From the Department of Molecular Biology, Princeton
University, Lewis Thomas Laboratory, Princeton, New Jersey 08544, and § Department of Molecular, Cell, and Developmental
Biology, Haverford College, Haverford, Pennsylvania 19041
Received for publication, January 9, 2001, and in revised form, March 23, 2001
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ABSTRACT |
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Smad proteins mediate transforming growth factor
TGF- The Smad proteins are functionally divided into three distinct classes:
(i) co-mediator Smads (Co-Smads), namely, Smad4 in mammals and Smad10
(also known as Smad4 A TGF- The Smad proteins are conserved across species, particularly in the
N-terminal MH1 domain and the C-terminal MH2 domain. The MH2 domain, to
which most of the tumor-derived mutations map, is responsible for
receptor recognition, transactivation, interaction with transcription
factors, and homo- and hetero-oligomerization among Smads. The MH1
domain, on the other hand, exhibits sequence-specific DNA binding
activity and negatively regulates the functions of the MH2 domain.
Formation of a heterocomplex between Co-Smad and R-Smad is
indispensable for the signaling process (2). The only known Co-Smad in
mammals, Smad4, forms a homotrimer in a
concentration-dependent manner both in vivo and
in vitro (9-11). The R-Smads, however, exhibit several
distinct oligomeric states at the basal state (11, 12). The complex
between Co-Smad and R-Smad has been suggested to be a heterohexamer
(9), a heterotrimer (10, 12), or, more recently, a heterodimer (11).
Understanding this stoichiometry has important implications for
understanding the molecular mechanisms of transcriptional regulation by
Smad proteins. To address this controversy, we have undertaken a
biochemical and biophysical approach, using purified homogeneous Smad
proteins. Results from both gel filtration and ultracentrifugation
analyses demonstrate that Smad2 and Smad4 form a stable heterodimer. In addition, 15 tumor-derived missense mutations were introduced into
these two Smad proteins to assess several prevailing models of
heterocomplex formation. Our results suggest a novel arrangement between Smad2 and Smad4.
Site-directed Mutagenesis and Protein Preparation--
Point
mutations were generated using a standard polymerase chain
reaction-based cloning strategy, and the identities of individual clones were verified through double-strand plasmid sequencing. All
Smad4 proteins were overexpressed in Escherichia coli strain BL21(DE3) at room temperature as a GST-fusion protein using a pGEX-2T
vector (Amersham Pharmacia Biotech). The soluble fraction of the
GST-Smad4 fusion in the E. coli lysate was purified over a
glutathione-Sepharose column, cleaved by thrombin, and further purified
by anion-exchange chromatography (Source-15Q; Amersham Pharmacia
Biotech) and gel filtration chromatography (Superdex-75, Amersham
Pharmacia Biotech). All recombinant Smad2 proteins were overexpressed
in E. coli strain BL21(DE3) at room temperature using a
pET-3d vector (Novagen). The soluble fraction of the cell lysate was
fractionated over a SP-Sepharose column and further purified by
anion-exchange chromatography (Source-15Q) and gel filtration
chromatography (Superdex-75). The identities of all proteins were
confirmed by mass spectroscopy. The concentrations of Smad4 and Smad2
were determined by spectroscopic measurement at 280 nm. All recombinant
proteins were characterized by gel filtration and dynamic light scattering.
Interaction Assay by Size Exclusion Chromatography--
Size
exclusion chromatography, using a Superdex-200 column (10/30; Amersham
Pharmacia Biotech), was employed to examine the interaction between
Smad4 and Smad2. In all cases, Smad2 is incubated with Smad4 at 4 °C
for at least 45 min to allow equilibrium to be reached. The flow rate
was 0.5 ml/min, and the buffer contained 25 mM Tris, pH
8.0, 150 mM NaCl, and 2 mM dithiothreitol. All fractions were collected at 0.5 ml each. Aliquots of relevant fractions
were mixed with SDS sample buffer and subjected to SDS-polyacrylamide gel electrophoresis. The proteins were visualized by Coomassie Blue staining. The column was calibrated with molecular mass standards.
