PU.1-mediated Transcription Is Enhanced by HMG-I(Y)-dependent Structural Mechanisms*

Robert T. Lewis, Amy Andreucci, and Barbara S. NikolajczykDagger

From the Departments of Medicine and Microbiology, Boston University School of Medicine, Boston, Massachusetts 02118

Received for publication, September 25, 2000, and in revised form, December 15, 2000


    ABSTRACT
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The ets transcription factor PU.1 is an important regulator of the immunoglobulin heavy chain gene intronic enhancer, or µ enhancer. However, PU.1 is only one component of the large multiprotein complex required for B cell-specific enhancer activation. The transcriptional coactivator HMG-I(Y), a protein demonstrated to physically interact with PU.1, increases PU.1 affinity for the µ enhancer µB element, indicating that HMG-I(Y) may play a role in the transcriptionally active µ enhanceosome. Increased PU.1 affinity is not mediated by HMG-I(Y)-induced changes in DNA structure. Investigation of alternative mechanisms to explain the HMG-I(Y)-mediated increase in PU.1/µ enhancer binding demonstrated, by trypsin and chymotrypsin mapping, that interaction between PU.1 and HMG-I(Y) in solution induces a structural change in PU.1. In the presence of HMG-I(Y) and wild-type µ enhancer DNA, PU.1 becomes more chymotrypsin resistant, suggesting an additional change in PU.1 structure upon HMG-I(Y)-induced PU.1/DNA binding. From these results, we suggest that increased DNA affinity under limiting PU.1 concentrations is mediated by an HMG-I(Y)-induced structural change in PU.1. In functional assays, HMG-I(Y) further augments transcriptional synergy between PU.1 and another member of the ets family, Ets-1, indicating that HMG-I(Y) is a functional component of the active enhancer complex. These studies suggest a new mechanism for HMG-I(Y)-mediated coactivation; HMG-I(Y) forms protein-protein interactions with a transcription factor, which alters the three-dimensional structure of the factor, resulting in enhanced DNA binding and transcriptional activation. This mechanism may be important for transcriptional activation under conditions of limiting transcription factor concentration, such as at the low levels of PU.1 expressed in B cells.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The immunoglobulin heavy chain intronic enhancer, or µ enhancer, is a regulatory region required for B lymphocyte-specific Igµ recombination and transcription (1, 2). Interaction between members of the ets family of transcription factors controls µ enhancer activation through the core enhancer sites µA and µB (3). The µB activator is the ETS protein PU.1, a hematopoietic-specific transcription factor (4, 5). PU.1 is required for B lymphocyte development, although overexpression of PU.1 promotes macrophage formation to the detriment of B cell development (6-8). Functional assays have demonstrated that the µ enhancer-binding proteins must be arranged in the proper stereospecific configuration for B cell-restricted activity characteristic of the enhancer in vivo (9). Although additional candidate members of the µ enhanceosome have been investigated (3, 10, 11), it is clear the complete composition of the transcriptionally active enhanceosome is unknown.

Assembly of an active enhanceosome can require not only transcription factors, but also DNA structure-determining proteins, such as HMG-I1 and its splice variant HMG-Y, referred to in the literature as HMG-I(Y) (12, 13). HMG-I(Y) can bind DNA in a sequence-specific manner, although DNA binding becomes promiscuous under modest overexpression in experimental systems (14). Because HMG-I(Y) lacks a transcriptional activation domain, any positive effects of the protein on transcription are thought to be mediated by its ability to define DNA structure and perhaps enhance binding affinity of activating transcription factors (15, 16). Alternatively, HMG-I(Y) can inhibit transcription through competitive binding of transcription factor interaction sequences (17) or modifying DNA topology (18, 19).

Transcriptional regulation by HMG-I(Y) is best understood for the interferon-beta enhancer, wherein HMG-I(Y) activates transcription through determining appropriate DNA structure (16). However, HMG-I(Y) utilizes multiple mechanisms to regulate numerous unrelated promoters. One example is the IL-2 promoter, on which HMG-I(Y)/DNA contact is unnecessary for HMG-I(Y)-mediated increases in transcription factor binding (15). On this promoter, HMG-I(Y) selectively enhances binding of c-Rel but not RelA to the promoter NF-kappa B site (20). For a second promoter, the IL-2 receptor alpha  promoter (IL-2Ralpha ), physical and functional interactions between HMG-I(Y) and an ets transcription factor, Elf-1, have been reported. Specifically, HMG-I(Y) and Elf-1 synergize to activate IL-2Ralpha in cells even in the absence of additional cotransfected transcription factors (21). Recent evidence indicates that HMG-I(Y) binds to the surface of IL-2Ralpha promoter DNA packaged into a precisely positioned nucleosome (13). Because HMG-I(Y) binds this positioned nucleosome in a directional manner, it was suggested that in addition to synergizing with Elf-1 to activate transcription, HMG-I(Y) plays a role in determining nucleosome positioning and remodeling, which in turn regulates transcription. Although the hypothesis that HMG-I(Y) dictates chromatin structure was not addressed directly, the study suggests yet another interesting mechanism for HMG-I(Y) as a transcriptional coactivator.

In addition to the multiple mechanisms proposed for HMG-I(Y)-mediated transcription, the function of HMG-I(Y) can be modulated by post-translational modification. For example, HMG-I(Y) inhibits transcription from the IgE promoter until a signal transduction cascade results in HMG-I(Y) phosphorylation, decreasing HMG-I(Y)/DNA binding and allowing transcription (22). Alternatively, the role of HMG-I(Y) in enhanceosome assembly can be influenced by its acetylation status (23). Whether post-translational modification determines the mechanism by which HMG-I(Y) alters transcriptional activation is untested.

