Identification and Characterization of Constitutively Active Smad2 Mutants: Evaluation of Formation of Smad Complex and Subcellular Distribution
Masayuki Funaba and
Lawrence S. Mathews
Department of Biological Chemistry University of Michigan
Ann Arbor, Michigan 48109-0606
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ABSTRACT
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Smads mediate activin, transforming growth
factor ß (TGFß), and bone morphogenetic protein signaling from
receptors to nuclei. According to the current model, activated
activin/TGFß receptors phosphorylate the carboxyl-terminal serines of
Smad2 and Smad3 (SSMS-COOH); phosphorylated Smad2/3 oligomerizes with
Smad4, translocates to the nucleus, and modulates transcription of
defined genes. To test key features of this model in detail, we
explored the construction of constitutively active Smad2 mutants.
To mimic phosphorylated Smad2, we made two Smad2 mutants with acidic
amino acid substitutions of carboxyl-terminal serines: Smad22E
(Ser465, 467Glu) and Smad23E (Ser464, 465, 467Glu). The mutants
enhanced basal transcriptional activity in a mink lung epithelial
cell line, L17. In a Smad4-deficient cell line, SW480.7, Smad22E did
not affect basal signaling; however, cotransfection with full-length
Smad4, but not transfection of Smad4 alone, resulted in
enhanced basal transcriptional activity, suggesting that the
constitutively active Smad2 mutant also requires Smad4 for function.
In vitro protein interaction analysis revealed that
Smad22E bound more tightly to Smad4 than did wild-type Smad2;
dissociation constants were 270 ± 66 nM
for wild-type Smad2:Smad4 complexes and 79 ± 18
nM for Smad22E:Smad4 complexes.
Determination of the subcellular localization of Smad2
revealed that a greater percentage of Smad22E was localized in the
nucleus than wild-type Smad2. These results suggest that Smad2
phosphorylation results in both tighter binding to Smad4 and increased
nuclear concentration; those changes may be responsible for
transcriptional activation by Smad2.
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INTRODUCTION
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Members of the transforming growth factor ß (TGFß)
family, including activins, TGFßs, and bone morphogenetic
proteins (BMPs), are protein growth and differentiation factors that
manifest an extraordinary variety of biological activities (1). These
proteins signal through the sequential activation of two cell surface
receptors, termed type I and type II, both of which are protein
serine-threonine kinases (2, 3). Extensive efforts to elucidate the
downstream signaling mechanisms led to the discovery of a series of
Smad proteins and trials for clarification of Smad signaling
mechanisms.
Smads are categorized into three subclasses: receptor-regulated Smads,
which include Smad2 and 3 (activin/TGFß receptor regulated)
and Smad1, 5, and 8 (BMP receptor regulated); the common-partner
Smad (Smad4); and the inhibitory Smads (Smad6 and 7) (2, 3, 4).
Receptor complexes activated by appropriate ligands phosphorylate
conserved serine residues at the carboxyl terminus of
receptor-regulated Smads (5, 6, 7), which promotes association with a
common partner Smad, Smad4 (8, 9, 10). The complexes of
receptor-regulated Smad and Smad4 translocate into the nucleus and
participate in the activation of target gene transcription either by
associating with other transcription factors or by direct DNA binding
(24, 11).
Recently, on the basis of this phosphorylationrelated Smad
activation model, detailed studies on Smad signaling mechanism have
been conducted. As for receptor-regulated phosphorylation, SARA (for
Smad anchor for receptor activation), identified as a Smad2 binding
protein, was suggested to facilitate Smad phosphorylation by
localization of unphosphorylated Smad in proximity to the activated
receptor complexes (12). In addition, several nuclear proteins,
including p300, the AP-1 complex, TFE3, vitamin D receptor, Evi-1,
c-Ski, and SnoN, have been found to interact with Smads and influence
Smad function (13, 14, 15, 16, 17, 18, 19, 20, 21).
