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The Journal of Immunology, 2005, 175: 5160-5169.
Copyright © 2005 by The American Association of Immunologists

Protein Acetylation Regulates Both PU.1 Transactivation and Ig{kappa} 3' Enhancer Activity1

Yuchen Bai2, Lakshmi Srinivasan, Leslie Perkins and Michael L. Atchison3

Department of Animal Biology, University of Pennsylvania School of Veterinary Medicine, Philadelphia, PA 19104


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Ig{kappa} gene expression and chromatin structure change during B cell development. At the pre-B cell stage, the locus is relatively hypoacetylated on histone H3, whereas it is hyperacetylated at the plasma cell stage. We find in this study that the histone deacetylase inhibitor, trichostatin A (TSA) stimulated 3' enhancer activity through the PU.1 binding site. TSA also stimulated PU.1 transactivation potential. PU.1 activity was increased by the coactivator acetyltransferase protein, p300, and p300 physically interacted with PU.1 residues 7–30. PU.1 served as a substrate for p300 and was acetylated on lysine residues 170, 171, 206, and 208. Mutation of PU.1 lysines 170 and 171 did not affect PU.1 DNA binding, but did lower the ability of PU.1 to activate transcription in association with p300. Lysine 170 was acetylated in pre-B cells and plasmacytoma cells, but TSA treatment did not stimulate PU.1 acetylation at this residue arguing that a second mechanism can stimulate 3' enhancer activity. Using chromatin immunoprecipitation assays we found that TSA caused preferential acetylation of histone H3 at the 3' enhancer. The relevance of these studies for PU.1 function in transcription and hemopoietic development is discussed.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Immunoglobulin gene rearrangement is an ordered process in which H chain gene rearrangement precedes L chain gene rearrangement (1). H chain DJ rearrangement occurs in pro-B cells, followed by VDJ rearrangement, whereas L chain genes undergo VJ rearrangement at the pre-B cell stage (1). B cells are defined as having both H and L chain genes rearranged. The same enzymatic machinery and DNA recognition sequences are used for rearrangement of H and L chain genes, and therefore, the mechanism of temporal specificity between the loci remains unclear. Transcriptional activation or chromatin accessibility models have been proposed to account for these differences (2).

The Ig{kappa} locus is controlled by two stage-specific enhancers, the intron enhancer, and the 3' enhancer (3). Both enhancers are inactive at the pro-B and pre-B cell stages, but are functional at the B cell and plasma cell stages (4, 5, 6). Both enhancers also contribute to the ability to rearrange the {kappa} locus in vivo (3, 7). Coincident with the development of enhancer activity and subsequent transcription, DNA at the locus changes from a heavily methylated, DNase I-resistant structure, to a hypomethylated, DNase I-sensitive structure (8, 9, 10, 11, 12, 13). The factors directly responsible for these changes are unknown, but presumably include the transcription factors that bind to the Ig{kappa} enhancers.

A number of studies explored chromatin accessibility at various developmental stages (3, 7, 10, 14). These studies revealed some site-specific changes in protein occupancy during development. For instance, hypersensitivity at certain sites in the 3' enhancer is increased in pre-B cells compared with pro-B cells (14), and these changes may contribute to control of {kappa} locus expression. However, simple protein accessibility to the {kappa} locus is not a sufficient mechanism for controlling transcription or recombination in pro-B cells, because in vivo footprinting studies showed proteins bound at the {kappa} enhancers before the onset of transcriptional or recombinational activity (14). Instead, increased activity of the 3' enhancer at later B cell stages must be controlled by the composition of proteins bound to the regulatory elements or to post-translational modifications of histones or regulatory proteins.

An important motif in the 3' enhancer is the PU.1/Pip (IFN regulatory factor-4) composite element. This motif is dependent upon transcription factor PU.1, which is necessary for the recruitment of Pip to DNA (15, 16, 17, 18). PU.1 is an erythroblast transformation-specific (ETS)4 domain transcription factor necessary for development of multiple hemopoietic lineages (19, 20, 21, 22, 23, 24, 25, 26, 27, 28). PU.1 homozygous mutants fail to develop lymphoid, myeloid, and granulocyte lineages and die in late gestation (27). Levels of PU.1 are implicated in regulating the lymphoid and myeloid cell fates (23), and PU.1 is down-regulated during erythrogenesis (26, 29, 30). Reduced levels of PU.1 also contribute to acute myeloid leukemia (31). PU.1 can physically interact with CREB binding protein (CBP) and elevated PU.1 expression can lead to changes in global histone acetylation levels in vivo (32, 33). PU.1 is also implicated in controlling chromatin structure at the Ig H chain enhancer (34). Therefore, PU.1 may influence both global histone acetylation patterns as well as local chromatin structure.

The factors that control {kappa} locus chromatin structure are presently unknown, although Pax5 has been suggested to play an important role (35). Previously we showed that H3 acetylation at the 3' enhancer is increased greatly in pre-B cells by treatment with the histone deacetylase inhibitor, trichostatin A (36). We find in this study that trichostatin A (TSA) treatment increased 3' enhancer activity, and this increase was largely through the PU.1/Pip composite element. The PU.1 transactivation potential was also increased in the presence of TSA or when cotransfected with the histone acetyltransferase (HAT) protein, p300. We find that PU.1 physically interacts with p300 and is a substrate for p300 acetylation with acetylation sites at lysines 170, 171, 206, and 208. We found that PU.1 is acetylated in vivo, and this acetylation may be important for PU.1 transactivation with p300. However, TSA treatment did not increase PU.1 lysine 170 acetylation, arguing that either other PU.1 lysines are targeted, or a second mechanism is needed for TSA induction of enhancer activity. Using chromatin immunoprecipitation methods we found that TSA causes preferential acetylation of histone H3 at the 3' enhancer. Our results provide evidence of two mechanisms for controlling 3' enhancer activity.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Chromatin immunoprecipitation (ChIP) assays

