(Received for publication, September 30, 1996, and in revised form, December 16, 1996)
From the Rosenstiel Basic Medical Sciences Research Center and
Departments of Biology and Biochemistry,
Brandeis University, Waltham, Massachusetts 02254-9110
The lymphoid-specific immunoglobulin µ heavy chain gene intron enhancer (µE) contains multiple binding sites for trans-acting nuclear factors. We have used a combination of in vitro and in vivo assays to reconstruct protein-DNA interactions on a minimal B cell-specific µ enhancer that contains three motifs, µA, µB, and µE3. Using ETS-domain proteins that transactivate the minimal enhancer in non-lymphoid cells, we show that (i) PU.1 binds coordinately to both µA and µB sites in vitro and (ii) in the presence of Ets-1, this factor binds to the µA site and PU.1 to the µB site. Two factors, TFE3 and USF, bind to the µE3 element. When the ETS proteins are present together with µE3 binding proteins, a three-protein-DNA complex is generated. Furthermore, we provide evidence for protein-protein interactions between Ets-1 and PU.1 proteins that bind to µA and µB sites, and between Ets-1 and TFE3 bound to the µA and µE3 sites. We propose that this domain of the µ enhancer is assembled into a nucleoprotein complex that contains two tissue-restricted ETS domain proteins that recognize DNA from the same side of the helix and one ubiquitously expressed bHLH-leucine zipper protein that binds between them, recognizing its site from a different side of the helix.
The immunoglobulin µ heavy chain (IgH) gene intron enhancer is a tissue-specific regulatory element that is necessary for expression of the IgH gene (1, 2). Since the proposal of Alt and colleagues of the correlation between transcription and immunoglobulin gene rearrangements (3-5), the µ enhancer has been considered a likely candidate to be the sequence that regulates rearrangements at this locus. Recently, this hypothesis has been directly verified in mice containing disrupted µ enhancer alleles (6, 7). Thus, the enhancer is activated at very early stages of B cell differentiation, prior to the onset of DH to JH rearrangements. In addition, experiments in transgenic mice have shown that this enhancer is sufficient to direct expression of heterologous genes in B and T lymphoid cells (8-11).
The µ enhancer contains binding sites for multiple DNA binding proteins, and tissue-specific activity has been proposed to be the consequence of both positive and negative regulatory proteins. Positive regulation is effected by lymphoid-specific transcription activators, such as those that bind to the µA, µB (12-15), and octamer elements (16-18). Negative regulation is effected by proteins found in non-lymphoid cells that may suppress enhancer activity (19-22), such as those that bind to the µE5 site within the enhancer (23, 24). In addition, there are several sites within the enhancer referred to as E motifs. These elements, µE1-µE5, fit the consensus CANNTG and bind proteins belonging to the basic helix-loop-helix family of transcription factors (25) that are expressed in both lymphoid and non-lymphoid cells. We have previously shown that mutation of either µA or µB elements abrogates B cell-specific activity of a 170 nucleotide enhancer (µ170) that contains both these elements as well as µE1, µE3, and µE5. Mutation of individual E motifs in the enhancer reduces, but does not abolish, enhancer activity (2). For example, a mutation in µE3 results in a 2-fold reduction in the activity of µ170 (14). Such results indicate functional redundancy among the several E motifs present in the µ enhancer. To simplify the enhancer for more detailed characterization, we have previously defined a 70-nucleotide minimal µ enhancer (µ70). This fragment contains only the µA, µB, and µE3 elements and activates transcription from a heterologous promoter in S194 plasma cells, when assayed either as a monomer or a dimer. Mutation of the µE3 element in µ70 results in a more drastic reduction of activity compared to the corresponding mutation in µ170. Based on these results we have proposed that the µA and µB elements define a minimal enhancer whose activity is raised by the presence of proximal E motifs (14).
The µA and µB sites bind members of the ETS domain protein family (14, 26, 27). Binding of the B cell and macrophage-specific factor PU.1 to the wild type µB element and a panel of three nucleotide substitution mutants within the element correlates with the ability of the mutants to activate transcription. These results suggested that PU.1 was a likely candidate for a functionally relevant µB-binding protein. The µA site binds the lymphoid-restricted Ets-1 protein, and mutations in µA, but not µB, affect Ets-1 binding to the µ enhancer in vitro. Furthermore, the minimal enhancer can be transactivated by co-expressing PU.1 and Ets-1 in COS non-lymphoid cells, thereby strengthening the idea that these factors can complement the lack of lymphoid-specific factors required for µ enhancer activity (14).
In this study we report the detailed interactions of ETS domain proteins and bHLH proteins with the minimal µ enhancer in vitro. Methylation and ethylation interference assays show that the contacts made by PU.1 protein on the µB element are very similar to those made by Ets-1 protein bound to its cognate DNA site. These results indicate that these two most distantly related members of the ETS family recognize DNA similarly. Furthermore, electrophoretic mobility shift and DNase I footprint assays demonstrate that PU.1 protein binds coordinately to both µA and µB elements, although the µA site has a significantly lower affinity for PU.1 compared to µB. In contrast, full length Ets-1 protein binds only to the µA site, within the range of concentrations used in these experiments. When both proteins are present together, Ets-1 and PU.1 bind to the µA and µB sites respectively to generate a tri-molecular complex. The third cis component of the minimal µ enhancer is the µE3 element. Two µE3 binding proteins, TFE3 and USF, can bind to µ enhancer DNA in the presence of ETS domain proteins to generate a three protein-DNA complex in which all three sites, µA, µB, and µE3, are occupied. Furthermore, we provide evidence for protein-protein interaction between PU.1 and Ets-1, and Ets-1 and TFE3. We propose that the µ70 enhancer is activated by a nucleoprotein complex that contains two lymphoid-specific ETS-domain proteins that recognize DNA from one face of the helix and one ubiquitous bHLH-leucine zipper protein that binds between them, probably recognizing the DNA from a different side.
