more effective than overexpression of either domain alone. Because overexpression of the CH1 domain hardly affected Zac transactivation, transcriptional squelching appeared rather unlikely. In support of this idea, p53 displayed a differential response (data not shown) with the CH1 and CH3 domains, producing moderate and potent inhibition, respectively (17). Conceptually, competition between ectopically expressed p300 domains and endogenous p300 for Zac binding indicated that coactivation required both the KIX and CH3 domains. Additionally, overexpression of the CH3 domain, which competed solely the interaction of the zinc fingers with the CH3 domain of endogenous p300, pointed to a regulatory role of the zinc fingers in coactivation via p300.
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KIX, p300CH3, and p300KIXCH3 as shown in Fig. 8A.
Subsequent EMSA with the DR element evidenced indistinguishable supershifts between p300 (Fig. 8B, lane 4) and derivatives in which the KIX or CH3 domains were singly deleted (lanes 6 and 8), whereas absence of both domains completely prevented the formation of Zac-p300 complexes (lane 10). Similar results were obtained for the PAL element (data not shown). Reporter assays conducted in PA-TU cells showed that cotransfection of p300KIX failed to enhance Zac transactivation on the DR or PAL elements (Fig. 8C), while p300CH3 even inhibited residual Zac activity (Fig. 8D). As expected, p300KIXCH3 failed to affect transactivation in agreement with the absence of stable Zac-p300 complexes (data not shown). These experiments strengthened the concept that coordinated p300 binding via the KIX and CH3 domains mediates Zac coactivation.
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To exclude any other potential effectors that could assist binding to p300 and modify HAT activity in vivo, we performed in vitro HAT assays with recombinant Zac in the presence of its DNA elements and in vitro-translated p300 protein. Recent reports suggest that small peptides are suitable substrates for kinetic or mechanistic studies of p300-HAT activity (8, 47). Therefore, we compared firstly core histones to a peptide (corresponding to the first 24 amino acids of histone H4) that is coupled to biotin to allow rapid quantification of acetylation products. A kinetic analysis with saturating amounts of core histones or H4 peptide and [3H]acetyl-CoA revealed closely related acetylation reactions with a linear range between 1 and 7 min (Fig. 10A). We additionally tested various concentrations of core histones and H4 peptide, using as the shortest possible period an incubation time of 2 min, and plotted the incorporated radioactivity (3H acetylation) in a double-reciprocal graph. The deduced Km values for core histones [Km(ch), 21 µM] and H4 peptide [Km(h), 25 µM] were strongly related (Fig. 10B), confirming the validity of H4 peptides as substrates for p300-mediated acetylation.
The addition of Zac dose-dependently enhanced HAT activity, reaching a maximum at equimolar ratios of Zac and p300 (Fig. 10C). Higher Zac doses, however, caused a decline, probably due to competition for limited amounts of available p300 binding sites, leading to one-sided contacts blocking p300 activation. In contrast to the biphasic response for increasing Zac concentrations, we observed a steady decline for ZacC1, probably due to binding solely to the CH3 domain. We conducted this experiment also in the presence of ZacZF6, which binds to perfect DR elements (18) and p300 indistinguishably from wild-type Zac (data not shown). However, ZacZF6 did not significantly regulate HAT activity, strengthening the regulatory role of the zinc fingers (Fig. 10C).
Mutational and crystallographic studies suggest that nuclear acetyltransferases share a structurally conserved central core domain, which mediates acetyl-CoA binding and catalysis, whereas the sequence variability within a structurally related framework of the N- and C-terminal domains determines substrate binding specificity. A recent report, however, demonstrated that amino acid residues in the middle of the p300-HAT domain, which is the evolutionarily best-conserved subregion, are intimately involved in both acetyl-CoA and histone binding (8). Given this functional entity, we studied in parallel the influences of Zac on p300 histone and acetyl-CoA binding with differing concentrations of H4 peptide or [3H]acetyl-CoA and an excess of the Zac DR element. We plotted the incorporated radioactivity (3H acetylation) in a double-reciprocal graph to deduce Km values for histone [Km(h)] and acetyl-CoA binding [Km(a)]. We conducted these experiments in the absence or presence of Zac, ZacC1, or ZacZF6. Interestingly, the addition of Zac enhanced histone and acetyl-CoA binding [Km(h), 16 µM; Km(a), 14 µM] compared to p300 alone [Km(h), 25 µM; Km(a), 20 µM] (Fig. 10D). In contrast, ZacC1 reduced [Km(h), 47 µM; Km(a), 26 µM], while ZacZF6 failed to regulate p300-HAT activity [Km(h), 25 µM; Km(a), 21 µM]. Results for Zac and ZacC1 in the presence of the DR element were identical to those measured in the presence of the PAL element (data not shown). Since ZacZF6 fails to bind the PAL element (18), we tested regulation of HAT activity in this case solely in the presence of the DR element. Moreover, Zac but not ZacC1 or ZacZF6 seemed to increase 3H-acetylation rates (see below).