S194 cells (3.4 x 108) and 11 x 108 3-1 cells were fixed with 1.1% (v/v) formaldehyde, 100 mM NaCl, 0.5 mM EGTA, and 50 mM Tris-HCl (pH 8.0) in growth medium at 37°C for 10 min, then at 4°C for 50 min. Formaldehyde was quenched by adding 0.05 vol 2.5 M glycine. Fixed cells were washed with PBS, incubated for 15 min in 15 ml of 10 mM Tris-HCl (pH 8.0), 10 mM EDTA, 0.5 mM EGTA, and 0.25% (v/v) Triton X-100, followed by 15 min in 15 ml of 10 mM Tris-HCl (pH 8.0), 1 mM EDTA, 0.5 mM EGTA, and 200 mM NaCl, and finally sonicated in 1 ml of 10 mM Tris-HCl (pH 8.0), 1 mM EDTA, 0.5 mM EGTA, 1% (w/v) SDS plus 1 mM PMSF and a protease inhibitor mixture (1/100; P8340; Sigma-Aldrich). After sonication, cell debris was removed by centrifugation. Chromatin extracts were diluted to 6 OD 260 U/ml in IP buffer (140 mM NaCl, 1% (w/v) Triton X-100, 0.1% (w/v) sodium deoxycholate, 1 mM PMSF, 100 µg/ml yeast tRNA, and 100 µg/ml BSA); preincubated for 1 h at 4°C with 10 µl/ml 50% (v/v) protein A-agarose (Invitrogen Life Technologies); reconstituted in PBS, and washed several times in IP buffer. Aliquots (600 µl) were incubated with 24 µg of preimmune, anti-acetylH3, anti-acetyl H4 (Upstate Biotechnology), or p300 (Santa Cruz Biotechnology) Abs and incubated overnight at 4°C. Complexes were separated into bound and unbound complexes with protein A-agarose and cross-links were reversed by treatment at 65°C overnight. After treatment with RNase A and proteinase K, samples were extracted first with phenol/chloroform, then with chloroform, and precipitated with 2 vol of ethanol and 10 µg of glycogen (Roche). PCR was performed on 5-, 10-, and 20-ng DNA aliquots at 25 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 30 s using primers specific for the Ig{kappa} 3' enhancer, the intron enhancer, or the C{kappa} constant region (see Table I). Samples were dot-blotted to Hybond-N+ (Amersham Biosciences) and hybridized to a 1.1-kb EcoRI-SacI DNA fragment containing the 3' enhancer, a 473-bp AluI fragment containing the intron enhancer, or a 1.7-kb HindIII-BglII fragment containing the constant region.


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  		Table I. Primers used for construction of mutant plasmids or for ChIP PCR

 
Plasmids and recombinant proteins

Various full-length or mutant PU.1 plasmids were originally generated from a pBluescript KS+ PU.1 plasmid. PU.1 site-directed mutations were generated by overlap extension PCR (37) with primers containing the mutated sequences (Table I), followed by digestion with appropriate restriction enzymes. PCRs contained 30 ng of template DNA; 300 ng of each primer, 50 mM each of dATP, dCTP, dGTP, and dTTP; 10 mM Tris-HCl (pH 8.3); 50 mM KCl; 1.5 mM MgCl2; 0.001% (w/v) gelatin (PerkinElmer/Cetus); 2.5 U of AmpliTaq polymerase; and 2.5 U of Taq extender (Stratagene). The amplification cycle consisted of 35 cycles of 1 min at 95°C, 1 min at 50°C, and 2 min at 72°C, followed by one cycle of extension for 4 min at 72°C.

PCR products were either subcloned into KS+ pBluescript or directly cloned into pGEX-2T bacterial expression vector in-frame with the GST coding region. The pGEX plasmid PU.1 {Delta}201–272 was generated from pGEX-PU.1 by digestion with KpnI, followed by religation. The pGEX plasmids PU.1 {Delta}223–272 and {Delta}243–272 were generated by PCR with 3' primers p222 and p242, respectively (Table I). Plasmids PU.1 {Delta}7–30, {Delta}30–100, {Delta}119–160, and {Delta}245–272 were described by Pongubala et al. (15); {Delta}201–272 and {Delta}255–272 were described by Perkel and Atchison (38).

Plasmid CMV-p300 was a gift from Dr. P. Liberman (Wistar Institute, Philadelphia, PA). CMV-PU.1 was described by Pongubala et al. (16). (Oligo2)4TKCAT, (Oligo5)4TKCAT, (Oligo7)4TKCAT, and GALTKCAT were described by Pongubala and Atchison (4).

In vitro acetylation assays

The HAT domain of p300 or p300/CBP-associated factor (P/CAF) fused to GST were expressed in Escherichia coli (BL21), bound to glutathione-Sepharose beads, and eluted with buffer containing 25 mM glutathione. Eluted proteins were further purified on PD-10 columns (Sephadex G-25M; Amersham Biosciences), and eluted with buffer containing 20 mM Tris-HCl (pH 8.0), 0.5 mM EDTA, 100 mM KCl, 20% glycerol, 0.5 mM DTT, and 1 mM PMSF. Before storage at –80°C proteins were concentrated by Centricon (YW30; Amicon). Substrate proteins (GST fusions or chicken erythrocyte histones; Sigma-Aldrich) were incubated with 0.25 µCi of [3H]acetyl-coenzyme A (Amersham Biosciences) and 0.2 µg of purified enzyme in 30 µl of acetylation buffer containing 50 mM sodium butyrate. Reaction mixtures were incubated at 30°C for 30 min, stopped by addition of Laemmli buffer, and resolved by electrophoresis on 10% SDS-polyacrylamide gels. Gels were stained with Coomassie Blue to verify protein quantity, then subjected to autoradiography to evaluate acetylation activity.