Expression and Purification of Recombinant Proteins
GST.PUThe coding region of PU.1 was amplified using
polymerase chain reaction and cloned into SmaI-digested
pGEX-3X: 5 primer, 5
-G GGT AGG CCT ATG TTA CAG GCG TGC AAA ATG-3
;
and 3
primer, 5
-CTG CAC GCT CTG CAG CTC TGT G-3
. The 5
primer
introduces three additional amino acids between the coding regions for
glutathione S-transferase and PU.1. GST fusion proteins were
purified as described previously (28). An overnight culture of HB101
cells transformed with the appropriate plasmid was diluted 1:10 in
LB/ampicillin medium and grown for an additional hour. Expression of
the recombinant protein was induced by the addition of
isopropyl-1-thio-
-D-galactopyranoside to a final
concentration of 0.4 mM for a further 3-5 h. After induction the bacteria were harvested and resuspended in NETN buffer
(10 mM Tris, pH 8.0, 100 mM NaCl, l
mM EDTA, 0.5% Nonidet P-40). Extracts were prepared by
sonication (three pulses for 10 s each) at 0 °C. The insoluble
debris was removed by centrifugation, and the fusion protein was
purified from the supernatant by adsorption to glutathione-agarose.
Briefly, sonicated supernatant was mixed with the affinity matrix at a
ratio of 1 ml of a 50% slurry of beads for 1 liter of bacterial
culture, with shaking for 1-2 h at 4 °C. The slurry was poured into
a 10-ml column and washed with 50 column volumes of cold NETN three
times. Adsorbed proteins were eluted by the addition of reduced
glutathione at a concentration of 5 mM. One column volume
eluates were collected and analyzed by
SDS-PAGE,1 and the most concentrated
fractions were pooled and dialyzed against buffer D. Aliquots were
frozen in liquid nitrogen and stored at
70 °C.
A 1.2-kilobase pair PvuII-EcoR I fragment containing murine PU.1 cDNA was treated with the Klenow fragment of DNA polymerase plus dNTPs and cloned into XhoI-digested, Klenow-treated pET14b plasmid (Novagen). For protein expression, the plasmids were transformed into the BL21 bacterial strain, and proteins were purified as specified by the manufacturer (Novagen Inc.).
His.EtsThe coding region of murine Ets-1 was isolated as a BamHI fragment from the previously described expression vector pEVRF0-Ets (14). After filling in the ends, this fragment was cloned into pET-14b cut with BamHI and treated with Klenow to create blunt ends. His.Ets was expressed in Escherichia coli strain BL21 and purified by affinity chromatography.
Methylation and Ethylation Interference Assays
DNA fragments (residues 359-433) (29) from the wild-type,
µA, µB
, and M103 mutant enhancers
(14) were isolated as Sau3AI-BamHI fragments
(note that the BamHI site was introduced by site-specific mutagenesis of the first core site, and the mutation has no effect in
transient transfection assays) and cloned into pSP72 cut with BamHI. For labeling each strand, these plasmids were
linearized either with EcoR I or XbaI, treated
with calf intestinal phosphatase (Boehringer Mannheim), phosphorylated
with polynucleotide kinase and [
-32P]ATP, recut with
XbaI or EcoRI, respectively, and a 102-base pair
DNA probe purified by preparative acrylamide gel electrophoresis. Typical kinased preparations contained 4-6 × 106
Cerenkov counts/min/100 ng of DNA.
Methylation interference assays were carried out using probes described above and GST.PU fusion protein. In vitro electrophoretic mobility shift assays were used to titrate protein levels to obtain 30-50% of the probe retained in the nucleoprotein complex. Ten analytic binding reactions were pooled and loaded onto a preparative 4% acrylamide gel and free and bound DNA extracted by electroelution. Subsequent steps were as described (30).
Ethylation interference assays were performed as described (31) using the above probes and GST.PU fusion proteins. Alkali cleaved DNA was analyzed by electrophoresis through 8% polyacrylamide gels containing urea.
Yeast Two-hybrid Assay
The procedures of Gyuris et al. (32) were followed as described previously. LexA-PU.1 fusion proteins were generated in the vector pEG202. Full-length PU.1 (1-272) was isolated as a PvuII-EcoRI fragment, and a truncated PU.1 (101-272) was isolated as a NcoI-EcoRI fragment from pBS.PU. After treatment with Klenow enzyme to create blunt ends the fragments were cloned into pEG202 cut with EcoRI and filled in with Klenow.
Fusion proteins with the B42 transcription activation domain were
generated in the vector pJG4-5. pJG-PU, containing the full-length PU.1 gene cloned 3 of the transactivation domain was produced by
cloning the PvuII-EcoRI PU.1 fragment from pBS.PU
into pJG4-5 cut with EcoRI and treated with Klenow. The
XhoI-EcoRI fragment of pJG4-5 was replaced by
the XhoI-EcoRI polylinker fragment from pSP72 to
generate pJG72. pJG-Ets was generated by cloning the Ets-1 containing
BamHI fragment from pEVRF-Ets into pJG72 cut with
BamHI. The reporter plasmid used in these assays was
pSH18-34 containing the GAL1 promoter with eight LexA binding sites
fused to a lac Z gene. Quantification of
-galactosidase activity was carried out using a liquid assay as described previously (33).