Previous analysis demonstrated a physical interaction between HMG-I(Y) and PU.1, but the functional consequences of this interaction are unknown (24). Because HMG-I(Y) is a transcriptional coactivator of numerous promoters and enhancers through multiple mechanisms, we questioned whether HMG-I(Y)/PU.1 interaction mediates PU.1-dependent µ enhancer activation. To address this question, we initially tested whether HMG-I(Y) alters PU.1/µ enhancer binding. Because HMG-I(Y) increased PU.1/µB binding at low PU.1 concentrations characteristic of PU.1 levels in B cells, we examined the likely mechanism underlying this phenomenon. We have defined a novel, protein structure-based mechanism of HMG-I(Y) action unique from the DNA structural mechanisms previously described. Finally, we demonstrate that HMG-I(Y) is a functional coactivator of the µ enhancer. Overall, our studies mechanistically define a new structural component of the transcriptionally active µ enhancer and support the hypothesis that the µ enhanceosome requires HMG-I(Y) as an ETS protein coactivator.

    EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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Recombinant Proteins and Antibodies-- GST-tagged HMG-I plasmid was a generous gift of Dimitris Thanos (Columbia University), and the His-PU.1 construct has been previously described (25). Purification of recombinant protein was accomplished by standard methods (25). Because HMG-I and HMG-Y are considered functionally equivalent in transcriptional co-activation assays. The nomenclature HMG-I(Y) will be used throughout to avoid confusion with similarly designated HMG family members. Purified Sp1 and the Sp1-specific antibody were kindly provided by Herb Cohen (Boston Medical Center). Additional antibodies were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).

DNA Binding Assays-- Electrophoretic mobility shift assays (EMSAs), in vitro DNase I footprinting, and methylation interference assays have been previously described (9, 25, 26). For EMSA analysis, a PstI-BamHI fragment (base pairs 376-433 in the numbering system of Ephrussi et al. (27)) or a PvuII-BamHI fragment (base pairs 383-433) of the µ enhancer was analyzed. For supershift assays, 1 µl of anti-GST, anti-HMG-I(Y), or anti-PU.1 antibody at supershift concentrations (Santa Cruz Biotechnology) were added to the EMSA reaction prior to adding radiolabeled DNA. For methylation interference and footprint assays, we scanned a HinfI-DdeI µ170 enhancer fragment (base pairs 346-51). Circular permutation EMSAs in the pBend vector (28) and DNA bending calculations have been described in detail (25) and were based on earlier work (29, 30).

Oligonucleotides-- The coding strands of the annealed oligonucleotides are as follows: PRDII-NRDI, 5'-AGT GGG AAA TTC CTC TGA ATA GAG A-3'; µ enhancer, 5'-CAG CTG GCA GGA AGC AGG TCA TGT GGC AAG GCT ATT TGG GGA AGG GAA-3'; µB-enhancer, 5'-CAG CTG GCA GGA AGC AGG TCA TGT GGC AAG GCT ACC CGG GGA AGG GAA-3' (mutated residues are underlined).

All oligonucleotides were annealed to their complementary strands and purified on an 8% acrylamide gel before use.

Limited Protease Digestion-- Recombinant His-PU.1 was incubated with 100-250 ng of HMG-I(Y) in 10 µl of buffer D (20 mM HEPES, pH 7.9, 100 mM KCl, 0.2 mM EDTA, 0.5 mM dithiothreitol, 20% glycerol) plus 1.5 µg of bovine serum albumin for 10 min at RT (21 °C). The amount of PU.1 used was standardized based on DNA binding units and was sufficient to bind ~1 ng of DNA. The amount of HMG-I(Y) required to induce structural changes in PU.1 was determined empirically and was similar to the amount needed to increase PU.1/DNA binding in EMSA analysis. Trypsin or chymotrypsin analysis of DNA-bound PU.1 was done under EMSA conditions, with all components scaled up ~5-fold (1 µg of PU.1, 95 ng of HMG-I(Y), 2 ng of DNA, 1.5 µg of bovine serum albumin, 4.5 µl of buffer D, 100 ng of poly(dI-dC)·(dI-dC) in a 15-µl total volume). 0.5 mg/ml trypsin solution (Life Technologies, Inc.) was diluted 1:10 to 1:80 in buffer D on ice immediately prior to use. For chymotrypsin, a 1 mg/ml solution was diluted 1:4 to 1:16 prior to use. Preliminary experiments demonstrated that 1 µl of diluted trypsin or chymotrypsin added to the PU.1-containing mixture for 10 min at room temperature gave an optimal range of partial digest patterns for data analysis. Reactions were stopped by the addition of 5 µl of SDS-polyacrylamide gel electrophoresis loading buffer and boiling for 3 min. Protease products were separated on 12% SDS-polyacrylamide gels and blotted onto polyvinylidene difluoride membranes for detection with anti-PU.1 antibodies (1:200) according to the manufacturer (Santa Cruz Biotechnology). The approximate location of the enhanced trypsin cleavage site in PU.1 was estimated based on the molecular weight of the partial cleavage product relative to protein standards.

GST Pull Downs-- GST pull-down assays were performed according to Giese et al. (31) as modified by Tian et al. (32). Proteins were detected on Western blots with the appropriate antibodies after separation on 12% (PU.1) or 8% (Sp1) SDS-polyacrylamide gels. All blots were reprobed with anti-GST antibody to verify the addition of the approximately equivalent amounts of the appropriate GST-tagged proteins to the sample.

Transfections-- NIH 3T3 cells were grown in DMEM, 10% calf serum, 0.05% penicillin/streptomycin prior to transfection using the CaPO4 method. Schneider S2 Drosophila melanogaster cells were grown in Schneider cell medium (Life Technologies, Inc.) supplemented with 12.5% heat-inactivated fetal bovine serum and penicillin/streptomycin. We transfected 5 × 106 S2 cells with 5 µg of reporter DNA plus 2 µg of transcription factor or control pPAC DNA using the CaPO4 method. All cells were harvested 44-48 h post-transfection, and the presence of the CAT reporter protein was assayed by CAT enzyme-linked immunosorbent assay (Roche Molecular Biochemicals).