EMSAs

Nuclear extracts were prepared from S194 plasmacytoma cells essentially as previously described (39), and binding reactions were performed as previously described (15). Briefly, EMSA was performed with 0.1 ng of labeled DNA probe (10,000 cpm) in a 20-µl reaction mixture containing 2 µg poly(dI-dC), 10 mM Tris-HCl (pH 7.5), 50 mM NaCl, 1 mM EDTA, 1 mM DTT, 5% glycerol, and 8 µg of nuclear extract or protein made by in vitro translation. Proteins made by in vitro transcription and translation were prepared using a coupled transcription and translation kit (Promega) with T3 or T7 RNA polymerase. The PU.1-Pip-binding site from the {kappa}3' enhancer used as probe is CTTTGAGGAACTGAAAACAGAACCT (Oligo 5). Samples were electrophoresed on 4% polyacrylamide gels in 6.7 mM Tris-HCl (pH 7.5), 3.3 mM sodium acetate, and 1 mM EDTA.

Cell culture and transfections

NIH-3T3 cells were grown in DMEM supplemented with 10% FCS and transfected by the calcium phosphate coprecipitation method (40). Cells were harvested 48 h after transfection. Each transfection contained 1 µg of a {beta}-galactosidase-expressing plasmid to normalize transfection efficiencies, 3–5 µg of reporter plasmid, and 3–5 µg of either empty expression vector (CMV) or CMV effectors. Transfection efficiencies were normalized using {beta}-galactosidase activity, and CAT assays, followed by TLC, were performed as described by Gorman et al. (41). The percent CAT activity was calculated by scintillation counting of the acetylated product and substrate spots. In each case, transfections were performed three to five times. S194 and 3-1 cells were grown and transfected as previously described (4). TSA treatment was performed for 24 h at a concentration of 33 nM.

Preparation of GST fusion proteins

GST fusion proteins were prepared essentially as previously described (42). Ten-milliliter cultures of E. coli BL21 cells containing the appropriate plasmid were inoculated overnight. The following morning, cells were diluted 20 times and incubated for another 3 h. Cultures were then induced with 0.25 mM isopropyl {beta}-D-thiogalactoside for 2–3 h at 30°C. Cells were spun down and subjected to a freeze-thaw cycle, followed by resuspension in 20 mM Tris-HCl (pH 8), 100 mM NaCl, 1 mM EDTA, 0.5% Nonidet P-40 (NETN) containing lysozyme, PMSF, leupeptin, and aprotinin. After incubation for 20 min on ice, cells received five 15-s sonication bursts. The suspension was centrifuged at 10,000 rpm in a Sorvall RTH-750 rotor for 10 min, and the remaining supernatant was incubated at 4°C with 0.5 ml of glutathione beads (50% slurry) for 2 h. The beads were spun down and washed three times with 5 ml of NETN before storage at 4°C.

GST chromatography

Reactions consisted of GST fusion protein or an equivalent amount of GST protein alone incubated with 5–15 µl of 35S-labeled protein prepared by in vitro transcription and translation in a 100-µl reaction containing NETN. Samples were rocked for 2 h at 4°C and washed at least five times with 450 µl of NETN. Samples were electrophoresed on 10% SDS-polyacrylamide gels for 1 h at 160 V, dried, and subjected to autoradiography.

Coimmunoprecipitation assays

NIH-3T3 cells were transfected with the appropriate plasmids and lysed in buffer containing 20 mM Tris (pH 7.5), 100 mM NaCl, 0.5% Nonidet P-40, 0.5 mM EDTA, and protease inhibitors. After removal of cell debris by centrifugation, lysates were immunoprecipitated with anti-FLAG M2 affinity agarose (Sigma-Aldrich) for 3 h at 4°C. Proteins bound to the affinity beads were washed extensively with lysis buffer, separated by SDS-PAGE, and then subjected to a Western blot procedure with anti-PU.1 and anti-p300 (Santa Cruz Biotechnology) Abs.

Generation of PU.1 acetyl lysine 170-specific antiserum

Rabbit antisera was prepared by Research Genetics against keyhole limpet hemocyanin-conjugated PU.1 peptide CGFTGSK-acetylK-KIRLY. The resulting antiserum was affinity purified, then cross-absorbed against the nonacetylated peptide. The specificity of the serum for acetylated PU.1 was tested by Western blot analysis with acetylated and unacetylated PU.1 (see Fig. 5C).



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  		FIGURE 5. PU.1 is acetylated in vivo. A, S194 cells were metabolically labeled with tritiated acetate for 90 min. Cell lysates were prepared and incubated with either preimmune or anti-PU.1 Abs, and bound proteins were fractionated by SDS-PAGE. The identity of the Ab used is shown above the lanes. B, Whole cell extracts from 3-1 and S194 cells were subjected to the Western blot procedure with anti-PU.1, anti-PU.1 acetyl lysine 170, or anti-actin Abs. C, The acetyl lysine 170 Ab is specific for PU.1 acetylated on lysine 170. Bacterially made GST-PU.1 protein was either untreated (lane 1) or acetylated with increasing quantities of p300 HAT domain (lanes 2–5) and Western blotted with anti-acetyl lysine PU.1 (top) or anti-PU.1 antisera (bottom).