Analytical Ultracentrifugation--
Protein samples were
prepared in 10 mM Tris-HCl, pH 8.0, 150 mM
NaCl, and 1 mm dithiothreitol. All sedimentation equilibrium experiments were carried out at 4 °C using a Beckman Optima XL-A analytical ultracentrifuge equipped with an An60 Ti rotor and using
six-channel, 12-mm path length, charcoal-filled Epon centerpieces and
quartz windows. Loading concentrations included 2.5, 5, 10, and 20 µM complex. Data were collected at four rotor speeds
(8,000, 11,000, 14,000, and 17,000 rpm) and represent the average of 20 scans using a scan step size of 0.001 cm. Partial specific volumes and
solution density were calculated using the Sednterp program (13). Data
were analyzed using the HID program from the Analytical Ultracentrifugation Facility at the University of Connecticut (Storrs,
CT). The ultracentrifuge figure was composed using IGOR Pro version
3.16 (WaveMetrics Inc., Lake Oswego, OR).
Electrophoretic Mobility Shift Assays--
All assays were
performed at 4 °C to offset the heat generated by electrophoresis.
The 6% polyacrylamide (37.5:1 acrylamide:bisacrylamide) gels were used
under the buffer condition of 65 mM Tris, pH 8.5, and 65 mM boric acid. All protein samples were prepared in a 40 µl volume containing 25 mM Tris, pH 8.0, 150 mM NaCl, 2 mM dithiothreitol, and 5% glycerol.
After prerunning the gels for 15 min, half of each sample (20 µl) was
loaded into each lane and subjected to electrophoresis with a constant
electric field of 15 V/cm. The gels were stained with Coomassie
Blue, destained, and dried for photography. After scanning the gels
with a densitometer, the unbound fractions of Smad4 were used to fit a
binary interaction equation, from which the estimate for the binding
affinity was obtained.
GST-mediated Pull-down Assay--
Approximately 0.4 mg of a
recombinant SARA fragment (residues 665-721) was bound to 200 µl of glutathione resin as a GST-fusion protein. The resin was washed
with 400 µl of buffer four times to remove excess unbound SARA (Smad
anchor for receptor activation) or other contaminants, and then 600 µg of a Smad2-Smad4 complex was allowed to flow through the resin.
After extensive washing with an assay buffer containing 25 mM Tris, pH 8.0, 150 mM NaCl, and 2 mM dithiothreitol, GST-SARA was eluted with 5 mM reduced glutathione, and all fractions were visualized
by SDS-polyacrylamide gel electrophoresis with Coomassie Blue staining.
Smad2 Forms a Stable Heterodimer with Smad4--
The full-length
Smad2 is unable to form a complex with Smad4 in the absence of
phosphorylation in its C-terminal SS*MS* sequence. This is likely due
to an autoinhibitory interaction between the MH1 and MH2 domains that
can be relieved by phosphorylation (14). Indeed, removal of the MH1
domain in Smads results in constitutively active transcriptional
activity (15) and allows the formation of a stable heterocomplex in the
absence of phosphorylation (9, 14, 16). Hence, we chose to focus on the
MH1-deleted proteins of Smad2 (residues 241-467) and Smad4 (residues
251-552). These recombinant proteins were overexpressed in bacteria
and purified to homogeneity. After confirmation of protein identity by
mass spectroscopy, both proteins were subjected to gel filtration and dynamic light scattering analysis to ensure that they are well folded
and exhibit good solution properties.
To examine the stoichiometry between Smad2 and Smad4, we devised an
in vitro interaction assay employing size exclusion
chromatography. We also used equilibrium sedimentation analytical
ultracentrifugation to characterize the molecular mass of the
hetero-oligomer. Because Smad proteins exhibit
concentration-dependent homo-oligomerization, we selected
the concentration ranges in which both proteins behave as monomers on
gel filtration analysis (Fig.