DNase I Footprinting
The DdeI-HinfI fragment (residues
346-518) (29) from the wild-type, µA,
µB
, and M103 mutant enhancers was treated with Klenow
enzyme and cloned into pSP72 cut with EcoRV. To generate
probes for footprinting assays the plasmids were linearized with either
EcoRI or BglII, dephosphorylated with calf
intestinal phosphatase and radiolabeled with [
-32P]ATP
and polynucleotide kinase. The fragment was obtained by recutting with
either BglII or EcoRI, respectively, and isolated from preparative polyacrylamide gels. For footprinting reactions, 20,000 cpm of probe were incubated with PU.1 proteins in a final volume
of 50 µl containing 25 µl of DNA probe mix (1 µg of poly(dI-dC), 4% polyvinyl alcohol) containing 50 µg of bovine serum albumin in
Buffer D. After incubation for 10 min on ice 50 µl of 10 mM MgCl2 and 5 mM CaCl2
were added, followed by DNase I for 1 min to a final concentration of
17 µg/ml, determined empirically. DNase I digestions were quenched by
the addition of 90 µl of stop solution (20 mM EDTA, pH
8.0, 1% SDS, 0.2 M NaCl, 250 µg/ml tRNA), and the DNA
was purified after one extraction with phenol/chloroform (1:1) and
precipitated with ethanol. Samples were analyzed by electrophoresis
through denaturing 6% polyacrylamide/urea gels. Gels were dried onto
Whatman 3MM paper and visualized by autoradiography.
Partial Proteolysis Assays
His.Ets-1 alone, or together with His.TFE3, His-p50, or bovine serum albumin, was incubated in a final volume of 12 µl (containing 20 mM Hepes, pH 7.9, 100 mM KCl, 0.2 mM EDTA, 0.5 mM dithiothreitol, and 20% glycerol) for 10 min at room temperature. Proteolysis was initiated by the addition of 1 µl of 150 ng/µl trypsin and carried out for 15 min at room temperature. Reactions were quenched by the addition of 2 × SDS sample buffer and heating to 100 °C for 3 min, and proteins were separated by 12% SDS-PAGE. The proteins were transferred to nitrocellulose membranes and probed using an anti-Ets-1 antibody (Santa Cruz SC350, 1:1000). Chemiluminescence detection was carried out using the ECL immunodetection protocol according to the manufacturer's specifications (Amersham Corp.). His.Ets-1 and His.TFE3 were used at 400 ng/reaction, bovine serum albumin and p50 were used at 2 µg/reaction, and DNA was 20 ng of a 51-base pair double-stranded synthetic oligonucleotide containing the µA, µE3, and µB motifs.
Transient Transfection Assays
COS and NIH3T3 cells were transfected with calcium phosphate using reporter plasmids containing wild-type, or mutated, µ70 dimers as described previously (14). Transfections contained 2 µg of reporter and 2 µg of each transactivator. In those cases where only one, or no, transactivator was used, total DNA was made up to 6 µg using an empty expression vector. 48 h after transfection whole cell extracts were prepared and CAT enzyme assayed using 75 µg of heat-treated extracts for 1 h. S194 cells were transfected by the DEAE-dextran procedure using 5 µg of reporter plasmids and 2 µg of TFE3 expression vector, or the empty expression vector. Extracts prepared 48 h later were assayed for CAT enzyme expression by ELISA (CAT ELISA, Boehringer Mannheim).
To identify
nucleotides involved in PU.1 recognition of the µB element, we used
methylation and ethylation interference assays. Preparative binding
reactions were carried out with partially methylated µ enhancer DNA
followed by separation of the protein bound DNA and free DNA by
non-denaturing gel electrophoresis. Bound and free DNA were
electroeluted, treated with piperidine to effect strand cleavage, and
analyzed by electrophoresis through urea-containing sequencing gels. On
the coding strand, methylation of four contiguous guanines affected
PU.1 binding as shown by the significant depletion of the bands
corresponding to these residues in the bound DNA (Fig.
1A, compare lanes 4-6). On the noncoding strand, bands corresponding to three contiguous adenines were
diminished in the bound DNA (Fig. 1A, compare lanes
1-3) identifying these residues as being important for
nucleoprotein complex formation.
Interference assays to identify contacts of
PU.1 with the µ enhancer µB element. A, methylation
interference assay. A DNA fragment encompassing residues 359-433 of
the murine µ heavy chain gene enhancer was labeled with
[-32P]ATP and polynucleotide kinase on either the
coding or the noncoding strand, as described under "Experimental
Procedures." Probes were partially methylated using dimethyl sulfate
and used in in vitro binding assays with purified GST.PU.1
fusion protein. The protein-bound DNA and the unbound DNA were
separated by electrophoresis through nondenaturing polyacrylamide gels,
and the DNA was isolated by electroelution as described. Bound and free
DNAs were cleaved by piperidine treatment and analyzed by
electrophoresis through 6% polyacrylamide gels containing 8 M urea. Free (lanes 1, 3, 4, and 6), represents unbound probe DNA isolated
from a preparative mobility shift assay gel, and complex
(lanes 2 and 5) represents probe DNA present in
the nucleoprotein complex. Positions and sequences of the µB and µA
sites are indicated. On the noncoding strand, the missing bands in the
complex lane correspond to the three A residues and on the
coding strand to the four G residues within the µB element.
B, backbone phosphate contacts identified by ethylation
interference assays. Radioactive probes were as described in
A, except that chemical modification was induced by
treatment with ethyl-nitrosourea as described. Free and
Complex labels above the lanes are the same as described
above, and the positions of the µB and µA sites are indicated.
C, summary of interference assay results. Helical
representation of 12 nucleotides containing the µB element, TTTGGGGAA on the coding strand (read from top to
bottom). Location of phosphates whose ethylation interferes with
protein binding are marked with filled circles. In the
sequence shown beside the helix, residues marked with
asterisks are those whose methylation interferes with
protein binding and vertical lines represent phosphates
identified in the ethylation interference assays. The phosphodiester
bond between A and C on the noncoding strand (indicated by the
vertical arrow in the sequence) becomes hypersensitive to
DNase I when PU.1 binds to the µB element. Location of this
hypersensitive site is shown by the horizontal arrow on the
left of the helix. The horizontal lines, labeled
M102, M103, and M104, correspond to
mutations within the µB element that have been characterized by
transfections in B cell lines (12). Mutant 102 and 104 change the 3 Ts
to 3 Cs and the GAA to CCC, respectively, and are inactive in B cells.