 
Metabolic labeling experiments

S194 cells were labeled for 90 min with [3H]acetate (16 Ci/mmol; ICN Biomedical) in the presence of 33 nM TSA. Cells were lysed in 2 ml of 350 mM NaCl, 50 mM Tris-HCl (pH 7.5), 0.5% IGEPAL (Rhone-Poulenc), 1 mM EDTA, 0.5 mM DTT, 10 mM sodium butyrate, 0.2 mM PMSF, and 1 µg/ml each of aprotinin, pepstatin, and leupeptin. The sample was centrifuged at 10,000 x g for 5 min and diluted to 150 mM NaCl with 1 mM EDTA, 0.5 mM DTT, 10 mM sodium butyrate, and protease inhibitors. Equal quantities of cell lysate were precipitated with either preimmune or anti-PU.1 Abs coupled to protein A-agarose. Precipitates were washed five times with 150 mM NaCl, 50 mM Tris-HCl (pH 7.5), 1 mM EDTA, 0.5 mM DTT, 10 mM sodium butyrate, and protease inhibitors. Samples were fractionated by SDS-PAGE, and the gel was treated with Autofluor (Amersham Biosciences), dried, and exposed to x-ray film.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
TSA induces 3' enhancer activity and PU.1 transactivation

Previously, we showed that TSA could stimulate histone H3 acetylation at the Ig{kappa} 3' enhancer (36). The 3' enhancer is implicated in numerous important functions, including transcription, somatic rearrangement, and somatic mutation (6, 43, 44, 45, 46, 47). To explore whether TSA treatment could augment 3' enhancer activity, we transfected the pre-B cell line 3-1 with a reporter plasmid containing the 3' enhancer 132-bp core segment driving the CAT gene (CORETKCAT) (4). This reporter construct shows weak activity in pre-B cells (4). After transfection, cells were treated with either 33 nM TSA or vehicle control. Interestingly, TSA treatment caused a dramatic 7-fold increase in enhancer activity (Fig. 1A, lanes 1 and 2). The 3' enhancer contains several segments that show enhancer activity when multimerized (4, 48). These segments bind to c-Fos and c-Jun, PU.1 and Pip, or E47 and Pip (4, 48, 49). To determine the TSA-responsive region of the 3' enhancer, we performed transfections with reporter plasmids containing multimers of these enhancer elements in the presence or the absence of TSA. Weak TSA induction (2- to 3-fold) was observed with the c-Fos/c-Jun and the E47/Pip reporters (Fig. 1, lanes 3, 4, 7, and 8). This induction level was similar to that of the empty TKCAT reporter (2-fold; lanes 9 and 10). However, the activity of the PU.1/Pip reporter was very dramatically increased by TSA treatment (10-fold; Fig. 1, lanes 5 and 6). Because the PU.1/Pip reporter, but not the E47/Pip reporter, was induced by TSA, we reasoned that PU.1 was the most likely target of TSA induction. To confirm this, we performed transfections in NIH-3T3 cells (which lack PU.1) with the PU.1/Pip reporter plasmid and a plasmid expressing PU.1 (CMV-PU.1). TSA treatment resulted in a very dramatic increase in PU.1 transcriptional activity (10- to 20-fold; Fig. 1B). Therefore, PU.1 transcriptional activity can be greatly stimulated by an agent that inhibits histone deacetylase activity. In summary, our results show that histone deacetylase inhibition stimulated 3' enhancer activity and increased PU.1 transactivation function. Because DNA binding by PU.1 is not affected by TSA treatment (36), a distinct mechanism is responsible for increased PU.1 transactivation by TSA.



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  		FIGURE 1. TSA stimulates Ig{kappa} 3' enhancer activity by targeting PU.1. A, The pre-B cell line 3-1 was transfected with reporter plasmids containing either the 3' enhancer core (lanes 1 and 2) or multimers of enhancer fragments that bind to c-Fos and c-Jun (lanes 3 and 4), PU.1 and Pip (lanes 5 and 6), or E47 and Pip (lanes 7 and 8). Empty TKCAT vector is shown in lanes 9 and 10. One-half of each transfection received 33 nM TSA for 18 h. The presence (+) or the absence (–) of TSA is indicated above each lane. B, NIH-3T3 cells were transfected with CMV-PU.1 and a PU.1-responsive reporter plasmid. The presence or the absence of TSA is indicated above the lanes. C, p300 stimulates PU.1 transcription. NIH-3T3 cells were transfected with CMV-PU.1, a PU.1-reponsive reporter plasmid, and various coactivator proteins with HAT activity. The identity of the coactivator plasmid is shown above each lane. D, Map of the PU.1 mutants used for p300 activation mapping studies. Locations of the activation domain, the proline, glutamic acid, serine, threonine (PEST) domain, and the ETS DNA binding domain are shown. E, PU.1 residues 7–30 are needed for p300 activation. NIH-3T3 transfections contained various PU.1 deletion constructs (shown above the lanes) and the PU.1-responsive reporter. The presence (+) or the absence (–) of p300 expression plasmid is indicated above the lanes.

 
Proteins with HAT activity can induce PU.1-dependent transcription

A variety of coactivator proteins, including p300, CBP, general control nonderepressible, and P/CAF, have HAT activity. In light of our TSA results, described above, we tested the ability of these proteins to induce PU.1 transactivation potential. CMV-PU.1 was transfected with each coactivator, and transcriptional activity was assessed. PU.1 activity was stimulated by most coactivator proteins, but transactivation was stimulated the most by p300 (3-fold; Fig. 1C). P/CAF and CBP induced activity by ~1.5- to 2-fold (Fig. 1C). General control nonderepressible expression had little effect on PU.1 transactivation.