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Previous analysis demonstrated that PU.1 interacts with HMG-I(Y) (24). Our series of protein-protein interaction assays confirmed that PU.1 forms a stable complex with HMG-I(Y) in solution. Specifically, GST-tagged HMG-I(Y) precipitated full-length recombinant PU.1 in a GST pull-down assay as detected with a PU.1-specific antibody (Fig. 1A, lanes 1 and 3). In contrast, the GST tag alone did not stably interact with PU.1 (lanes 2 and 4). Recombinant PU.1 alone, shown in lane 7, comigrates with the HMG-I(Y)-interacting species in lanes 1 and 3. All lanes contained the appropriate GST-tagged proteins in approximately equivalent amounts, as demonstrated by reprobing the Western blots with an anti-GST antibody (data not shown). In addition, this result was confirmed by probing the HMG-I(Y)-interacting protein fraction with an anti-histidine antibody, which specifically detects the His tag covalently bound to the PU.1 protein. Results were identical to those shown with the anti-PU.1 antibody. To further test whether PU.1-HMG-I(Y) interaction was specific, GST-HMG-I(Y) was incubated with an unrelated transcription factor, Sp1. Sp1 was absent in Western analysis of the HMG-I(Y)-interacting protein fraction (Fig. 1B, lane 2), despite strong detection of Sp1 loaded directly on the gel (lane 5). Predictably, Sp1 failed to interact with GST as well (lane 1). These data clearly demonstrate that PU.1 and HMG-I(Y) specifically interact in solution.


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Fig. 1.   GST pull-down assays. A, Western analysis using alpha -PU.1 antibody after two independent pull downs with GST-tagged HMG-I(Y) (lanes 1 and 3) or GST alone (lanes 2 and 4). Recombinant proteins were loaded directly onto the gel as negative (lanes 5 and 6) and positive (lane 7) controls for alpha -PU.1 specificity. B, control GST pull down. Western analysis using alpha -Sp1 after pull down of purified Sp1 with GST alone (lane 1) or GST-tagged HMG-I(Y) (lane 2). Control recombinant proteins were loaded directly onto gel as indicated (lanes 3-5).

PU.1 is a critical regulator of µ enhancer transcriptional activation mediated through PU.1/µB binding. (3, 25). These in vitro analyses demonstrated that relatively high concentrations of PU.1 are required for PU.1-µB complex formation (see also data below; Fig. 2, A and C, lane 1). Interestingly, this complex is undetectable in EMSA analysis using B cell nuclear extracts and is very weak in analysis of macrophage nuclear extracts, which contain ~3-4 times the amount of PU.1 detected in B cells (9). Because HMG-I(Y)-transcription factor interaction increases binding of the transcription factor to its cognate DNA element in several model systems (12, 15, 20, 33, 34), we questioned whether physical interaction between HMG-I(Y) and PU.1 potentially facilitates PU.1-µB binding at low levels of PU.1 that more closely mimic conditions found in the B cell. We titrated PU.1 levels to establish the amount of PU.1 unable to form a detectable PU.1-µ enhancer complex in EMSA analysis (Fig. 2A, lane 2). Upon the addition of various amounts of HMG-I(Y) (lanes 3-5) but not the GST tag alone (lanes 6-8), a PU.1-µ enhancer complex was detected, indicating that HMG-I(Y) specifically increased PU.1 affinity for the µ enhancer. Both the less abundant upper and more abundant lower complexes comigrated with the complexes formed between high levels of PU.1 and µ enhancer DNA (lane 1). Because recombinant PU.1 prepared under denaturing conditions contains approximately equimolar amounts of the upper and lower complexes (data not shown), the upper complex probably represents misfolded recombinant PU.1 capable of DNA binding, while the lower complex is more similar to PU.1 synthesized in vivo. Note the absence of the upper complex in an independent PU.1 preparation shown in Fig. 2C. Increased PU.1/µ enhancer binding required appropriate stoichiometry between PU.1 and HMG-I(Y), as evidenced by data demonstrating that either sub- or superoptimal HMG-I(Y) concentrations had a minimal effect on PU.1 binding (Fig. 2, A, lanes 3 and 5, and B, lanes 3-6). As expected, HMG-I(Y)-mediated PU.1 binding required an intact µB site (Fig. 2B, lanes 7 and 8), confirming that PU.1 sequence specificity was not substantially altered in the presence of HMG-I(Y). We further demonstrated that PU.1, but not HMG-I(Y), is present in the HMG-I(Y)-induced EMSA complex by antibody supershift analysis (Fig. 2C). Under low PU.1 concentrations, the complex formed between PU.1 and µ enhancer DNA only in the presence of HMG-I(Y) (lane 3) was quantitatively supershifted by an alpha -PU.1 antibody (top arrow, lane 4). In contrast, the addition of either an alpha -GST antibody specific for the GST tag of the recombinant HMG-I(Y) (lane 5) or an alpha -HMG-I(Y) antibody (data not shown) had no effect on the EMSA complex. These results suggest that the EMSA complex contains PU.1 plus µ enhancer DNA, but not HMG-I(Y). These analyses were complemented by DNase I footprint analyses, wherein HMG-I(Y) increased PU.1 binding to the µB site (Fig. 2D, compare lanes 3 and 4) as evidenced by formation of a DNase I-hypersensitive site (arrow) characteristic of PU.1 binding µB at high PU.1 concentrations (lane 2). High levels of HMG-I(Y) inhibited PU.1/µB interaction in the footprint assays (lane 5), consistent with the hypothesis that appropriate HMG-I(Y)/PU.1 stoichiometry is critical for increased PU.1/DNA interaction. Although previous reports noted that HMG-I(Y) could competitively inhibit transcription factor binding (17), we hypothesize that inhibition of PU.1 binding at high HMG-I(Y) concentrations is due to sequence nonspecific HMG-I(Y)/µ enhancer interaction (discussed below). Our results are also consistent with previous analyses showing that overexpression of HMG-I(Y) substantially compromises the specificity of the protein, potentially blocking the µB site (14). In summary, these analyses demonstrate that PU.1-µB binding increases in the presence of HMG-I(Y).