1A). The elution volume for
Smad2 corresponds to a molecular mass of ~22 kDa (Fig. 1A,
panel 1 (panels in Fig. 1A are numbered from
top to bottom)), consistent with its calculated
molecular mass of 25 kDa. The elution volume for Smad4, which contains
a 30-residue flexible loop (residues 460-490) between helices H3 and
H4 (9), corresponds to a molecular mass of about 39 kDa (Fig. 1A,
panel 2), in reasonable agreement with its calculated molecular
mass of 33 kDa. When equimolar amounts of Smad2 and Smad4 were used,
the vast majority of Smad2 was shifted to earlier fractions, indicating
a 1:1 stoichiometry (Fig. 1A, panel 3). In addition, the
elution volume for the complex corresponds to an apparent molecular
mass of ~54 kDa, consistent with the expected mass of a complex
composed of one Smad2 and one Smad4 (Fig. 1A, panel 3). To
further demonstrate the formation of a heterodimer, 1.5 molar
equivalents of Smad2 were mixed with one molar equivalent of Smad4. In
this case, the excess amount of Smad2 was eluted from the size
exclusion column as a monomer (Fig. 1A, panel 4). During the
course of size exclusion chromatography, there is little dissociation
between Smad2 and Smad4, suggesting stable complex formation.
Nevertheless, excess Smad2 (Fig. 1A, panel 4) or higher
concentrations of both proteins (Fig. 1A, panel 5) led to
more complete formation of a heterodimer.
To estimate the binding affinity between Smad2 and Smad4 and to further
confirm the formation of a heterodimer, electrophoretic mobility shift
assays under nondenaturing conditions were employed (Fig.
1B). Under these conditions, Smad2 does not enter the gel (Fig. 1B, lane 1) whereas Smad4 migrates as a single band
(Fig. 1B, lane 2). With increasing concentrations of Smad2,
a distinct heterocomplex is formed (Fig. 1B, lanes 3-8).
Neither excess Smad4 (lanes 3 and 4) nor excess
Smad2 (lanes 6-8) resulted in more than one heterocomplex.
Quantitation of the binding experiment revealed a dissociation constant
of ~1 µM.
Formation of a functional Smad2-Smad4 complex may be antagonized by the
formation of a complex between SARA and Smad2 (17, 18). To further
demonstrate the functional relevance of the observed heterodimer, the
Smad2-Smad4 complex was applied to glutathione resin preimmobilized
with GST-SARA (Fig. 1C). As expected, the flow-through
fraction contained less Smad2 than the input (Fig. 1C, lane
5), suggesting disruption of a Smad2-Smad4 complex by SARA.
Indeed, the eluted fraction contained a SARA-Smad2 complex (Fig.
1C, lane 7), demonstrating that SARA does compete with Smad4 for binding to Smad2.
To complement the gel filtration analysis, the molecular mass of the
hetero-oligomer was analyzed by analytical ultracentrifugation. The
complex was prepared in 1:1 ratio, based on the conclusion from the gel
filtration results that this complex forms a heterodimer. The complex
was analyzed at four loading concentrations and four rotor speeds. At 5 and 10 µM, the protein complex is fully consistent with
that of a heterodimer, with molecular masses of 58,900 and 59,800 daltons, respectively (Table I). If Smad2
and Smad4 form heterotrimers instead, then the apparent molecular mass
would be significantly reduced because one of the two proteins would be
in significant excess and would have contributed to a significant reduction in the reported molecular mass. In Fig.
2, we show the fit of the 10 µM data to a heterodimer model (Fig. 2). At 2.5 µM, we see evidence for a dynamic equilibrium between
monomers and heterodimers, whereas at 20 µM, there is
some evidence for minor aggregation (Table I). Indeed, Smad2 by itself
starts to homo-oligomerize and/or aggregate at the 20 µM
concentration. The best-fit two-state model to account for the
aggregation is a heterodimer-hetero-octamer model. In summary,
ultracentrifugation analysis demonstrates that Smad2 and Smad4 form a
heterodimer.