Mutant 103 changes the 3 Gs to 3 As and is active in B cells.
Phosphate contacts were similarly assayed using ethylated probes in
in vitro binding assays. On the noncoding strand, ethylation of four phosphates at the 3 end of the µB site affected protein binding as shown by the absence of these bands in the bound DNA lanes
(Fig. 1B, lanes 1-3). Four additional phosphate
contacts were identified on the coding strand at the 5
end of the µB
element (Fig. 1B, lanes 4-6).
These results are summarized in the helix diagram and sequence shown in
Fig. 1C. The A and G residues identified by methylation interference studies are marked with asterisks and include
the two guanines that are part of the core GGA recognition site of ETS
domain proteins. Methylation at the N3 position of adenines lies in the
minor groove of the DNA and can affect protein binding by direct
interference or by altering DNA conformation. PU.1 binding was
diminished by methylation of 3 As on the noncoding strand lying up to
five nucleotides upstream of the core GGA. Phosphates identified by
ethylation interference are indicated by vertical lines in the sequence
and black circles in the helix diagram. Nye et al. (34) have
previously mapped at high resolution the DNA contacts made by Ets-1
protein. Methylation and ethylation interference assays described here,
as well as induction of a DNase I hypersensitive site on the noncoding
strand (see Fig. 6), indicate that the contacts made by PU.1 are
similar to those described for Ets-1. We conclude that these two most
divergent ETS domains recognize DNA by similar mechanisms.
Analysis of Mutation M103
We have previously analyzed a panel
of mutations within the µB sequence element by transfection into S194
plasma cells (12). Nucleotides modified by the three mutations
M102-104 are indicated by horizontal lines within the
sequence in Fig. 1C. In functional assays, M102 and M104
significantly decreased enhancer activity, whereas M103 had little or
no effect. The effects of mutations M102 and M104 are readily explained
by the strong adenine interferences that coincide with nucleotides
changed in M102 and alteration of the GGAA core, including the fourth G
identified by interference assays in M104. However, wild-type activity
of M103 is harder to explain since this mutation changed three guanine
residues that score strongly in the interference assay. In M103 the
three guanines indicated were changed to three adenines, so one
interpretation is that purine substitutions at these positions are
tolerated by the PU.1 protein. Alternatively, it was possible that PU.1 bound to a different GGAA sequence located 3 to the identified µB
site in M103 resulting in the observed enhancer activity. To distinguish between these possibilities, we studied PU.1 binding to
M103 in greater detail.
Methylation interference analysis showed that modification of the three
G residues located 3 of the M104 mutation did not interfere with PU.1
binding (Fig. 2A, lanes 6-8)
indicating that the putative downstream GGAA was not an alternate PU.1
binding site. Methylation of the remaining guanine (Fig. 1C,
position 10) in the µB element, that was unaltered by the M103
mutation, partially inhibited PU.1 binding. On the noncoding strand,
methylation of adenines at positions 4-6 (numbering as in Fig.
1C) affected PU.1 binding as observed with the wild-type
site, showing that the protein recognized the M103 mutant µB element.
Interestingly, we detected strong interferences with G and A residues
that make up the adjacent µA element which we have shown binds the
Ets-1 protein (Fig. 2A, lanes 6-8 marked
µA). No residues within the µA site on the noncoding
strand were identified by this assay (Fig. 2A, lanes
2-4).
Involvement of the µA site in the binding of PU.1 to M103 was further confirmed by ethylation interference assays. Ethylation of coding strand phosphates within the mutated µB sequence resulted in partial interference with PU.1 protein binding (Fig. 2B, lanes 7-9), whereas phosphate residues within the µA sequence interfered more strongly. On the noncoding strand as well, ethylation within the µA motif interfered strongly and those within the µB motif interfered weakly with PU.1 binding (Fig. 2B, lanes 2-4). First, these results show that PU.1 does not bind an alternate flanking GGAA site, downstream of the µB element, in the M103 mutation. DNase I footprinting experiments described below further confirm this observation. Second, significant interference caused by nucleotide or phosphate modifications within the µA site indicates that optimal binding of PU.1 to the M103 mutant requires dual recognition of both µA site and the mutated µB site. A lesser role of the µA element in the wild-type context suggests in that case most of the binding energy comes from PU.1/µB interaction.
DNase I Footprint Analysis of PU.1/µ Enhancer InteractionsOccupancy of both the µA and µB sites in the WT
and mutant enhancers was directly visualized by DNase I footprinting
assays. On the coding strand there was a conspicuous absence of bands corresponding to DNase I cleavage sites within the µB sequence with
no added protein (Fig. 3). We observed protection in two regions that correspond to the 5 end of the µB element and the 3
flank of the µB element. In addition, two weak DNase I hypersensitive sites that fell between the regions of protection described above, were
also induced in response to PU.1 binding (Fig. 3). Similarly, protection in the region of the µA element was also detectable, though weak. Surprisingly, a strong DNase I footprint was observed in a
region 3
of the µB element where several bands were protected and
one prominent hypersensitive band induced (Fig. 3). The µ enhancer
contains three elements homologous to the core element identified by
mutational analysis of the SV40 enhancer, and this footprint maps to
the region of the most downstream core site. Previous mutational
analysis concluded that these core sites were not required for B
cell-specific activity of the µ enhancer suggesting that PU.1 binding
to this site may represent a nonfunctional interaction (35).
PU.1 binding to all three sites on the enhancer was more clearly
discernible by analysis of the noncoding strand due to the induction of
a strong DNase I hypersensitive site within each element (Fig.