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Fig. 2.   DNA binding analysis with PU.1 and/or HMG-I(Y). A, EMSA analysis with 200 ng of PU.1 (lane 1) or 15 ng of PU.1 (lane 2) plus decreasing amounts of HMG-I(Y) (62.5, 31.2, and 15.6 ng, lanes 3-5, respectively) or GST (identical amounts as HMG-I(Y), lanes 6-8). B, EMSA analysis with 200 ng of PU.1 (lane 1) or 30 ng of PU.1 (lanes 2-6) plus 0, 240, 120, 60, or 30 ng of HMG-I(Y) (lanes 2-6, respectively). Lanes 7 and 8 contain 30 ng of PU.1 alone (lane 7) or with 30 ng of HMG-I(Y) (lane 8). Radiolabeled probe was the wild-type µ enhancer (lanes 1-6) or the enhancer mutated at the µB site (lanes 7 and 8). C, EMSA analysis with 200 ng of PU.1 (lane 1) or 30 ng of PU.1 (lanes 2-5) plus 30 ng of HMG-I(Y) (lanes 3-5). alpha -PU.1 or alpha -GST antibody was added to samples run in lane 4 or 5, respectively. D, DNase I footprint analysis with PU.1 and HMG-I(Y). The limited DNase I digestion pattern after incubation with bovine serum albumin (lane 1) or 900 ng recombinant PU.1 (lane 2) is shown. Lanes 3-5 were digested after incubation with 255 ng of PU.1 plus 0, 50, or 200 ng of HMG-I(Y), respectively. The DNase I digestion pattern after the addition of 50, 200, or 500 ng of HMG-I(Y) alone is shown in lanes 6-8, respectively. The arrow at the right highlights the characteristic PU.1-induced DNase I-hypersensitive site in µB. Specific HMG-I(Y)/DNA interaction, absent in µ enhancer analysis, is generally detectable on other DNA sequences in this range of protein concentrations.

Several possible mechanisms could explain how HMG-I(Y) increases PU.1-µB interaction. First, PU.1 and HMG-I(Y) may cooperatively bind µ enhancer DNA. This possibility is unlikely, due to the observation that despite the size of the HMG-I(Y) fusion protein (~34 kDa), a trimolecular HMG-I(Y)-PU.1-DNA complex is not detectable in EMSA analyses (Fig. 2). Because formation of a protein-DNA or protein-protein tethered complex in EMSA is dependent on a sufficiently low off-rate (i.e. HMG-I(Y) must remain bound to the DNA or to PU.1 for the duration of the gel run) the lack of an HMG-I(Y)-PU.1-µ enhancer complex in EMSA is not definitive proof that HMG-I(Y) is absent from the cellular PU.1-DNA complex. However, HMG-I(Y) did not footprint µ enhancer DNA (Fig. 2D, lanes 6-8), in contrast to clear HMG-I(Y) footprints on interferon-beta , HLA-DR, and IL-2 receptor alpha  DNA (14, 21, 34). This result suggests that HMG-I(Y) does not bind µ enhancer DNA with sequence specificity. Preliminary EMSA analysis using recombinant HMG-I(Y) and a µ enhancer DNA probe demonstrated HMG-I(Y) binds the µ enhancer DNA, but this complex is poorly competed by a 50-100-fold molar excess of the sequence-specific HMG-I(Y) binding site from the PRDII-NRDI region of the interferon-beta enhancer (14). This result suggests that the EMSA complex represents nonspecific HMG-I(Y)/DNA binding (data not shown). This conclusion was further substantiated by isolation of the putative HMG-I(Y)-µ enhancer EMSA complex for methylation interference analysis. This technique failed to detect binding interference by methylation at specific adenosine residues (data not shown), although adenosine residues are critical for HMG-I(Y) binding to the interferon-beta enhancer (14). Finally, formation of an HMG-I(Y)-DNA complex is not augmented by the presence of PU.1 in EMSA or DNase I footprint (data not shown) as would be expected if HMG-I(Y) and PU.1 bind cooperatively. The lack of a specific HMG-I(Y)-µ enhancer DNA complex even in the presence of PU.1 discounts the possibility that HMG-I(Y) and PU.1 bind µ enhancer DNA cooperatively.

A second possible mechanism explaining how HMG-I(Y) increases PU.1 binding is suggested by the role HMG-I(Y) plays in increasing NF-kappa B binding of the interferon-beta promoter (12). Falvo et al. (16) have suggested that HMG-I(Y) is a key regulator of an NF-kappa B-induced DNA bend in the promoter and that subtle changes in DNA structure by HMG-I(Y) may lead to increased NF-kappa B binding and subsequent transcriptional activation. Because PU.1 binding bends µ enhancer DNA (25), we hypothesized that HMG-I(Y)-induced changes in DNA structure, perhaps mediated through direct PU.1/HMG-I(Y) contact, may facilitate PU.1 binding and result in altered DNA bending angles similar to changes documented in the interferon-beta system. To test this hypothesis, we measured DNA bending in circular permutation assays as previously described (25). In this EMSA-based analysis, multiple probes are designed such that the protein binding site, in this case µB, is located at different distances from the end of each probe (Fig. 3A). If a protein alters DNA conformation upon binding, the protein-DNA complexes migrate through the EMSA gel with different Rf values. Calculations based on nucleoprotein migration estimate the degree of DNA distortion by the protein (Ref. 25 and references therein). Fig. 3B recapitulates previous analyses demonstrating that PU.1 induces an ~48° bend in µ enhancer DNA as measured by circular permutation (lanes 1-4). Identical analyses with low levels of PU.1 plus HMG-I(Y) (lanes 5-8) show that HMG-I(Y) alters the PU.1-induced DNA bend by only 3-5°, a change that is well within the error of the analysis. We conclude that HMG-I(Y) increases PU.1-µ enhancer binding by a mechanism independent from alterations in DNA bending.


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Fig. 3.   DNA bending analysis on PU.1 with or without HMG-I(Y). A, circular permutation fragments 1-4 differ in the length of DNA separating the µB binding site from the end of the DNA probe. A change in migration of the DNA-protein complex in EMSA signifies a change in DNA structure upon protein binding. B, PU.1-µ enhancer complex with probes 1-4 (lanes 1-4). EMSA pattern with PU.1 + HMG-I(Y) on probes 1-4 (lanes 5-8). Free probe is in excess in all lanes (not shown).