The Smad4 Activation Domain Is Required for the Formation of a
Stable Heterocomplex--
To investigate the sequence requirement for
the formation of a stable Smad2-Smad4 complex, we created deletion
mutants in Smad2 and Smad4 and examined their interaction with each
other. The results indicate that neither MH2 domain is sufficient for heterocomplex formation (Fig.
3A). In the case of Smad2, a
28-residue extension N-terminal to the MH2 domain (residues 269-467)
is required for stable interaction with Smad4 (Fig. 3A),
presumably due to the structural requirement that this fragment
contribute an additional Tumorigenic Mutations Disrupt the Formation of the Smad2-Smad4
Heterodimer--
Most of the tumor-derived missense mutations map to
the MH2 domain in Smad proteins (6, 9, 21). Because the formation of a
functional Smad2-Smad4 complex is important for signaling, some of
these missense mutations may act by disrupting this complex. Although
previous work shows that this is indeed the case in vivo (14), only four mutations were examined by immunoprecipitation, which
precluded a conclusion on whether or not these mutations directly
prevented formation of a heterocomplex.
To address this issue systematically, we generated a total of 18 missense mutations into either Smad2 (residues 241-467) or Smad4
(residues 251-552), purified the mutant proteins to homogeneity, and
examined their interactions with their wild-type counterparts. To rule
out the possibility of misfolding or aggregation, each mutant protein
was carefully compared with the wild-type Smad2 or Smad4 by gel
filtration, dynamic light scattering, and thermodenaturation analyses.
With the exception of three insoluble mutants (L440R and P445H in Smad2
and I527R in Smad4), these analyses demonstrated that each of the
mutant proteins was well folded and exhibited very similar solution
properties. Each of eight tumor-derived mutations was introduced in
both Smad2 and Smad4. For example, D450H in Smad2 has been reported in
colon cancers (22). Both D450H in Smad2 and the corresponding D537H in
Smad4 were generated. Each of the three tumor-derived mutations in
Smad4 (D351H, R361H, and V370D), as well as their corresponding
mutations in Smad2 (D300H, R310H, and V319D), disrupted the formation
of a heterocomplex (Table II; Fig.
4A). On the other hand, D450H
in Smad2 and D537H in Smad4 failed to disrupt the formation of a
heterodimer (Table II; Fig. 4A). Two additional tumorigenic
mutations (F346V in Smad2 and R420H in Smad4) and their corresponding
mutations (W398V in Smad4 and W368H in Smad2) exhibited no effects on
heterocomplex formation (Table II; Fig. 4). Interestingly, each of the
tumorigenic mutations in the original Smad and the corresponding
mutation in the other Smad has identical effects on heterocomplex
formation (Fig. 4B). This result suggests that Smad2 and
Smad4 may form a pseudosymmetric heterodimer, in which each of the two
Smad proteins uses similar surface motifs to interact with the other.
Because three of the four deleterious mutations affect residues that
are located in the L1 and L2 loop region (9) (Fig. 4B), this
loop-helix region must be directly involved in mediating the formation
of a heterodimer between Smad2 and Smad4. In support of this
observation, this loop-helix region contains the overwhelming majority
of the most highly conserved and solvent-exposed residues in Smad
proteins (9).
Comparison of Several Distinct Models for Heterocomplex
Formation--
Assessment of the effects of these tumorigenic
mutations and other reported evidence allows us to evaluate several
existing models on the formation of a heterocomplex between Smad2 and
Smad4. The initial heterohexamer model was proposed on the basis of
structural features of a homotrimeric Smad4 (9) but lacks experimental support. Subsequently, using gel filtration to assay overexpressed and
epitope-tagged Smad proteins in mammalian cells, it was shown that the
size of the heterocomplex between Smad2 and Smad4 is smaller than that
of a Smad2 homotrimer (12); however, for reasons that are unclear, a
heterotrimer model was proposed (12). More recently, using gel
filtration of endogenous Smad proteins in mammalian cells, it was
suggested that Smad2 forms a heterodimer with Smad4 (11).