4A, lanes 2-6). Addition of
increasing amounts of GST-PU.1 fusion protein resulted in a
dose-dependent increase of a band within the µB motif
that was barely detectable in the absence of protein. A similar band
was induced within the µA element (labeled µA) and the
third core site (at the bottom of the gel). The location of the
hypersensitive site within the µB sequence is shown in Fig.
1C and corresponds to the position where a similar band is
induced in response to Ets-1 binding to its cognate sequence element.
Using DNase I footprinting, Rivera et al. (26) have previously shown that PU.1 binds to both µA (, in their
nomenclature) and µB sequence elements.
Analysis of the µB and µA
(inactive)
enhancer mutations showed that in each case only one of these two sites
bound PU.1. With the µB
DNA probe, no hypersensitive
band was observed at the µB site in the range of protein
concentrations used (Fig. 4B, lanes 2-6), whereas the hypersensitive site at µA was still induced although higher concentrations of PU.1 were required. This is consistent with
competition analysis that showed that the µA site bound PU.1 approximately 8-fold more weakly (data not shown). Conversely, only the
hypersensitive band corresponding to µB site occupancy was seen when
the µA
enhancer was analyzed (Fig. 4C,
lanes 2-6). Both mutated enhancers still showed PU.1
binding to the third core site as evidenced by the DNase I
hypersensitive site in this region (Fig. 4A-C, lower part
of the gel). In contrast, the functional M103 mutation bound PU.1
efficiently to both the mutated µB and the wild-type µA elements as
shown by the generation of the characteristic DNase I hypersensitive
bands at both positions with increasing amounts of PU.1 protein (Fig.
4D, lanes 2-6). These experiments directly demonstrate that mutations in µA or µB elements that inactivate the
enhancer allow PU.1 binding to only one site, whereas both sites can be
filled in the functional M103 mutation despite the lower affinity of
the M103 µB site for PU.1. These observations provide an explanation
for the observed activity of the M103 enhancer in transfection assays.
We propose that (i) interaction with the proximal µA site and (ii)
the pool of available PU.1 protein in S194 cells compensate for the
decreased affinity of the M103 µB site.
A GST.PU.1 fusion protein was used in these studies of PU.1 binding to wild-type and mutated enhancers. To rule out a possible contribution of the GST portion of the fusion protein, we analyzed the binding of two other forms of PU.1 to the wild-type µ enhancer. First, the GST part of the GST.PU.1 fusion protein was proteolytically removed by digestion with Factor X. Alternatively, the PU.1 cDNA was cloned in the vector pET14b to generate a hexahistidine tagged fusion protein (His.PU) in bacteria. Both new PU.1 derivatives showed similar patterns of binding to the µA and µB sites of the µ enhancer (data not shown).
Binding of PU.1 and Ets-1 to Minimal µ EnhancerPU.1
binding to both µA and µB sites of the wild-type and M103 mutant µ enhancers, or to only one or the other site in the µA and µB
mutated enhancers, correlates
well with the functional characteristics of the mutations in S194 B
cells. That is, those enhancers where both sites are bound by PU.1 are
active in transient transfection assays, whereas those enhancers where
only one site is bound are inactive. However, in COS cell
co-transfection assays, both PU.1 and Ets-1 are required for optimal
enhancer activity, indicating that PU.1 binding to both sites is not
sufficient for transcriptional activation (14). To analyze binding of
both ETS-domain proteins, we expressed polyhistidine tagged full length
Ets-1 (His.Ets-1) in bacteria and used the purified protein in DNase I
footprint assays.
DNase I footprint analysis of His.Ets-1 resulted in a protection
pattern over the µA site that was substantially different from that
observed with PU.1. On the noncoding strand (Fig.
5A), in addition to the hypersensitive site
within µA that was induced with either PU.1 or Ets-1 (marked with an
asterisk), several bands were prominently protected against
DNase I digestion only with Ets-1 (indicated by the open
triangles next to lane 2), but not with PU.1. Second,
in the range of protein concentrations used, Ets-1 did not bind over
the µB site as shown by (i) the absence of the strong hypersensitive
band within µB that is induced upon PU.1 binding and (ii) no
protection of the weak bands indicated by the black triangles that are
protected by PU.1 (Fig. 5A, lanes 10-13).
Addition of Ets-1 protein, in the presence of PU.1, showed a pattern of
protection and hypersensitive sites that indicated binding of Ets-1 and
PU.1 to the µA and µB sites respectively (Fig. 5A,
lanes 7-9). Specifically, Ets-1 binding to the µA site was shown by the protections over the µA site, whereas PU.1 binding to the µB site was shown by induction of the strong hypersensitive site within this sequence. Because the constant amount of PU.1 used in
these experiments was sufficient to fill both sites (Fig. 5A, lane 6), these results indicate that Ets-1
bound preferentially to the µA site.
A similar pattern was observed on the coding strand (Fig. 5B). PU.1 alone protected two sets of bands within the µB element indicated by the black triangles and induced three DNase I hypersensitive sites indicated by asterisks. PU.1 protection within the µA element was quite weak. Ets-1 alone (Fig. 5B, lane 5) strongly protected several bands within the µA site, indicated by the open triangles, but did not protect any bands or induce DNase I hypersensitivity within the µB site. Thus, PU.1-dependent protection and hypersensitivity within the µB site and Ets-1-dependent protection of the µA sequence characterize binding of each protein to these sites. Incubation of both proteins together showed a composite protection pattern (Fig. 5B, lanes 6 and 7), indicating that both sites were simultaneously occupied by the ETS-domain proteins. We conclude that although PU.1 alone can bind both µA and µB sites, in the presence of Ets-1 these sites are occupied by the two different ETS proteins. Taken together with the COS cell transfection data, we propose that the trimolecular PU.1-Ets-1-µ enhancer complex is the functional ETS domain protein complex.