A third potential explanation for HMG-I(Y)-induced PU.1/DNA binding is that the physical interaction between HMG-I(Y) and PU.1 (Fig. 1A) changes the structure of PU.1 to a higher affinity DNA binding species. To test this possibility, PU.1 was incubated with HMG-I(Y), and then the complex was subjected to partial proteolysis with trypsin. Changes in trypsin digest patterns with PU.1 alone compared with PU.1 plus HMG-I(Y) would indicate that availability of lysine or arginine residues in PU.1 (i.e. PU.1 structure) is altered by the PU.1/HMG-I(Y) interaction. PU.1 was incubated with either the GST tag (Fig. 4A, lanes 1-4) or HMG-I(Y) (lanes 5-8) before digestion with one of three different levels of trypsin. Western blot analysis detected resulting tryptic peptides of PU.1 with a PU.1-specific antibody. Intermediate (lane 7) and low (lane 8) levels of trypsin preferentially produced a 27-kDa PU.1 peptide when PU.1 was preincubated with HMG-I(Y) but not GST alone (Fig. 4A, arrow). Because the antibody interacts with the C terminus of PU.1, the preferential trypsin cleavage site can be mapped to the activation domain of PU.1 (Fig. 4B, arrow). Under these conditions, the DNA-binding ETS domain at the C terminus of the protein remained intact. The small molecular weight peptide strongly detected in the HMG-I(Y) plus PU.1 samples under high trypsin conditions (lane 6, peptide e) was variably detected and hence may not represent an HMG-I(Y)-induced structural change in PU.1. Similar analysis with chymotrypsin, which hydrolyzes proteins C-terminal to tyrosine, phenylalanine, and tryptophan residues demonstrated complementary results (Fig. 4C). Specifically, a repeatable decrease in chymotrypsin sensitivity of PU.1 was demonstrated upon the addition of HMG-I(Y), but not GST, to PU.1 in solution (compare lanes 1-3 with lanes 4-6 and 7-9), suggesting that HMG-I(Y) changes PU.1 to a more chymotrypsin-resistant structure. A trivial possibility is that HMG-I(Y) may decrease chymotrypsin activity by direct inhibition of enzymatic activity. This possibility is highly unlikely given that HMG-I(Y) does not affect chymotrypsin activity over a large range of enzyme concentrations on either purified bovine serum albumin or immunoglobulin (data not shown). Overall, these data demonstrate that PU.1-HMG-I(Y) interaction in solution results in a change in PU.1 structure and support the hypothesis that structural alteration is the mechanistic explanation for increased PU.1-µ enhancer binding in the presence of HMG-I(Y).


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Fig. 4.   Structural analysis of PU.1 with limited protease digestion. A, full-length PU.1 incubated with GST alone (lanes 1-4) or GST-HMG-I(Y) (lanes 5-8) treated with trypsin as indicated and separated on a 12% SDS-polyacrylamide gel. PU.1 fragments were detected on Western blots with alpha -PU.1 antibody. The arrow at the right highlights 27-kDa PU.1 tryptic peptide preferentially formed in the presence of HMG-I(Y). a-e, major tryptic peptides, corresponding to peptides a-e in Fig. 5A. B, approximate location of trypsin cleavage site resulting in the enhanced 27-kDa PU.1 peptide in A, lanes 7 and 8. The 5' transactivation domain of PU.1 was defined on multimerized PU.1 sites. The PEST domain is a protein-protein interaction domain containing two casein kinase II consensus phosphorylation sites. ETS DBD is the DNA-binding ETS domain of PU.1. C, PU.1 (lanes 1-9) was incubated with 100 ng of GST (lanes 1-3), 100 ng of HMG-I(Y) (lanes 4-6), or 250 ng of HMG-I(Y) (lanes 7-9) before digestion with alpha -chymotrypsin as indicated. Digestion products were visualized as in A. Blots are representative of at least three independent experiments using at least two different PU.1 preparations.

We next questioned whether the HMG-I(Y)-induced change in PU.1 structure mimics structural alterations PU.1 undergoes upon binding DNA under limiting PU.1 concentrations. These experiments were done only in the presence of HMG-I(Y), because PU.1 alone does not bind DNA at the concentrations tested. Relatively low concentrations of PU.1 were incubated with HMG-I(Y) either alone (Fig. 5A, lanes 1-3) or with the addition of wild-type µ enhancer DNA (lanes 4-6) under conditions resulting in formation of a PU.1-µ enhancer complex in EMSA analysis (i.e. a scale up of Fig. 2C, lane 3). Trypsin digest products, visualized on Western blots with alpha -PU.1 antibodies, demonstrated that the 27-kDa PU.1 polypeptide preferentially formed in the presence of HMG-I(Y) (27-kDa arrow at right, Fig. 4, lanes 7 and 8; Fig. 5, lanes 3, 8, and 9) was absent under conditions in which PU.1 bound µ enhancer DNA (Fig. 5A, lanes 4-6). In contrast, the 27-kDa PU.1 polypeptide was formed upon the addition of both HMG-I(Y) and a µB-mutated enhancer (lanes 7-9), confirming that formation of the three-dimensional PU.1 structure characterized by the 27-kDa trypsin product is lost upon PU.1/µ enhancer binding.


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Fig. 5.   Structural analysis on DNA-bound PU.1. A, PU.1 was incubated with HMG-I(Y) alone (lanes 1-3) or in the presence of wild-type (lanes 4-6) or µB-mutated µ enhancer DNA (lanes 7-9) before digestion with decreasing amounts of trypsin. Resulting polypeptides were visualized as described for Fig. 4. Peptides a-e correspond to peptides a-e in Fig. 4A. Although the overall peptide pattern is the same, relative representation of peptides in Figs. 4A and 5A differs due to increased trypsin digestion and a shorter run time in Fig. 5A. The arrow at the right labeled 27 kDa highlights the PU.1 tryptic peptide preferentially formed in the presence of HMG-I(Y) without PU.1/DNA interaction. B, duplicate samples from A were digested with chymotrypsin and then visualized as described in the legend to Fig. 4. Peptide 1 (arrow) highlights full-length (chymotrypsin-resistant) PU.1.

