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
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ABSTRACT |
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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.
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- 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.
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).
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.
INTRODUCTION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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-
B site (20). For a second
promoter, the IL-2 receptor
promoter (IL-2R
), 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-2R
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-2R
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.
EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
GST pull-down assays. A,
Western analysis using -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
-PU.1
specificity. B, control GST pull down. Western analysis
using
-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 -PU.1 antibody (top
arrow, lane 4). In contrast, the
addition of either an
-GST antibody specific for the GST tag of the
recombinant HMG-I(Y) (lane 5) or an
-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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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-, HLA-DR, and IL-2 receptor
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-
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-
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-B
binding of the interferon-
promoter (12). Falvo et al. (16) have suggested that HMG-I(Y) is a key regulator of an
NF-
B-induced DNA bend in the promoter and that subtle changes in DNA
structure by HMG-I(Y) may lead to increased NF-
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-
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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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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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 -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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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.
|
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.
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DISCUSSION |
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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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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* 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.
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
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ABBREVIATIONS |
---|
The abbreviations used are:
HMG, high
mobility group protein;
IL, interleukin;
IL-2R, interleukin-2
receptor
;
EMSA, electrophoretic mobility shift assay;
GST, glutathione S-transferase;
CAT, chloramphenicol
acetyltransferase.
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