To determine the PU.1 sequences necessary for p300 induction, we performed transfections with various PU.1 deletion mutants (Fig. 1D). Deletion of PU.1 sequences 7–30 completely abolished p300 stimulation of PU.1 transactivation (Fig. 1E, lanes 1–4; 1.1-fold compared with 3.2-fold by wild-type PU.1). Residues 7–30 comprise a small portion of the PU.1 transactivation domain (50). Stimulation by p300 was relatively unaffected by deletion of PU.1 sequences 33–100 (Fig. 1, lanes 5 and 6; 2-fold) or sequences 119–160 (Fig. 1, lanes 11 and 12; 3.8-fold). Residues 33–100 constitute a major portion of the transactivation domain, and residues 119–160 define the PU.1 PEST domain. Deletion of the entire transactivation domain (residues 2–118) or the transactivation plus PEST domains (residues 2–160) abolished all PU.1 transcriptional activity. These mutants were also not able to respond to p300 stimulation (lanes 7–10). Therefore, p300 induction of PU.1 transactivation requires PU.1 residues 7–30.

PU.1 transactivation domain can physically interact with p300

Coactivator proteins are usually recruited to DNA by transcription factors. Activation of PU.1 by p300 suggested that these proteins might physically interact. PU.1 was previously shown to bind to CBP (32), but p300 interaction has not been tested. We found that GST-PU.1 interaction with p300 was readily detectable, whereas p300 showed little affinity for GST alone (Fig. 2A, lanes 1 and 2). Deletion of PU.1 residues 7–30 greatly reduced PU.1-p300 interaction, whereas deletion of PU.1 residues 33–100 had no effect (lanes 3–6). Therefore, physical interaction between PU.1 and p300 requires the same PU.1 segment needed for p300-dependent stimulation of transcription.



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  		FIGURE 2. PU.1 binds to p300. A, GST pulldown assays were performed with 35S-labeled p300 and the GST-PU.1 constructs indicated above the lanes. After incubation and washing, bound proteins were fractionated by electrophoresis on 10% polyacrylamide gels. The mobility of p300 is shown. B, Coimmunoprecipitation of PU.1 and p300. CMV-p300 was transfected into NIH-3T3 cells in the absence or the presence of CVM-FLAGPU.1. After immunoprecipitation with FLAG Ab, samples were subjected to Western blot with either p300 (top) or PU.1 (bottom) Ab. C, ChIP assays were performed with 3-1 cells using preimmune and anti-p300 Abs. Samples were subjected to PCR with the 3' enhancer primers, then dot-blotted against the 3' enhancer hybridization probe. One to 4% inputs are shown below.

 
To determine whether PU.1 and p300 interacted in vivo, we performed coimmunoprecipitation experiments. A CMV-p300 expression plasmid was transfected into NIH-3T3 cells either alone or in the presence of a CMV-FLAG-PU.1 expression plasmid. Samples were immunoprecipitated with FLAG Ab, then subjected to Western blot with either p300 or PU.1 Abs. Although p300 was present in both input samples (Fig. 2B, lanes 1 and 2), it was only detected in the anti-FLAG immunoprecipitates when FLAG-PU.1 was present (Fig. 2B, lanes 3 and 4). Therefore, PU.1 and p300 can interact in vivo. Because PU.1 binds to the Ig{kappa} 3' enhancer in B cells (36), we tested whether p300 was also recruited to the enhancer in vivo. ChIP experiments with anti-p300 Abs showed that p300 was indeed recruited to the 3' enhancer in vivo (Fig. 2C).

PU.1 is a substrate for p300 acetylation

The physical interaction between PU.1 and p300 raised the possibility that PU.1 might be a substrate for p300 acetylation. To test this, we performed in vitro p300 acetylation assays with GST-PU.1 using a truncated p53 protein and histones as positive controls. As expected, histones and the p53 truncated fragment were acetylated by p300 (Fig. 3A, lanes 2 and 3). Interestingly, p300 also acetylated GST-PU.1 nearly as efficiently as p53 (lane 4). In contrast, P/CAF efficiently acetylated p53, but failed to recognize PU.1 as a substrate (lanes 5–7).



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  		FIGURE 3. PU.1 is a substrate for p300-mediated acetylation. A, In vitro acetylation assays were performed with the HAT domain of p300 (lanes 1–4) or P/CAF (lanes 5–7). Assays were performed with BSA, chicken erythrocyte histones, p53 residues 300–393, or GST-PU.1. The substrate and acetylase proteins are listed above each lane. B, In vitro acetylation assays were performed with the p300 HAT domain and the various PU.1 mutants indicated above the lanes.

 
To define the p300 acetylation target site in PU.1, we tested several PU.1 deletions. Deletion of the C-terminal 30 aa had no effect on PU.1 acetylation (Fig. 3B, lanes 3 and 4, and Fig. 4A, constructs 1 and 2). However, no acetylation was observed with PU.1 construct PU.1 1–75 (Fig. 3B, lane 2, and Fig. 4A, construct 5). We tested additional PU.1 deletion mutants for acetylation by p300. PU.1 1–222 was acetylated as efficiently as wild-type PU.1 (Fig. 4A, construct 3), whereas PU.1 1–200 (Fig. 4A, construct 4) was slightly less efficiently acetylated. Therefore, the major sites of PU.1 acetylation lie between residues 75 and 222. Within this region, there are only nine lysine residues (lysines 169, 170, 171, 188, 196, 198, 206, 208, and 219). In the context of the PU.1 1–200 construct, we mutated the cluster of three lysines at positions 169, 170, and 171 to arginine residues. This completely eliminated p300 acetylation, indicating that one or more of these residues was acetylated by p300 and that residues 196 and 198 are not substrates (Fig. 4A, construct 6). Individual mutation of each lysine residue indicated that lysine 170 is the major target of p300 acetylation, residue 171 is a minor target site, and lysine 169 appears not to be acetylated (Fig. 4A, constructs 7–9).