In the heterotrimer model, the organization of the three subunits is
believed to be very similar to that of the Smad4 homotrimer, except
that one or two copies of Smad4 are now replaced by Smad2 (10, 12)
(Fig. 5A). This model is
inconsistent with several lines of experimental data. First, our
mutational data contradict this model. Asp-450 in Smad2 and the
corresponding Asp-537 in Smad4 play a central role in this model, each
making three intersubunit hydrogen bonds (9). Mutation of this residue
is expected to completely disrupt a network of hydrogen bonds, leading
to the disruption of the heterotrimeric packing (Fig. 5A).
However, neither D450H in Smad2 nor D537H in Smad4 disrupted the
formation of a heterocomplex (Table II; Fig. 5). Second, according to
the heterotrimer model, SARA binding to Smad2 should not interfere with
Smad2-Smad4 interactions because the SARA-binding surface of Smad2 is
far away from the proposed heterotrimeric interface (19); however, SARA-Smad2 and Smad2-Smad4 complexes appear to antagonize each other
(17, 18) (Fig. 1). Third, the heterotrimer model is inconsistent with
the observation that the SAD domain is important for the formation of a
heterocomplex (Fig. 3) because SAD is located in the periphery of the
proposed heterotrimer (Fig. 5A). Fourth, a Smad2-Smad4
heterotrimer would have a larger molecular mass than that of a Smad2
homotrimer, which is inconsistent with the observation that the
apparent molecular mass of a functional complex between Smad2 and Smad4
is appreciably smaller than that of a Smad2 homotrimer or a Smad4
homotrimer (11, 12). Fifth, according to the crystal structure of a
transcriptionally active Smad4 fragment, the C-terminal phosphorylated
SS*MS* motif in Smad2 may bind a specific site where two sulfate ions
were found within 4 Å of each other (10). The linear distance between
C
In our studies, we used the unphosphorylated Smad2 and Smad4 proteins.
Could this affect our final conclusion? We think that the answer is
likely to be no. The major role of phosphorylation is to relieve the
inhibitory effect of the MH1 domain and to release R-Smads from SARA
and other proteins (14, 17). With the removal of the MH1 domain, the
resulting Smad proteins are fully able to form heterocomplex between
R-Smad and Co-Smad and are constitutively active in transcriptional
assays (14-16). In fact, the MH2 domain of Smad2 in Xenopus
is constitutively localized to the nucleus, and its
developmental phenotype closely resembles that of the full-length Smad2
upon activin signaling (23). In addition, phosphorylation occurs in the
C-terminal end of R-Smads, which is flexible in solution and disordered
in all crystal structures (9, 10, 19). Thus, phosphorylation is very
unlikely to alter the stoichiometry of a stable heterocomplex. It has
been reported that substitution of the C-terminal Ser residues in
R-Smad by Glu resulted in increased binding affinities between R-Smad and Co-Smad (24), presumably because the carboxylate side chain in Glu
residues mimics the phosphate group. To assess the effect of such
substitutions, we created a mutant Smad2 with the two C-terminal Ser
residues replaced by Glu and purified this protein to homogeneity.
Using gel filtration (Fig. 5B) and electrophoretic mobility
shift assays (data not shown), this mutant Smad2 (SEME) is shown to
form a stable heterodimer with Smad4 with a modest increase in binding
affinity (Table II). Using this mutant Smad2, we also repeated all of
the experiments reported in this study. In all cases, the results are
identical to those for the wild-type Smad2. Despite this agreement, we
caution against interpretation of results by using this mutant to
substitute for the fully phosphorylated Smad2 protein. The reason is
clear: a phosphorylated Ser is stereochemically different from the Glu
residue, and the phosphorylated SS*MS* motif in Smad may show stringent
specificity for binding.