One possible explanation for the trypsin digest results is that PU.1 returns to its original solution structure after binding DNA, perhaps due to dissociation of HMG-I(Y). A second possibility is that PU.1 takes on a third structure that is not apparent at the level of trypsin cleavage patterns. To differentiate between these two possible explanations for the lack of trypsin hypersensitivity in DNA-bound PU.1, we repeated limited protease digest analysis with chymotrypsin (Fig. 5B). For these studies, PU.1 was incubated with µ enhancer DNA under conditions requiring HMG-I(Y) for formation of a PU.1-DNA complex. The reaction products were then subjected to limited chymotrypsin digestion. DNA-bound PU.1 is cleaved substantially less by chymotrypsin as compared with PU.1 in solution (compare the amount of peptide 2 in lanes 1-3 and lanes 4-6) or PU.1 incubated with a µB mutated enhancer fragment (lanes 7-9). Notable is the chymotrypsin-resistant full-length PU.1 protein (peptide 1) present only under conditions of PU.1/DNA binding (lanes 4-5). Overall, the chymotrypsin hydrolysis experiments indicate that PU.1 is most chymotrypsin-sensitive in solution, less sensitive in the presence of HMG-I(Y) (Fig. 4C), and relatively resistant in the presence of both HMG-I(Y) and target DNA (Fig. 5B). Because HMG-I(Y) is required for PU.1 to bind DNA under these experimental conditions, the simplest interpretation of the protease digestion data is as follows. Physical interaction between HMG-I(Y) and PU.1 results in a conformational change of PU.1, perhaps to a higher affinity DNA structure, resulting in formation of a PU.1-HMG-I(Y)-µ enhancer trimolecular complex. The stability of this proposed trimolecular complex is unclear, although EMSA analysis suggests that HMG-I(Y) may dissociate from DNA-bound PU.1. Chymotrypsin analysis suggests that PU.1 reconfigures into a third structure that is more chymotrypsin-resistant. Although there are limitations of both EMSA and structural determination by protease digestion, the data presented favor the following hypothesis: PU.1 has three unique structures determined by the presence or absence of HMG-I(Y) and DNA.

Experiments thus far have examined the potential mechanism of PU.1 coactivation by HMG-I(Y). Although HMG-I(Y) acts as a transcriptional coactivator with a second member of the ets transcription factor family, Elf-1 (21), function of the proposed HMG-I(Y)-PU.1 complex has not been demonstrated. To test the functional significance of PU.1/HMG-I(Y) interaction in cells we completed transient transfection assays in which a CAT reporter gene is transcriptionally activated by the µ enhancer. Because the full-length µ enhancer is transcriptionally silent in non-B cells (9), we assayed function of the tripartite µ70 enhancer defined by Nelsen et al. (3). This enhancer contains two ETS binding sites (µA, which binds Ets-1, and µB, which binds PU.1) flanking a basic helix-loop-helix-leucine zipper protein binding site, µE3. Dimerized µ70 (µ70)2 is synergistically activated by ectopic Ets-1 and PU.1 in conjunction with an endogenous basic helix-loop-helix-leucine zipper protein in both COS and NIH 3T3 cells (3, 32). A potential complication of functional assays was that all mammalian cell lines express endogenous HMG-I(Y), so the effect of ectopic HMG-I(Y) on PU.1-mediated activation of µ70 might be unclear given that HMG-I(Y) levels are critical for increased PU.1 binding (Fig. 2). Our initial studies therefore examined whether PU.1 and Ets-1 transactivate µ70 in the D. melanogaster Schneider cell line S2, a line that lacks endogenous HMG-I(Y) (35). Fig. 6A demonstrates that although Ets-1 or PU.1 alone activate only low levels of µ70-dependent transcription (1- and 3-fold of vector alone, respectively), Ets-1 plus PU.1 synergize to activate transcription ~10-fold in S2 cells compared with the pPAC Drosophila expression vector alone. Furthermore, cotransfection of Ets-1 plus HMG-I(Y) expression plasmids does not activate µ70-dependent transcription compared with empty expression vector alone. Similarly, cotransfection of HMG-I(Y) and PU.1 results in no increased activation of µ70 as compared with PU.1 alone (Fig. 6B). To test whether the combination of PU.1, Ets-1, and HMG-I(Y) activated µ70-dependent transcription over and above synergy of PU.1 plus Ets-1, all three expression vectors were transiently transfected into S2 cells followed by measurement of CAT reporter protein. At high levels of PU.1 (P1000, 1000 ng of PU.1 in Fig. 6A), HMG-I(Y) did not further activate transcription substantially (data not shown). We suggest that at high concentrations of PU.1, HMG-I(Y)-mediated increases in PU.1/µB interaction are no longer required for optimal PU.1 loading onto µB. To test this hypothesis, we analyzed transcriptional activation under conditions in which PU.1/Ets-1 transactivation was limited (i.e. 100 ng of PU.1 and 50 ng of Ets-1 expression vector; Fig. 6B). That combination of transcription factors activated µ70 ~5-fold over empty pPAC vector alone, compared with 3-fold activation with 100 ng of PU.1 alone. However, under limiting conditions for PU.1 and Ets-1, the addition of HMG-I(Y) increased transcriptional activation of µ70 12-fold over empty vector alone and 2.5-2.8-fold over Ets-1 plus PU.1 alone. Although the reproducible 2-3-fold increase in HMG-I(Y)-mediated transcriptional activity is not dramatically greater than activity with Ets-1 plus PU.1 alone, this level of transcriptional coactivation is characteristic of HMG-I(Y) in multiple enhanceosome complexes (15, 20, 34). Interestingly, suboptimally low or high HMG-I(Y) levels resulted in little synergy with PU.1 and Ets-1 (Fig. 6B). These data are consistent with biochemical results demonstrating that excessively low or high HMG-I(Y) levels cannot augment PU.1/µ enhancer binding in EMSAs or DNase I footprinting. Finally, mutation of either the Ets-1 binding µA site or the PU.1-binding µB site destroyed Ets-1/PU.1/HMG-I(Y) transcriptional synergy (Fig. 6C). We argue from these data that HMG-I(Y) synergizes with a PU.1-Ets-1 core activation complex to activate the µ70 enhancer in a µA-µB-dependent manner.