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  		FIGURE 4. Identification of p300 acetylation sites in PU.1. A, PU.1 constructs used for p300 acetylation assays are diagrammed. The diagram shows PU.1 amino acid residues present in each construct, and Xs mark the positions of lysine residues mutated to arginine. The right panel indicates the relative level of p300 acetylation of each mutant construct as well as the specific lysine residues that are mutated to arginine. Relative acetylation was determined by comparison of multiple film exposures of HAT assay gels. Each + represents ~25% of the level observed with wild-type PU.1, which was defined as 100%. B, Representative HAT results showing relative acetylation levels. C, PU.1 sequences 161–220. *, Major p300 acetylation sites are marked by an asterisk (lysines 170, 206, and 208); {circ}, the minor site at lysine 171.

 
In contrast to the results reported above with PU.1 residues 1–200, mutation of lysine residues 169, 170, and 171 in the context of PU.1 sequences 1–222 did not completely eliminate PU.1 acetylation (Fig. 4A, construct 10). This suggested that lysines 206 or 208 might be acetylated by p300. Indeed, a PU.1 protein with lysine residues 169, 170, 171, 196, and 198 mutated to arginine yielded weak acetylation (Fig. 4A, construct 11). In addition mutant K169, 170, 171, and 206R (Fig. 4A, construct 12) or K169, 170, 171, and 208R (Fig. 4A, construct 13) showed weak acetylation, but mutation of residues K169, 170, 171, 206, and 208R (Fig. 4A, construct 14) abolished all acetylation. Similarly, mutant K169, 170, 171, 196, 198, 206, and 208R (Fig. 4A, construct 15) was not acetylated. Therefore, lysines 206 and 208 are weak p300 acetylation sites. We tested the same series of point mutations in the context of PU.1 sequences 1–242. These mutations yielded identical conclusions (Fig. 4A, constructs 16–21). The acetylation level of each mutant is summarized in Fig. 4A, and primary data showing acetylation of representative mutants are shown in Fig. 4B. The above results indicate that PU.1 lysine residues 170, 206, and 208 are major p300 acetylation target sites, and lysine 171 is a minor site. The p300 acetylation target sites in the PU.1 sequence are shown in Fig. 4C.

PU.1 is acetylated in vivo

Our acetylation studies above were performed with purified wild-type and mutant PU.1 proteins. To determine whether PU.1 is acetylated in vivo, we performed metabolic labeling studies with tritiated acetate in S194 cells. Cell lysates were prepared, and samples were incubated with either preimmune or anti-PU.1 Abs. The anti-PU.1 Ab precipitated a labeled protein with the electrophoretic mobility expected for PU.1 (~43 kDa; Fig. 5A), indicating that PU.1 can be acetylated in vivo.

Because PU.1 lysine 170 is a major acetylation site in vitro, we prepared antiserum specific for PU.1 acetylated on lysine 170 and used this antiserum to determine the acetylation status of PU.1 lysine 170 at the pre-B and plasma cell stages. Western blots of whole cell extracts from 3-1 and S194 cells were probed in parallel with PU.1 antisera and acetyl lysine 170-specific antisera. Interestingly, PU.1 was acetylated on lysine 170 at both developmental stages (Fig. 5B). The specificity of the Ab for acetylated PU.1 was shown by observing greater reactivity against p300 acetylated PU.1 than with unacetylated PU.1 protein (Fig. 5C).

Functional consequences of PU.1 acetylation

The major PU.1 acetylation sites that we identified in this study reside within the PU.1 DNA binding domain. Therefore, acetylation could potentially alter the ability of PU.1 to bind to DNA. We tested the DNA-binding ability of acetylated and unacetylated PU.1 by EMSA and found no discernable difference (although the fraction of acetylated PU.1 molecules was uncertain; data not shown). We also tested whether mutation of the PU.1 acetylation sites would alter DNA binding. PU.1 mutant proteins K170,171R and K206,208R were tested by EMSA for binding to the PU.1 site in the Ig{kappa} 3' enhancer and for their ability to recruit Pip to its adjacent DNA binding site. Mutant K170,171R bound to DNA and recruited Pip as efficiently as wild-type PU.1 (Fig. 6A, lanes 1–4). In contrast, mutant K206,208R bound to DNA poorly and was inefficient at recruiting Pip to DNA (lanes 5 and 6).