On the basis of our biochemical and biophysical analysis, Smad2 clearly
forms a heterodimer with Smad4. There are three scenarios for a
heterodimer. Two scenarios involve packing interactions similar to that
of the homotrimer, except that the relative positions of Smad2 and
Smad4 could be switched (Fig. 5C); a third scenario involves
a novel interaction interface, possibly psuedosymmetric. Our current
data support the third scenario because the results with mutations
D450H in Smad2 (D537H in Smad4) and D351H (D300H in Smad2) and R361H
(R310H in Smad2) in Smad4 are not compatible with either of the first
two scenarios.
Smad2 versus Smad3--
Given the strong sequence similarity among
R-Smads, it appears likely that Smad4 forms a heterodimer with other
R-Smads. For example, Smad2 shares 92% sequence identity with Smad3,
and both proteins are involved in signaling by TGF-
In summary, we conclude that the R-Smad Smad2 forms a stable
heterodimer with Co-Smad Smad4. The formation of this complex requires
the SAD domain in Smad4 and can be disrupted by a number of
tumor-derived missense mutations in both Smad2 and Smad4. This finding
should have broad implications in the interpretations of a range of
biological experiments.
signaling from the cell membrane to the nucleus. Upon
phosphorylation by the activated receptor kinases, the
receptor-regulated Smad, such as Smad2, forms a heterocomplex with the
co-mediator Smad, Smad4. This heterocomplex is then translocated into
the nucleus, where it associates with other transcription factors and
regulates expression of ligand-responsive genes. The stoichiometry
between receptor-regulated Smad and co-mediator Smad is important for
understanding the molecular mechanisms of the signaling process. Using
purified recombinant proteins, we demonstrate that Smad2 and Smad4 form
a stable heterodimer and that the Smad4 activation domain is important
for the formation of this complex. Many tumor-derived missense
mutations disrupt the formation of this heterocomplex in in
vitro interaction assays. Mapping these mutations onto the
structures of Smad4 and Smad2 identifies a symmetric interface between
these two Smad proteins. Importantly, two previous models on the
formation of a heterocomplex are incompatible with our
observations and other reported evidence.
INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS AND DISCUSSION
REFERENCES
1 signaling
regulates a broad range of cellular responses, including growth,
differentiation, and cell fate specification, in all animals (1, 2).
TGF-
signaling from the cell membrane to the nucleus is mediated by
the Smad family of proteins, which contains at least nine distinct
members in vertebrates and two of which, Smad2 and Smad4, have been
identified as tumor suppressors in humans (3-6).
) in Xenopus, which participate in
signaling by diverse TGF-
family members, (ii) receptor-regulated Smads (R-Smads), including Smad1, Smad2, Smad3, Smad5, and Smad8, each
of which is involved in a specific signaling pathway, and (iii)
inhibitory Smads, which include Smad6 and Smad7 and negatively regulate
these pathways (3-5, 7).
response is initiated by the binding of a specific TGF-
ligand to a pair of specific transmembrane receptors, the type I and II
receptors, leading to the activation of the Ser/Thr kinase in the
cytoplasmic domain of the type I receptor (8). The signal is then
propagated by the type I receptor-mediated phosphorylation of specific
R-Smads. For example, Smad1, Smad5, and Smad8 are phosphorylated by the
bone morphogenetic protein receptors, whereas Smad2 and Smad3
are phosphorylated by the activin and TGF-
receptors. The
phosphorylated R-Smad hetero-oligomerizes with Co-Smad Smad4,
translocates into the nucleus, and associates with sequence-specific
DNA-binding protein(s), resulting in the positive or negative
regulation of ligand-responsive genes.
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS AND DISCUSSION
REFERENCES
RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS AND DISCUSSION
REFERENCES
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Fig. 1.