Together, our data indicate (i) that equimolar ratios of Zac and p300 most efficiently induce HAT activity, (ii) that coordinated Zac binding similarly increases p300 histone and acetyl-CoA binding, (iii) that ZF6 and ZF7 exert an important regulatory function, since binding to the CH3 domain alone inverts this enhancement, and (iv) that the zinc fingers are indispensable to HAT regulation.
To validate this concept, we further studied the regulation of p300KIX and p300CH3. Kinetic analysis displayed robust HAT activity similar to p300 (see below). The apparent Km values of p300KIX [Km(h), 19 µM; Km(a), 19 µM] mirrored the ones of p300 [Km(h), 25 µM; Km(a), 20 µM], while they clearly increased by two- and threefold, respectively [Km(h), 51 µM; Km(a), 60 µM] in the case of p300CH3 (Fig. 10E and F; for a survey, see Fig. 11D). This suggested a critical role of the CH3 domain in modulating the affinity of p300 substrate binding. In accord with results given above, binding of Zac solely to the CH3 domain of p300KIX relatively impaired the affinities [Km(h), 26 µM; Km(a), 24 µM], whereas binding to the KIX domain of p300CH3 was largely ineffective. These results evidenced the differential but interdependent functions of the KIX and CH3 domains in Zac-mediated regulation of p300 histone and acetyl-CoA binding.
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C1, or ZacZF6. Linear least-squares fit was performed over the linear portion of data to deduce maximal rates of vz. Interestingly, equimolar amounts of Zac robustly increased vz by 67%, whereas ZacC1 impeded vz by 17% (Fig. 11A). Again, ZacZF6 failed to regulate HAT activity, reinforcing the essential role of the zinc fingers in this process. We additionally studied Zac regulatory effects on p300KIX and p300CH3 (Fig. 11B and C). p300KIX resembled wild-type p300, while the vz of p300CH3 paradoxically increased by 79%. This could indicate a more dynamic role of the CH3 domain in p300-HAT function by its activating substrate binding at low concentrations and by its inhibiting catalytic activity at high ones. Irrespective of this topic, the addition of Zac reduced the vz of p300KIX by 25% and of p300CH3 by 7%. Hence, Zac binding to either the KIX or CH3 domain alone failed to enhance or even reduced p300 acetylation rates. Therefore, in accord with Zac-mediated enhancement of p300 substrate binding, regulation of the catalytic activity strictly required coordinated binding of the C1 region and of the zinc fingers to p300. Coordinated Zac binding regulates in vivo p300-HAT activity. To validate whether Zac regulation of in vitro HAT activity mirrored regulation of p300-HAT activity in vivo, we cotransfected the DR element and HA-tagged Zac or derivatives (18) in the absence or presence of Flag-tagged p300 or derivatives into PA-TU cells. Following cultivation for 24 h, we carried out ChIP experiments with an antibody directed against acetylated histone H4. While H4 acetylation of the reporter (Fig. 12B, lane 2) increased weakly or not at all following cotransfection of either Zac or p300 alone (lanes 5 and 8), we detected strikingly increased H4 acetylation following cotransfection of Zac and p300 together (lane 11). Moreover, p300 recruitment to the reporter plasmid strictly correlated with the presence of Zac (compare lanes 7 and 10).
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C1, and ZacZF6 in the absence or presence of ectopic p300 for their capability to support acetylation. In the absence of p300, they uniformly showed low levels of H4 acetylation (Fig. 12C, lanes 2, 5, and 8). In contrast, Zac but not ZacC1 or ZacZF6 strongly enhanced H4 acetylation in the presence of p300 (Fig. 12C, compare lane 11 with lanes 14 and 17). Importantly, all Zac constructs recruited p300 similarly (lanes 10, 13, and 16), demonstrating that coordinated Zac binding determined regulation of p300-HAT activity. To corroborate further this idea, we investigated H4 acetylation by p300 and its derivatives in the absence or presence of Zac. Neither p300 nor any of its derivatives enhanced H4 acetylation in the absence of Zac (Fig. 12D, lanes 2, 5, and 8). In contrast, solely p300 (lane 11) but neither p300KIX nor p300CH3 (lanes 14 and 17) conferred potent H4 acetylation in the presence of Zac. In sum, these data faithfully reflect the specific mode of Zac-p300 interaction deduced from the preceding sections and closely recapitulate the conclusions from the in vitro HAT assays. |
DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Coordinated Zac binding regulates p300 enzymatic activity. The typical activator operates by binding to specific sites on DNA and contacting with its activating region the multiprotein machinery that directs transcription. According to this model, the specificity and magnitude of activation depend largely on the DNA-binding address of the activator and its capacity to recruit the transcriptional machinery (23). In continuation of this concept, recruitment of p300/CBP is a key event in triggering transcriptional activation through these coactivators. Consequently, the level of transcription and/or HAT activity induced by coactivator-dependent transcriptional factors is postulated to correlate with the strength of the transcription factor-coactivator interaction (22, 40).