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Fig. 6.   Functional analysis of HMG-I(Y) on the µ enhancer. Transcriptional activity of a dimer of the tripartite µ enhancer (µ70)2 as measured by CAT reporter protein is shown. P, E, and H, PU.1, Ets-1, and HMG-I(Y) expression vectors, respectively. Numbers preceding letters denote ng of each expression vector transfected into the cells. A, synergy between PU.1 and Ets-1 in the S2 transfection system is shown by the E50 + P1000 bar; the rightmost bars show transactivation of the enhancer by Ets-1 plus HMG-I(Y). Comparisons are based on CAT measurements of samples transfected with the empty expression vector pPAC (leftmost bar). B, lack of synergy between PU.1 and HMG-I(Y) activating the µ enhancer (left bars) and synergy between PU.1 + Ets-1 + HMG-I(Y) under identical conditions (right bars). C, activation of mutated µ enhancer constructs in the presence of PU.1 + Ets-1 + HMG-I(Y). WT, µA-, and µB- designate the wild-type enhancer or the µ enhancer mutated at the µA or µB site, respectively. D, synergy among PU.1, Ets-1, and HMG-I(Y) in the mammalian cell line, NIH 3T3. Mock-transfected sample contains no DNA. pBluescript, used to equalize DNA amounts between transfections, was transfected alone as a negative control (bar at left). Values represent the amount of CAT protein relative to CAT-negative samples and are representative of 2-5 transfections done in duplicate (A-C) or triplicate (D). Error bars show the range of individual samples as compared with the average of those samples.

Although synergy among PU.1, Ets-1, and HMG-I(Y) was demonstrated in an HMG-I(Y)-negative Drosophila S2 cell background, we questioned the importance of this finding in mammalian cells containing endogenous HMG-I(Y). Because Ets-1 and PU.1 synergize to activate µ70 activity in NIH 3T3 cells (32), we tested whether ectopic HMG-I(Y) coupled with PU.1 and Ets-1 increased µ70 activity in 3T3 cells over PU.1 + Ets-1 alone (Fig. 6D). First, to confirm PU.1/Ets-1 synergy in 3T3 cells we transiently cotransfected cells with mammalian Ets-1 and PU.1 expression vectors along with the µ70 reporter construct. Ets-1 and PU.1 synergized to activate transcription 14-fold over irrelevant pBluescript DNA alone, as compared with 3.9- and 3.7-fold for Ets-1 or PU.1 alone, respectively. The addition of HMG-I(Y), PU.1, and Ets-1 together resulted in 38-fold transcriptional activation compared with the pBluescript control, a 2.7-fold increase over PU.1 + Ets-1 synergy. Thus, HMG-I(Y) coactivated PU.1/Ets-1-dependent transcriptional activity to approximately the same extent in 3T3s and Drosophila S2 cells. Overall, optimal amounts of HMG-I(Y) in the S2 system (25 ng, Fig. 6B) were 2-fold less than the amount of HMG-I(Y) resulting in high synergy in 3T3 cells (50 ng, Fig. 6D). Because transfection conditions for these two cell lines differed, and the results were not equalized based on the level of HMG-I(Y) expression in the two systems, significance of HMG-I(Y) levels cannot be compared directly. We believe the consistent increase of µ enhancer activity in these unrelated cell types is compelling enough to conclude that HMG-I(Y) coactivates µ enhancer-mediated transcription.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

These data clearly demonstrate that HMG-I(Y) increases PU.1/DNA interaction at the µ enhancer µB site in the absence of specific HMG-I(Y)/µ enhancer binding. Increased binding can be explained by a demonstrated HMG-I(Y)-induced structural change in PU.1, a new mechanism for transcriptional coactivation by HMG-I(Y). The absence of a trimolecular PU.1-HMG-I(Y)-µ enhancer complex in EMSA assays indicates that HMG-I(Y) may dissociate from PU.1 upon DNA binding, and chymotrypsin analysis is consistent with the hypothesis that PU.1 forms a unique structure upon HMG-I(Y)-induced DNA binding. Our data further suggest a functional role for HMG-I(Y) in µ enhancer activity, specifically a 2-3-fold increase in PU.1/Ets-1-mediated activity upon the addition of HMG-I(Y) to a transient transfection system. Overall, we propose that HMG-I(Y) is an important structural and functional regulator of PU.1-induced transcriptional activation.

Previous analysis has implicated members of the HMG family in transcriptional activation of the B cell-specific µ enhancer. HAF-1 and HAF-2 were originally identified based on binding and activation of the human µ enhancer, probably in conjunction with PU.1 (36). Like HMG-I(Y), neither HMG protein binds the mouse µ enhancer, but unlike HMG-I(Y), coactivation of the mouse enhancer by HAF-1/2 was not tested. In contrast to HMG-I(Y), HAF-1 and HAF-2 contain transcriptional activation domains; therefore, it is unlikely that activation of the human enhancer by a HAF/PU.1 combination is mechanistically identical to activation of the mouse enhancer by HMG-I(Y)/PU.1 synergy. The involvement of HMG family members in activation of both mouse and human enhancers may, however, reflect increased PU.1 binding due to structural alteration as suggested by our studies.

Enzymatic and circular dichroism analysis demonstrated that the ETS domain protein Ets-1 changes structure upon DNA binding (37, 38). In contrast, preliminary experiments show that dramatic protein structural changes are not detected after PU.1/DNA interaction. Instead, PU.1 structural changes are induced by interaction with a transcriptional coactivator, HMG-I(Y), in solution. This analysis suggests that Ets-1 and PU.1, although members of the same family of transcriptional activators, achieve activation though distinct mechanisms. Further evidence that Ets-1 and PU.1 activate transcription by different means was uncovered by Erman and Sen (39), who demonstrated that the full-length Ets-1 protein was necessary for activation of the µ enhancer, but the DNA binding domain of PU.1 alone was sufficient for maximal activation under the same conditions. The difference in the mechanism of transcriptional activation implied by both structural and functional analyses might be anticipated because Ets-1 and PU.1 are the most distantly related of ets family members (40).