Further Functional Analysis of the µ70 EnhancerThe minimal µ enhancer contains three elements, µA, µB, and µE3. A monomer, or a dimer, of this enhancer activates a heterologous promoter in S194 plasma cells, and we have previously shown that all three sites are necessary for enhancer activity in these cells. The µA and µB sites bind tissue-restricted ETS domain proteins, whereas the µE3 site binds several factors that contain a basic helix-loop-helix plus a leucine zipper domain (bHLH-zip), such as TFE3 and USF. Unlike the ETS domain proteins, µE3 binding proteins are ubiquitously expressed in a variety of cell types. However, in the earliest in vivo footprinting experiments, Ephrussi et al. (29) had shown that the µE3 site was bound by a factor in B cells only. To resolve the apparent inconsistency between the observed expression and in vivo DNA binding by µE3 proteins we proposed that the µ enhancer was inaccessible to µE3 binding proteins in non-B cells (36). Alternatively, it is possible that there exist B cell-specific forms of µE3 binding proteins which are necessary for µ enhancer function in B cells. Because the µ enhancer activates transcription in B cells only, it has been difficult to ascertain whether E binding proteins expressed in non-B cells can activate this enhancer or not.
Co-expression of PU.1 and Ets-1 transactivates the minimal µ enhancer in COS non-lymphoid cells (14). To determine whether the µE3 sequence contributed to enhancer activity in COS cells, we used a reporter plasmid containing a µE3 mutated enhancer fragment. This mutation decreases activity of the minimal enhancer by approximately 80% in S194 cells (14). In COS cell co-transfections, the combination of PU.1 and Ets-1 transactivated the minimal enhancer strongly, and mutation of either the µA, µB, or µE3 elements reduced activity approximately 10-fold (Fig. 6A). Therefore, the site usage in COS cells in the presence of transfected PU.1 and Ets-1 closely paralleled that seen in S194 cells in the absence of additional transfected trans-activators. Although we cannot rule out the possibility that the transfected PU.1 and (or) Ets-1 induced the synthesis of a B cell-specific form of a µE3 binding protein, we favor the interpretation that COS cell µE3 binding proteins can activate the µ enhancer in the presence of a co-transfected PU.1 and Ets-1.
To extend these results we assayed µ enhancer activity in a second non-lymphoid cell line. In NIH3T3 cells as well, co-expression of PU.1 and Ets-1 transactivated a minimal µ enhancer containing reporter plasmid (Fig. 6B), and mutation of either µA, µB, or µE3 sites significantly reduced enhancer function. We conclude that µE3 binding proteins present in these non-lymphoid cells can activate the µ enhancer only in the presence of transfected ETS domain genes. Furthermore, the combined transactivation and mutational analysis suggests that a two-protein DNA complex is not sufficient to reconstitute a functional enhancer. For example, since PU.1, or Ets-1, alone do not activate the enhancer despite the presence of endogenous µE3-binding proteins, it suggests that any one ETS protein plus a µE3-binding protein is not sufficient for enhancer function. Conversely, lack of activity of the µE3 mutation suggests that binding of both ETS proteins is also insufficient for activity. We therefore propose that the three-protein-DNA complex consisting of two ETS domain proteins and a µE3-binding protein is required for µ70 enhancer activity.
In the studies described above, we observed a requirement for
endogenous µE3-binding proteins present in non-lymphoid cells to
activate the minimal µ enhancer in the presence of transfected ETS
proteins. We further examined the effects of expressing exogenous µE3-binding proteins in the presence of endogenous ETS proteins that
activate the enhancer in B cells. As shown earlier, in S194 cells the
µ70 dimer is a functional enhancer whose activity requires both the
µE3 site and the µB site (Fig. 7, bars
marked with a minus sign, indicating the absence of
co-transfected TFE3 protein). Co-transfection of a murine TFE3
expression vector increased the activity of the µ70 enhancer
approximately 2-fold. Increased activity was dependent on all three
sites being intact as shown by the significantly reduced activity of
the µE3 or µB
reporter plasmids (Fig.
7, bars marked with a plus sign). These results
further strengthen the proposal that occupancy of all three sites in
the minimal enhancer are required for optimal enhancer function,
irrespective of whether the function is assayed in B cells or
non-lymphoid cells.
ETS and bHLH Proteins form a Multiprotein Complex on the µ Enhancer
Because transfection analyses suggested that µE3-binding proteins present in non-lymphoid cells can activate the µ enhancer, we used cloned, ubiquitously expressed bHLH zip proteins to determine whether a three protein-DNA complex could form on the µ enhancer. Full-length TFE3 (37, 38), a truncated TFE3 containing the DNA binding domain (39) and full-length USF (40) were expressed in bacteria as GST fusion proteins. DNase I footprint analysis showed that each protein bound to the µE3 element generating a protected region that extended toward the µA site (data not shown).
Co-incubation of µE3 binding proteins and PU.1, followed by DNase I
footprinting resulted in a pattern consistent with the simultaneous
occupancy of all three sites, µA, µB, and µE3. As described in
the preceding sections, addition of increasing amounts of PU.1 alone
generates two strong DNase I hypersensitive sites indicating occupancy
of both µA and µB sites (Fig. 8A,
lanes 2-6). In addition to these, several weaker bands in
both sites were protected against DNase I digestion. No protection was
observed in the core of the µE3 element, although three bands in the
3 end of the µA bracket were protected by either µE3-binding
proteins alone (data not shown) or by PU.1 binding to the µA site
(Fig. 8A, lanes 2-6) indicating a region of
partial overlap between the binding proteins. For the co-incubation
experiments we used a fixed amount of µE3 binding proteins that
results in complete protection over the µE3 element and added
increasing amounts of GST.PU corresponding to lanes 2-6 of
Fig. 8A. Under these conditions, we observed protection of
the µE3 sequences indicating occupancy of this site, as well as the
strong induction of DNase I hypersensitive sites within the µA and
µB motifs, indicating that both of these sites were filled as well
(Fig. 8A, lanes 7-21). We conclude that all
three elements of the minimal µ enhancer can be simultaneously occupied by factors: ETS domain proteins, such as PU.1, binding to the
µA and µB elements, and bHLH proteins, such as TFE3 and USF,
binding to the µE3 element. Analysis of the coding DNA strand was
consistent with this conclusion (data not shown).