In contrast, we show here that coordinated binding of Zac to p300 regulates HAT activity. In support of our concept, dominant negative mutants of the hepatocyte nuclear factor 1 paradoxically exhibit even stronger interaction with either CBP or PCAF than the wild-type protein; however, CBP and PCAF recruited by these mutants lack HAT activity (43). This suggests that the specific mode of interaction between a transcription factor and a coactivator regulates HAT activity, possibly by allosteric mechanisms. In the present study, we developed a comprehensive model for Zac binding to p300 by assigning possible interactions from a functional perspective based on complementary in vitro and in vivo experiments. Our model proposes that the Zac C1 region and the zinc fingers function as possible allosteric effectors of HAT activity by coordinated binding to the KIX and CH3 domains. Mechanistically, Zac increases the affinity for histone and acetyl-CoA binding and the catalytic activity. Since Zac binding to either the KIX or the CH3 domain alone confers no or even negative regulation, respectively, the KIX and CH3 domains must operate in a differential but interdependent all-or-nothing manner to mediate Zac coactivation. Our results represent the first demonstration of how a transcription factor can coordinately modulate various enzymatic properties of a HAT. Further, they raise the possibility that allosteric regulation of enzyme activity might extend to transcription factor-coactivator interactions, leading us to speculate that this type of regulation may be of general importance, potentially opening new opportunities for more selective targeting of aberrant HAT activity in disease.
Multitasking C2H2 zinc fingers link Zac DNA binding to the regulation of HAT activity. While several proteins, including Zac, contain multiple C2H2 fingers which typically comprise the DNA recognition motif, other classes of zinc fingers act exclusively in protein-only or protein-lipid interactions. Among the latter is the RING family of zinc fingers that is involved in ubiquitination processes, phosphate inositol signaling (FYVE and PHD domains), and the assembly of large protein complexes (LIM, TAZ, and PHD domains) (32). A remarkably small number of DNA-bound C2H2 zinc fingers interacts with other proteins (30); for those that do, recent reports emphasized the importance of posttranslational modifications including acetylation (33, 59).
Zinc finger domains are common, relatively small protein motifs that fold around one or more zinc ions. Classic C2H2 zinc fingers conform to the well-characterized ßß-fold, comprising a short ß-hairpin followed by a 10- to 11-residue -helix. Residues near the N-terminal half of the -helix are primarily responsible for contacts to nucleotides, binding in an antiparallel manner to DNA, with the N terminus directed into the major groove and the C terminus directed towards the surrounding environment. Therefore, DNA recognition and p300 binding most likely localize to separate surfaces of Zac ZF6 and ZF7 and illustrate that domains as small as 30 to 40 residues may represent two (or perhaps more) separable functional surfaces in transcriptional regulation. If so, what is the advantage of such a functionally linked entity? Most likely, coupling DNA binding to regulation of HAT activity would provide a potent link to transcription. While a large group of transcription factors may conform to the classic recruitment model, in which HAT activity is rather passively tethered, a subgroup, as exemplified by Zac, may regulate HAT activity more dynamically. Kinetic profiles of p300 occupancy in vivo predict that sustained association marks genes containing CpG islands, where local methylation and histone acetylation exert significant regulatory influence (41). Zac DNA-binding sites are GC rich, a feature typical of CpG islands (3). A restraint chromatin structure at Zac- or other DNA-binding sites may require potent HAT activity to promote transcription. Moreover, biochemical studies implicate the amino-terminal tails of histones both in folding of nucleosomes into higher order chromatin (12) and in directly controlling transcription factor binding to nucleosomal DNA (26, 53). Histone acetylation stabilizes the binding of transacting factors and might occur either before or concurrently with the interactions of transcription factors with their binding sites in chromatin (26, 53). From this perspective, the link between Zac DNA binding to HAT regulation would provide additional stability to the binding of Zac to nucleosomal DNA.