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  		FIGURE 6. Effect of mutation of PU.1-acetylated residues on PU.1 function. A, Wild-type PU.1 or mutated PU.1 (K170,171R and K206,208R) proteins were prepared by in vitro transcription and translation. Proteins were subjected to EMSA with the PU.1-Pip DNA binding site probe from the Ig{kappa} 3' enhancer. Proteins were assayed for DNA binding either alone or with S194 nuclear extract (NE), which provided a source of Pip to assess PU.1-Pip cooperative DNA binding. The proteins present in each assay are indicated above the lanes. Arrows indicate the positions of PU.1-DNA and PU.1-Pip-DNA complexes. B, Mutation of lysines 170 and 171 reduces p300-mediated stimulation of PU.1 transcriptional activity. NIH-3T3 cells were transfected with a PU.1-responsive reporter and plasmids expressing either wild-type PU.1 or the PU.1K170,171R mutant in the absence (–) or the presence (+) of p300. Although the K170,171R mutant activated transcription similar to wild-type PU.1, its induction by p300 was lower than that of the wild type. Error bars show the SD from the mean of three to six experiments. The lower panel shows a representative CAT assay. C, TSA does not increase PU.1 acetylation on lysine 170. 3-1, and S194 nuclear extracts were prepared from untreated and TSA-treated cells. Western blots with anti-PU.1 and anti-acetyl lysine 170 PU.1 Abs indicated no change in PU.1 acetylation in response to TSA treatment.

 
Because the K170,171R mutant showed normal DNA binding characteristics, we tested whether the transcriptional activity of this mutant was similar to that of wild-type PU.1 in response to p300 cotransfection. Interestingly, the K170,171R mutant was compromised in its ability to be stimulated by p300 (Fig. 6B). Therefore, lysine residues 170 and 171, which serve as target sites of p300 acetylation, are also necessary for maximal p300 stimulation of PU.1 transcriptional activity.

TSA causes increased PU.1 transactivation (Fig. 1B) and increased 3' enhancer activity (Fig. 1A). If PU.1 acetylation was responsible for these inductions, one would expect TSA treatment to increase the acetylation of PU.1. However, we did not find evidence that TSA treatment caused elevated PU.1 acetylation on lysine 170. 3-1 pre-B and S194 plasmacytoma cells were left untreated or were treated with TSA, and whole cell lysates were subjected to Western blot analyses with anti-PU.1 and anti-acetyl lysine 170 PU.1 antisera. TSA treatment did not result in a substantial increase in PU.1 lysine 170 acetylation (Fig. 6C). It should be noted that acetylation of other PU.1 lysine residues could potentially be increased by TSA. However, our results are also consistent with the possibility that TSA induction of 3' enhancer activity involves a second acetylation mechanism.

TSA induces H3 acetylation preferentially at the 3' enhancer

We previously showed that TSA can induce histone H3 acetylation at the 3' enhancer (36). If this increase in H3 acetylation plays a role in 3' enhancer activity, one would expect H3 acetylation to be greater at the 3' enhancer than at other segments of the Ig{kappa} gene. To test this, ChIP experiments were performed on 3-1 cells either treated with TSA or left untreated, and acetylation of histones H3 and H4 was measured at the intron enhancer, constant region, or 3' enhancer. We found histone H4 acetylation was increased between 2- and 3-fold by TSA at all locations across the locus (Fig. 7, lanes 3, 4, 7, 8, 11, and 12). In contrast, TSA treatment caused a nonuniform induction of histone H3 acetylation. H3 acetylation increased ~6-fold at the intron enhancer and ~3-fold at the constant region (lanes 1, 2, 5, and 6). Most impressively, TSA caused a 15- to 20-fold induction of H3 acetylation at the 3' enhancer (lanes 9 and 10). Thus, TSA preferentially activates H3 acetylation at the 3' enhancer compared with other locations within the {kappa} locus, suggesting that this acetylation contributes to increased enhancer activity. Thus, protein acetylation can influence Ig{kappa} 3' enhancer activity by two distinct mechanisms. First, acetylation can cause increased transactivation by PU.1. Second, acetylation of histone H3 correlates with increased enhancer activity.



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  		FIGURE 7. TSA preferentially induces H3 acetylation at the 3' enhancer. A map of the Ig{kappa} locus is shown at the top, depicting locations of the intron enhancer, the C{kappa} constant region, and the 3' enhancer. At the bottom are results of ChIP assays from 3-1 pre-B cells either untreated or treated with TSA, followed by immunoprecipitation with anti-acetyl H3 or anti-acetyl H4 Abs, and PCR amplification with primers specific for the intron enhancer, C{kappa} constant region, or 3' enhancer. The percent input bound was calculated and normalized to the level bound in untreated samples, which was defined as 100%. Error bars show the SD from three to 11 measurements.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Our results indicate that protein acetylation can influence Ig{kappa} 3' enhancer activity by two distinct mechanisms. First, the PU.1 transactivation function was increased by either inhibition of histone deacetylases via TSA treatment or cotransfection with acetyltransferase protein p300. PU.1 also served as a substrate for p300-dependent acetylation, and this acetylation was necessary for maximal PU.1 transactivation in association with p300. Thus, acetylation of PU.1 can increase PU.1 transactivation and, as a result, enhancer activity. However, because PU.1 acetylation on lysine 170 was unchanged in response to TSA, acetylation of this residue is not responsible for the increased PU.1 transactivation in response to TSA. Other PU.1 lysine residues could potentially be involved in this response. Second, elevated acetylation of histone H3 at the 3' enhancer in response to TSA treatment correlated with increased 3' enhancer activity. This suggests a chromatin structure mechanism for increased enhancer activity. Possibly, the TSA induction of enhancer activity is mediated by the chromatin structure changes we detected via H3 acetylation.