Because enhancer activity requires Ets-1 in addition to PU.1, we also analyzed the binding of PU.1, Ets-1 and USF to the µ enhancer (Fig. 8B). On the coding strand, the footprints generated by Ets-1 and USF overlap considerably. Inclusion of all three proteins together resulted in a composite pattern (Fig. 8B, lane 6), indicating that all three proteins bound simultaneously to the enhancer.
Protein-Protein Interactions on the µ EnhancerVisualization of the minimal enhancer sequence in
double-helical form shows that the GGAA cores of the µA and µB
sites lie on the same side of the double helix. Thus, ETS domain
proteins bound to these sites are well positioned to interact with each other. We further investigated whether µA, µB, and µE3 binding proteins interacted directly with each other. Full-length PU.1, or an
N-terminal deletion mutant, were cloned in frame 3 of a LexA DNA
binding domain in the expression vector pEG202 (Fig. 9A) (32). PU.1 and Ets-1 sequences were also
cloned into the pJG vector containing the strong B42 transcription
activation domain (Fig. 9A, lines 4-6), and as
negative controls we used two other pJG derivatives shown in Fig.
9A (lines 7 and 8). Yeast strains
containing different combinations of the Lex and pJG derivatives and a
lac Z reporter plasmid were selected and assayed for lac Z expression.
Co-expression of PU.1 or Ets-1, containing pJG derivatives together
with PU.1 containing pEG derivatives consistently resulted in lac Z
expression as assayed by development of blue colonies in the presence
of X-gal. In contrast, pJG-Crk or pJG-Per co-expression with
pEG-PU(1-272) yielded only white colored colonies. To quantify the
results, three colonies were randomly selected from yeast strains
summarized in Fig. 9B, and
-galactosidase activity was quantified in extracts prepared from cultures. The average
-galactosidase activity obtained is shown in Fig. 9B. We
conclude that PU.1 can interact with itself or with Ets-1. Furthermore,
this interaction requires only the C-terminal 171 amino acids of PU.1
that contain the ETS domain.
We assayed possible interactions between Ets-1 and TFE3 proteins by
partial proteolysis. His.Ets-1 alone, or preincubated with other
factors, was treated with trypsin and assayed by immunoblotting after
SDS-PAGE. The anti-Ets-1 antibody used was directed against a
C-terminal peptide epitope and therefore reveals Ets-1 derivatives that
retain an intact C terminus. Brief treatment with trypsin showed a
major proteolytic fragment of an apparent molecular mass of 45 kDa
(Fig. 10A, lanes 2 and
7). Co-incubation of Ets-1 with TFE3 decreased the amount of
cutting at this site as indicated by the detectable amount of full
length Ets-1 under these conditions (Fig. 10A, lanes
3 and 8). This suggests that Ets-1 and TFE3 may associate in solution. To check whether DNA bound Ets-1 and TFE3 could
also interact we included a fragment of the µ enhancer in these
assays. In the presence of DNA, tryptic fragmentation of Ets-1 was
inhibited even further (Fig. 10A, lanes 4 and
9). The observed effects were specific for Ets-1/TFE3
interactions because neither the p50 subunit of NF-B nor bovine
serum albumin inhibited Ets-1 proteolysis by trypsin (Fig.
10A, lanes 5 and 6). Based on the size
of the tryptic fragment in SDS-PAGE, we estimate that the site of
proteolysis is toward the N terminus of Ets-1 as shown (Fig.
10B). These observations suggest that Ets-1 and TFE3 can interact on the µ enhancer.
We have previously shown that the µ70 domain of the Ig µ enhancer, that encompasses µA, µB, and µE3 elements, is a B cell-specific transcription activator. Transcription activity can be reconstituted in non-lymphoid cells by co-expressing the ETS domain proteins PU.1 and Ets-1 that bind to the µB and µA sites, respectively. Here we show that µ70 activity in non-lymphoid cells requires an intact µE3 element, suggesting that an endogenous µE3-binding protein is functionally recruited during the transactivation assay. Because transfection of PU.1 or Ets-1, individually, did not activate the µ70 enhancer in non-lymphoid cells, these observations suggest that neither ETS protein alone works efficiently together with the endogenous µE3 binding protein. Further evidence for the importance of all three sites was obtained by assessing the effects of TFE3 expression on µ70 activity in B cells that contain endogenous µA and µB binding proteins. Optimal transactivation of the µ70 enhancer by TFE3 also required intact µA and µB sites. Taken together, our analyses in lymphoid and non-lymphoid cell lines indicate that a three-protein-DNA complex, consisting of factors bound to the µA, µB, and µE3 sites, is required to activate the µ70 domain of the Ig enhancer.
We next examined the in vitro interactions of proteins that reconstituted minimal enhancer activity. We found that, despite a lower affinity for the µA site compared to the µB site, PU.1 bound both sites in vitro. These results provide a partial explanation for our previous observation that the M103 µB mutation that changes the core GGA in the PU.1 recognition site retains wild-type enhancer activity in B cells. Although the M103 mutation resulted in an approximately 4-fold weaker PU.1 site, the mutated enhancer bound PU.1 efficiently at both µA and µB sites. Furthermore, the contacts identified by interference assays for PU.1 recognition of the µB site were very similar to those described previously for Ets-1, indicating that these two most divergent ETS domain proteins recognize DNA similarly.