Based on the results presented here, we propose that this new mechanism, linking DNA binding to enzymatic regulation of transcription, may also apply to other activators which comprise C2H2 or other DNA recognition motifs and could be of general importance.
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ACKNOWLEDGMENTS |
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D.S. was supported by the DFG.
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FOOTNOTES |
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REFERENCES |
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2. Alam, S., D. Zinyk, L. Ma, and C. Schuurmans. 2005. Members of the Plag gene family are expressed in complementary and overlapping regions in the developing murine nervous system. Dev. Dyn. 234:772-782.[CrossRef][Medline]
3. Arima, T., T. Kamikihara, T. Hayashida, K. Kato, T. Inoue, Y. Shirayoshi, M. Oshimura, H. Soejima, T. Mukai, and N. Wake. 2005. Zac, Lit1 (Kcnq1Ot1) and P57(Kip2) (Cdkn1C) are in an imprinted gene network that may play a role in Beckwith-Wiedemann syndrome. Nucleic Acids Res. 33:2650-2660.
4. Basyuk, E., V. Coulon, D. A. Le, M. Coisy-Quivy, J. P. Moles, A. Gandarillas, and L. Journot. 2005. The candidate tumor suppressor gene ZAC is involved in keratinocyte differentiation and its expression is lost in basal cell carcinomas. Mol. Cancer Res. 3:483-492.
5. Bilanges, B., A. Varrault, E. Basyuk, C. Rodriguez, A. Mazumdar, C. Pantaloni, J. Bockaert, C. Theillet, D. Spengler, and L. Journot. 1999. Loss of expression of the candidate tumor suppressor gene ZAC in breast cancer cell lines and primary tumors. Oncogene 18:3979-3988.[CrossRef][Medline]
6. Bilanges, B., A. Varrault, A. Mazumdar, C. Pantaloni, A. Hoffmann, J. Bockaert, D. Spengler, and L. Journot. 2001. Alternative splicing of the imprinted candidate tumor suppressor gene ZAC regulates its antiproliferative and DNA binding activities. Oncogene 20:1246-1253.[CrossRef][Medline]
7. Blanco, J. C., S. Minucci, J. Lu, X. J. Yang, K. K. Walker, H. Chen, R. M. Evans, Y. Nakatani, and K. Ozato. 1998. The histone acetylase PCAF is a nuclear receptor coactivator. Genes Dev. 12:1638-1651.
8. Bordoli, L., S. Husser, U. Luthi, M. Netsch, H. Osmani, and R. Eckner. 2001. Functional analysis of the p300 acetyltransferase domain: the PHD finger of p300 but not of CBP is dispensable for enzymatic activity. Nucleic Acids Res. 29:4462-4471.
9. Ciani, E., A. Hoffmann, P. Schmidt, L. Journot, and D. Spengler. 1999. Induction of the PAC1-R (PACAP-type I receptor) gene by p53 and Zac. Mol. Brain Res. 69:290-294.[Medline]
10. Debes, J. D., L. J. Schmidt, H. Huang, and D. J. Tindall. 2002. p300 mediates androgen-independent transactivation of the androgen receptor by interleukin 6. Cancer Res. 62:5632-5636.
11. Eckner, R., M. E. Ewen, D. Newsome, M. Gerdes, J. A. DeCaprio, J. B. Lawrence, and D. M. Livingston. 1994. Molecular cloning and functional analysis of the adenovirus E1A-associated 300-kD protein (p300) reveals a protein with properties of a transcriptional adaptor. Genes Dev. 8:869-884.
12. Felsenfeld, G., and M. Groudine. 2003. Controlling the double helix. Nature 421:448-453.[CrossRef][Medline]
13. Fulco, M., R. L. Schiltz, S. Iezzi, M. T. King, P. Zhao, Y. Kashiwaya, E. Hoffman, R. L. Veech, and V. Sartorelli. 2003. Sir2 regulates skeletal muscle differentiation as a potential sensor of the redox state. Mol. Cell 12:51-62.[CrossRef][Medline]
14. Gayther, S. A., S. J. Batley, L. Linger, A. Bannister, K. Thorpe, S. F. Chin, Y. Daigo, P. Russell, A. Wilson, H. M. Sowter, J. D. Delhanty, B. A. Ponder, T. Kouzarides, and C. Caldas. 2000. Mutations truncating the EP300 acetylase in human cancers. Nat. Genet. 24:300-303.[CrossRef][Medline]
15. Gillardon, F., R. Hata, and K. A. Hossmann. 1998. Delayed up-regulation of Zac1 and PACAP type I receptor after transient focal cerebral ischemia in mice. Mol. Brain Res. 61:207-210.[Medline]
) are supershifted by anti-Flag antibodies (