Formation of a stable heterodimer between
Smad2 and Smad4. The Smad2 and Smad4 fragments contained residues
241-467 and 251-552, respectively. A, size exclusion
chromatography of Smad2 (250 µg; panel 1;
panels in A are numbered from top
to bottom), Smad4 (330 µg; panel 2), a
1:1 complex of Smad2-Smad4 (250 + 330 µg; panel 3), a
1.5:1 complex of Smad2-Smad4 (370 + 330 µg; panel 4), and
a concentrated 1:1 complex of Smad2-Smad4 (total input 2.5 mg;
panel 5). Relevant fractions were visualized on
SDS-polyacrylamide gel electrophoresis. The chromatographs and the
calibration of the Superdex-200 column are shown on the
right. The arrows indicate the starting points
for all four chromatographic runs. B, electrophoretic
mobility shift assays under nondenaturing conditions. C,
mutual exclusion of a Smad2-Smad4 complex and a Smad2-SARA complex.
GST-SARA was first bound to glutathione resin, and a stoichiometric
amount of a 1:1 Smad2-Smad4 complex was allowed to flow through the
resin. The resin was washed four times with assay buffer, and aliquots
of the last wash were visualized on SDS-polyacrylamide gel
electrophoresis.
Analytical ultracentrifugation analysis of a Smad2-Smad4 complex
-
heterotrimer (where both 2:1 or 1:2 ratios of
Smad2:Smad4 were considered) and both heterodimer and heterotrimer
self-association (1
-
N where 1 equals either the
heterodimer or heterotrimer unit and N = 2, 3, 4, 5, and 6 units of each heteromer). Least squares analysis disfavors all
models involving heterotrimer formation. The best two-state model is a
self-associating model for the heterodimer with N = 4.
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Fig. 2.
Smad2 and Smad4 form heterodimers by
ultracentrifugation analysis. Sedimentation equilibrium data and
fit derived from a one-state model where the molecular mass is fixed to
58,915 daltons, that of a heterodimer.
-strand that packs against a hydrophobic
surface (19). On the other hand, neither the MH2 domain of Smad4
(residues 319-552) (Fig. 3B, upper panel) nor a longer
fragment with 19 additional amino acids (300) was able to form a
stable heterocomplex with Smad2, as judged by their progressive
propensity toward dissociation upon size exclusion chromatography.
Inclusion of the full Smad4 activation domain (SAD) (20) restored its
ability to form a stable heterodimer with Smad2 (Fig. 3B, bottom
panel). The coupling of efficient formation of a heterocomplex
with the requirement for SAD may have functional implications for
TGF-
signaling.
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Fig. 3.
The SAD (residues 275-322) in Smad4 is
required for the formation of a stable heterodimer with Smad2.
A, a summary of the results obtained with Smad2 and Smad4
deletion mutants. The Smad4 mutants were assayed for their interaction
with Smad2 (residues 241-467), whereas the Smad2 mutants were examined
using Smad4 (residues 251-552). Gel filtration and GST-mediated
pull-down assays yielded consistent results. B, two
representative results showing the requirement for SAD in Smad4 for
complex formation with Smad2. In the top panel, equimolar
amounts of Smad4 (319) and Smad2 (182) were analyzed on gel
filtration after a 45-min preincubation. They dissociate during the
course of gel filtration. In the bottom panel, equimolar
amounts of the SAD-containing Smad4 (271) and Smad2 (241)
were co-eluted as a stable heterodimer.
Mutational analysis of a Smad2-Smad4 complex
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Fig. 4.