Overall, the importance of HMG-I(Y)-induced PU.1 structural changes in transcriptional activation remains to be determined. Our data suggest that the PU.1 structure induced by HMG-I(Y) represents a molecule with higher affinity for DNA as compared with native PU.1. This interpretation is consistent with increased PU.1 chymotrypsin resistance in the presence of either HMG-I(Y) or HMG-I(Y) + DNA. Higher affinity DNA binding would be important for recruiting PU.1 to DNA under conditions of limiting PU.1 concentration, such as conditions present in B cells. In support of this hypothesis, PU.1 binding to the µB site is undetectable in EMSA analyses of B cell nuclear extracts despite the demonstrations that 1) DNA-binding PU.1 is present in B cell nuclei and 2) recombinant PU.1 binds the µB site in a variety of analyses (3, 25, 41). Phosphorylation of PU.1 increases PU.1 recruitment of other transcription factors to DNA but does not appreciably change PU.1 affinity in DNA binding analyses (42), discounting the possibility that post-translational modification determines PU.1/DNA interaction in cells. The simplest explanation of these experimental observations is that B cell PU.1 requires either a structural determinant such as HMG-I(Y) or an unidentified cooperative binding protein to bind and activate the µ enhancer. The likely role that HMG-I(Y)-induced PU.1 structure plays in the three-dimensional arrangement of the active µ enhanceosome or in the multiprotein complex that increases µ enhancer chromatin accessibility, the targesome (43), remains a future direction of our work.

The DNase I footprinting data and S2 functional data demonstrated that too high a concentration of HMG-I(Y) decreased PU.1/µ enhancer binding and functional synergy. These data are reminiscent of other biological systems in which overexpression of a protein leads to quenching of a biological phenomenon. For example, overexpression of transcription factors can induce squelching, a phenomenon whereby an overabundant transcription factor nonspecifically blocks transcriptional activation (44). Whether the ratio of HMG-I(Y) to PU.1 in B cells lies within the range required for increased PU.1 binding and transcriptional activation is dependent on several factors, including the pool of HMG-I(Y) available for interaction with PU.1 versus DNA or other transcription factors. Similarly, the stoichiometry required for our in vitro binding analyses ignores the demonstrated interactions between PU.1 and other components of the µ enhanceosome, specifically Ets-1 (41), that occur in cells. Overall, although the data suggest that HMG-I(Y)/PU.1 stoichiometry is important, quantitating that ratio awaits extensive biochemical and cellular analysis.

An important conclusion from the functional data is that Ets-1 is required for HMG-I(Y) to augment µ enhancer-mediated transcription. Three possibilities exist: 1) HMG-I(Y) may interact with PU.1 but not Ets-1, with Ets-1/PU.1 synergy being mediated solely through direct PU.1-Ets-1 contact (41); 2) two molecules of HMG-I(Y) interact in two separate complexes with Ets-1 and PU.1, affecting µ enhancer activity through two (similar or different) mechanisms; or 3) HMG-I(Y) interacts with Ets-1 and PU.1 simultaneously, forming a trimolecular Ets-1-PU.1-HMG-I(Y) complex either in solution on DNA. Whether HMG-I(Y)/Ets-1 interaction is important for function is an area of active investigation.

Functional synergy between PU.1 and HMG-I(Y) in the presence of Ets-1 is admittedly weak despite the consistency of the results in multiple experimental systems. Although robust transcriptional synergy with HMG-I(Y) has been reported (21, 33, 45), many groups report similarly weak synergy with HMG-I(Y) in multiple experimental models (15, 20, 34). All assays published to date, like ours, have studied HMG-I(Y) function in the context of transiently transfected reporter plasmids. Although the precise structure of transiently transfected DNA remains controversial, this extrachromosomal DNA clearly lacks the complex structure characteristic of chromatin-packaged cellular DNA. Chromatin structure is a critical regulator of both general and tissue-specific gene expression (46), and HMG family members including HMG-I(Y) play a role in determining DNA structure and transcription from chromatin templates (47, 48); therefore, functional synergy between transcription factors and HMG-I(Y) must be reexamined in the context of chromatin. Because PU.1 physically bends DNA upon binding (25) and alters µ enhancer accessibility in the context of cellular chromatin (49), the role of HMG-I(Y)-induced PU.1 structural changes in this context is especially intriguing.

    ACKNOWLEDGEMENTS

We thank M. Atchison, S. Ono, T. Maniatis, and D. Thanos for contribution of HMG-I(Y) DNA clones. Gifts of S2 cells and the pPAC vector from V. Zannis and Sp1 reagents from H. Cohen are greatly appreciated. K. McCarthy kindly provided recombinant His-PU.1 for Fig. 5A. We appreciate critical comments on the manuscript from K. McCarthy and G. Viglianti.

    FOOTNOTES

* This work was supported through funding from the Evans Biomedical Research Foundation at Boston Medical Center and the American Cancer Society (Massachusetts Branch and Grant IRG-72-001-24).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: Depts. of Medicine and Microbiology, Boston University School of Medicine, 650 Albany Street X-438, Boston, MA 02118. Tel.: 617-638-7019; Fax: 617-638-7140; E-mail: bnikol@medicine.bu.edu.

Published, JBC Papers in Press, December 20, 2000, DOI 10.1074/jbc.M008726200

    ABBREVIATIONS

The abbreviations used are: HMG, high mobility group protein; IL, interleukin; IL-2Ralpha , interleukin-2 receptor alpha ; EMSA, electrophoretic mobility shift assay; GST, glutathione S-transferase; CAT, chloramphenicol acetyltransferase.

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RESULTS
DISCUSSION
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