It is tempting to speculate that PU.1 is involved in the localized H3 acetylation we observed at the 3' enhancer after TSA treatment. We attempted to demonstrate this by transfecting PU.1 and p300 into NIH-3T3 cells and assessing H3 acetylation at the endogenous Ig{kappa} 3' enhancer. However, we did not observe a convincing increase in H3 acetylation in response to PU.1 and p300 transfection (data not shown). This is consistent with other work showing that PU.1 does not increase nuclease accessibility at the Ig{kappa} locus in non-B lineage cells (51). Similarly, we found that stable expression of PU.1 in NIH-3T3 cells did not result in appreciable PU.1 binding at the endogenous Ig{kappa} locus, suggesting that PU.1 does not gain access to the locus in fibroblasts (data not shown). Our data suggest that the PU.1 and H3 acetylation mechanisms for increasing enhancer activity may be distinct. PU.1 knockdown studies in B cells may be required to address this issue.

Because PU.1 DNA binding in vivo is not appreciably changed by TSA treatment (36), PU.1 may respond to TSA treatment by differentially recruiting coactivators or corepressors to the enhancer. Consistent with this idea, PU.1 can recruit the corepressor protein, Grg4, to the IgH HS1,2 enhancer in conjunction with Pax-5 (52). We similarly previously showed that Pax5 can repress Ig{kappa} 3' enhancer function in cooperation with PU.1 (53). In contrast, our results show that PU.1 can cooperate with p300 to activate transcription, and that p300 can increase the transactivation potential of PU.1. Thus, at some stages of B cell development, PU.1 might associate with corepressor proteins that are replaced by coactivator proteins at other stages or under other conditions. Interestingly, PU.1 can inhibit CBP HAT activity in vivo, leading to changes in global acetylation patterns (33). It will be interesting to determine whether acetylation of PU.1 by p300 changes this inhibitory phenotype.

Our studies showed that efficient p300 stimulation of PU.1 transcriptional activity required two distinct PU.1 segments. First, PU.1 residues 7–30 were needed for p300 stimulation, and p300 physically interacted with PU.1 via these residues. This segment comprises a portion of the PU.1 transactivation domain defined in transient expression assays (50). Interaction with PU.1 residues 7–30 was somewhat surprising because the p300-related protein CBP requires PU.1 sequences 74–122 for physical interaction (32). Therefore, p300 and CBP appear to interact with PU.1 by distinct mechanisms. We previously showed that the PU.1 7–30 region is also the target of Pax5 (B cell-specific activator protein)-mediated repression of PU.1 transcriptional function (53). Cotransfection of p300 reversed the Pax5-mediated repression of PU.1 transactivation (53). Because Pax5 and PU.1 can recruit the corepressor Grg4 to DNA (52), competition between Grg4 and p300 for interaction with PU.1 could constitute a molecular switch between repression and activation. Additional studies will be needed to test this hypothesis.

The second PU.1 segment needed for maximal activity with p300 was lysine residues 170 and 171. It is likely that p300 is first recruited via PU.1 residues 7–30, and then p300 subsequently acetylates PU.1 on lysine residues 170, 171, 206, and 208. According to the crystal structure of the PU.1 Ets domain on DNA (54), lysine residues 170 and 171 should lie very near DNA just downstream of the PU.1 binding site. Similarly, lysines 206 and 208 lie near the sugar phosphate backbone. This would place the above-mentioned lysines in a position conducive to protein interactions with adjacently bound proteins. However, lysines 170 and 171 do not appear to influence the binding of Pip to its adjacent DNA binding site (Fig. 6A), and p300 stimulation of PU.1 transcription does not require Pip. Thus, p300 stimulation must target some function other than PU.1 recruitment of Pip DNA binding. It will be interesting to determine the roles of these acetylation sites on PU.1 repression in association with Grg4. The reduced binding observed with the K206,208R PU.1 mutant is intriguing. Acetylation of these residues could potentially be used to regulate DNA binding. This would represent a novel mechanism of regulating PU.1 function.

PU.1 is implicated in numerous developmental processes, including differentiation of B cells, T cells, macrophages, and erythroid cells (19, 20, 21, 22, 23, 25, 26, 27, 28, 29, 30, 55). The interplay between PU.1 and either GATA-1 or Pax5 (B cell-specific activator protein) has been proposed to regulate the development of either the erythroid or macrophage lineage, respectively (23, 29, 30, 53, 56). More recently, the effects of modified PU.1 expression levels have been described (23, 31). Low PU.1 expression levels in progenitor cells generally give rise to B cell development, whereas higher levels yield macrophage development (23). In addition, reducing PU.1 expression levels in vivo to 20% of wild-type levels leads to a high incidence of acute myeloid leukemia (31). Thus, changing PU.1 activity by either expression level or, perhaps, its function via protein interactions or post-translational modifications could have a very dramatic impact on biological processes. Exploring how the p300-dependent acetylation sites identified in this study relate to various PU.1 functions will be an important goal.


   Acknowledgments
 
We are very grateful for the advice and reagents provided by Shelly Berger and Lin Liu for p300 in vitro acetylation assays. We thank Paul Lieberman for coactivator expression plasmids. We also thank Barbara Nikolajczyk for helpful comments on the manuscript.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by National Institutes of Health Grant RO1GM42415 (to M.L.A.). Back

2 Current address: 145 King of Prussia Road, Wyeth-Ayrest Research Laboratories, Radnor, PA 19087. Back

3 Address correspondence and reprint requests to Dr. Michael Atchison, University of Pennsylvania School of Veterinary Medicine, 3800 Spruce Street, Philadelphia, PA 19104. E-mail address: atchison{at}vet.upenn.edu Back

4 Abbreviations used in this paper: ETS, erythroblast transformation specific; CBP, CREB binding protein; TSA, trichostatin A; HAT, histone acetyltransferase; ChIP, chromatin immunoprecipitation; P/CAF, p300/CBP-associated factor. Back

Received for publication January 19, 2005. Accepted for publication August 11, 2005.


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 Materials and Methods
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 Discussion
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