While this work was in progress, structural studies of the PU.1 and
Ets-1 ETS domains were reported (41-43). Our biochemical analysis of
PU.1/DNA interactions correlate closely with the x-ray crystal
structure of the PU.1 ETS domain in several aspects, but also point to
other interesting properties that are not easily evident from the
crystal structure. The most obvious similarity between the interference
and high resolution structures are the close correspondence of the
phosphodiester backbone contacts on both coding and noncoding strands,
and direct contact of PU.1 with nucleotides forming the core GGA
recognition site. Although no direct protein/DNA contacts were noted 5
of the GGAA sequence in the crystal structure, we found that three
adenosine residues on the noncoding strand located several nucleotides
upstream of the core scored strongly in methylation interference assays
(see Fig. 1C), suggesting possible involvement of these
residues in PU.1/µB interactions. Furthermore, the M102 mutation that
changes these nucleotides also reduced PU.1 binding and µ enhancer
activity significantly, suggesting that these residues contribute to
PU.1/DNA interactions. These observations indicate that the PU.1
recognition site extends at least five nucleotides upstream of the GGAA
sequence. Recognition of the upstream sequences may be mediated by
direct base-specific contacts by the protein, which were not detected in the crystal structure because of the particular sequence used in
that study. Alternatively, it is possible that sequence specificity at
these positions is determined by a combination of stereochemical and
electronic properties of the DNA helix (dependent on the sequence) without direct base recognition by the polypeptide. Because the methyl
group of methyl adenosines lies in the minor groove, these results also
indicate that the protein is located in the minor groove at this
position.
Our results differ most significantly from those of the Kodandapani et al. (41) by the identification of a strong DNase I hypersensitive site on the noncoding strand between the A and C residues (Fig. 1C). The crystal structure of the PU.1 ETS domain did not reveal any structural distortions in the DNA helix that could easily account for the generation of this hypersensitive site. We suggest that this site is caused by a PU.1-induced distortion of the helix. Furthermore, this distortion is probably not binding site-specific because a similar hypersensitive site was also induced within the µA element, when PU.1 was bound at that site.
In mobility shift assays full-length Ets-1 bound poorly to the µ enhancer. This was most likely due to the previously characterized inhibitory domain in Ets-1 (44-46) that lies just before the DNA binding ETS domain. Using DNase I footprint assays, we observed Ets-1 binding to the µA, but not the µB, site of the enhancer. When both ETS proteins were present together, Ets-1 bound to the µA site and PU.1 bound to the µB site to generate a ternary complex. Our observation that PU.1 and Ets-1 associate in the yeast two-hybrid assay supports the idea that the two proteins may interact when bound to the µ enhancer. We have recently shown that the ETS domain of PU.1 is sufficient to transactivate the enhancer together with Ets-1 (47), and it is interesting to note that the same domain of PU.1 is sufficient for interaction with Ets-1 in the two-hybrid assay. Our working model is that PU.1/Ets-1 interactions mediated by the ETS domain of PU.1 may provide a combined transactivation domain, either by jointly "touching" the basal transcription machinery directly or by jointly recruiting additional co-activators. Despite the correlation between the domains of PU.1 required for transcriptional activity and interaction with Ets-1, we note that the yeast two-hybrid assay does not closely resemble the B cell nucleus, and additional studies are required to prove the physiological relevance of this interaction.
As noted earlier, the two-protein-DNA complex discussed above is necessary, but not sufficient, for transcriptional activity of the µ70 enhancer. The third factor required to activate this enhancer is the µE3 binding protein(s). We show here that µE3 binding proteins can bind to the µ enhancer in the presence of ETS proteins to generate a three-protein-DNA complex, which we propose to be a functional unit of this B cell-specific enhancer. Crystal structures of bHLH zip family of transcription factors, of which TFE3 is one, have recently been reported (48, 49). These structures reveal that the basic residues take on an induced helical conformation and make contacts in the major groove of DNA. Contacts in the µA, µB, and µE3 regions suggest that the ETS-domain proteins recognize the µA and µB sites from the same side of the DNA helix, leaving the opposite side accessible to µE3 binding bHLH-zip proteins. The proposed structure has the interesting feature that the enhancer binding proteins appear to "coat" or "surround" the DNA. Perhaps this results in a weakening of the interaction of DNA with nucleosomal proteins that is manifested as altered chromatin structures in regions of the genome that contain active enhancers.
In addition we provide evidence that Ets-1 and TFE3 may interact directly. The trypsin sensitive site in Ets-1 that is masked in the presence of TFE3 is located in the N-terminal domain of Ets-1 that has no similarity to PU.1. The simplest interpretation of this observation is that the N-terminal domain of Ets-1 directly contacts TFE3. This is particularly interesting because we have shown that an N-terminal domain in Ets-1 is necessary for activation of the µ70 enhancer together with PU.1 (47). It is possible that this domain functions by recruiting TFE3 to the PU.1/Ets-1 bound µ enhancer, thus completing the functional quaternary nucleoprotein complex. Our studies suggest the following model of the µ70 enhancer. PU.1 and Ets-1 bind to the µB and µA sites of the enhancer, and interact via the PU.1 ETS domain. This interaction may be facilitated by DNA bending induced by PU.1 (50). TFE3 interacts with an N-terminal domain of Ets-1, while binding between the two ETS proteins on the other side of the DNA helix.
We are grateful to Dr. K. Calame for providing us plasmids for the bacterial expression of µE3 binding proteins. We thank B. Erman and D. Peisach for expertise in computer graphics, B. Nikolajczyk and K. Mama for comments on the manuscript, M. Rosbash and W. Jencks for helpful discussion, and Elaine Ames for preparation of the manuscript.