Tumorigenic mutations inactivate the
heterodimer between Smad2 and Smad4. Ten tumor-derived missense
mutations were introduced in Smad4 (251) and Smad2
(241). These mutant proteins were purified to homogeneity and
assayed for their ability to interact with their wild-type counterparts
by gel filtration. A, representative results showing that
three Smad4 mutants (indicated by a purple line) are unable
to form a heterocomplex with Smad2, whereas one Smad4 mutant and one
Smad2 mutant (highlighted by an orange line) retain their
ability to form a heterodimer. B, schematic representation
of the residues affected by the missense mutations. The MH2 domain of
Smad4 is shown in blue. Mutation of the purple
residues results in disruption of heterodimer formation. Mutation of
the orange residues does not affect the formation of a
heterodimer. Mutations in parentheses are introduced in
Smad2. The two corresponding mutations in Smad2 and Smad4 exhibit an
identical effect on the formation of a heterocomplex.
of residue 545 in Smad4 and the closest sulfate ion of the two is
over 32 Å, more than the head-to-tail distance of 8 residues, even if
they are in their most extended conformations. Thus, in order to reach this site, a minimum of 8 residues is required between the Smad2 residue that corresponds to residue 545 in Smad4 and the first phosphorylated Ser residue in Smad2; however, there are only 6 residues
in between them (9, 10). Finally, perhaps most importantly, if there
were a heterotrimer between Smad2 and Smad4, then we should have
observed this heterotrimer by either gel filtration or analytical
ultracentrifugation because the heterotrimer formation should be
heavily favored over an unstable heterodimer due to the cooperativity
involved. However, we did not obtain any evidence supporting the
existence of a heterotrimer.
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Fig. 5.
Comparison of proposed models of
heterocomplex between Co-Smad and R-Smad. In A and
C, Co-Smad and R-Smad are colored green and
blue, respectively. The critical interface residues from
Co-Smad and R-Smad are shown in purple and
orange, respectively. The SAD fragment is shown in
red. A, proposed heterotrimer model. This model
is inconsistent with several experimental observations (see "Results
and Discussion"). This model shows a 2:1 complex between Co-Smad and
R-Smad. The discussion in the text also applies to the other scenario,
in which Co-Smad and R-Smad form a 1:2 complex. B, formation
of a stable heterodimer between Smad4 (251) and a mutant Smad2
(241) in which the two C-terminal Ser residues (465 and 467) are
replaced by Glu (SEME). Equimolar amounts of these two
proteins were incubated together for 45 min before assay by size
exclusion chromatography. C, two possible models of a
heterodimer. Neither is consistent with our mutational analysis.
and activin.
Nevertheless, the basal states of Smad2 and Smad3 differ considerably
(11). Thus, the states of their heterocomplexes with Smad4 may also be
different, as is the case for their biological functions. For example,
Smad3 exhibits the highest sequence-specific DNA binding affinity; but
Smad2 does not bind DNA because of an obstructing insertion immediately
before the DNA-binding
-hairpin (25). Ectopic expression of the
ubiquitin E3 ligase, Smurf2, selectively reduces the
steady-state levels of Smad2 but not Smad3 (26). More importantly,
Smad3-null mice are viable, but Smad2-null mice are not. Thus, it
remains to be seen how Smad3 or Smad1 forms a heterocomplex with Smad4.
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ACKNOWLEDGEMENTS |
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We thank F. Hughson for critical reading of the manuscript and S. Kyin for peptide sequencing, DNA synthesis, and mass spectroscopy.
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FOOTNOTES |
---|
* This research was supported by National Institutes of Health Grant R01-CA82171.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.
¶ A Searle Scholar and a Rita Allen Scholar.
To whom correspondence should be addressed: Dept. of
Molecular Biology, Princeton University, Washington Road, Princeton, NJ
08544 Tel.: 609-258-6071; Fax: 609-258-6730; E-mail:
yshi@molbio.princeton.edu.
Published, JBC Papers in Press, March 27, 2001, DOI 10.1074/jbc.M100174200
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ABBREVIATIONS |
---|
The abbreviations used are:
TGF-, transforming growth factor
;
R-Smad, receptor-regulated Smad;
Co-Smad, co-mediator Smad;
SAD, Smad4 activation domain;
GST, glutathione S-transferase.
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