The model for phosphorylation-related Smad activation and function is
broadly accepted (2, 3, 4), but significant points remain to be verified;
for example, it is unclear whether phosphorylation directly influences
Smad2 binding to Smad4, or subsequent signaling events. In addition,
the current model derives primarily from qualitative data. To explore
the quantitative, biochemical basis for this model, in this study we
assessed the mechanistic implications of Smad2 phosphorylation. Because
it is not currently possible to generate large amounts of
stoichiometrically phosphorylated, purified Smad2, we demonstrated that
replacement of the phosphorylated serines with acidic amino acids
yielded a constitutively active form of Smad2. We have used that mutant
to investigate quantitatively the biochemical function of Smad2
phosphorylation.
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RESULTS
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Identification of Constitutively Active Mutants of Smad2
To identify constitutively active Smad2 mutants, we replaced the
carboxyl-terminal serines of Smad2 (SSMS-COOH) with glutamic acid to
mimic phosphorylation. These serines were previously identified as
ligand-induced phosphorylation sites (5, 6, 7); in particular, the last
two serine residues of Smad2 were phosphorylated in response to TGFß
stimulation (22, 23). To evaluate transcriptional activities of
Smad22E (Ser465, 467Glu) and -3E (Ser464, 465, 467Glu) in L17 cells,
we used three different luciferase-based reporter genes: 3TP-Lux, which
contains regulatory elements from the PAI-1 promoter (24); AR3-Lux,
which contains regulatory sequences from the Xenopus Mix.2
gene, and which requires coexpression of the transcription factor FAST1
for ligand induction (25); and SBE4-Lux, which contains optimized
Smad-binding regulatory sequences (26).
Neither wild-type Smad2 (WT-Smad2) nor the mutants affected
basal expression of 3TP-Lux and SBE4-Lux (Fig. 1A
). In addition, cotransfection of Smad4
also had no effect. On the contrary, WT-Smad2 resulted in a 2-fold
increase in basal expression of AR3-Lux, and cotransfection of Smad4
resulted in a further increase in AR3-Lux expression (Fig. 1A
). This
induction was dependent on FAST1 expression (data not shown). Smad22E
and Smad23E elevated basal expression of AR3-Lux 21-fold and
13-fold, respectively. Treatment with activin in the presence of either
wild-type or mutant Smad2 yielded additional AR3-Lux expression (Fig. 1B
). The increased basal expression in the presence of the acidic
mutants was approximately the same as the amount of activin-induced
expression in the presence of WT-Smad2. The absolute
activin-induced increase in all cases except for WT-Smad2 was
approximately the same, consistent with the activation of endogenous
Smads. We also examined other possible phosphorylation-mimic mutants;
432Thr-Ser-Thr, located at the L3 loop of the MH2
domain, were replaced with three glutamic acid residues, because the L3
loop was solvent exposed and protruded from the ß-sandwich core
structure (27) and involved in Smad-receptor interaction (28). However,
none of the reporter genes responded to this mutant (data not
shown).

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Figure 1. Smad22E and -3E Are Constitutively Active Mutants
in the Expression of AR3-Lux in L17 Cells
A, Effect of Smad2 mutants on 3TP-Lux, FAST1-dependent AR3-Lux, and
SBE4-Lux transcriptional responses. L17 cells were transiently
transfected with a reporter (3TP-Lux, AR3-Lux or SBE4-Lux),
ß-galactosidase, FAST1 for AR3-Lux, and wild-type (WT) or mutant
versions of Smad2 together with (hatched bar) or without
(solid bar) Smad4. Luciferase activity was normalized to
ß-galactosidase activity, and luciferase activity in the cell
lysates in the absence of exogenous Smads and FAST1 for each reporter
was set to 1. Data were expressed as the mean ±
SD of triplicates from a representative
experiment. B, Responses to activin by Smad2 mutants on expression of
AR3-Lux. L17 cells were transiently transfected with the AR3-Lux
reporter, ß-galactosidase, FAST1, Smad4, and WT or mutant versions of
Smad2 as indicated. Cells were treated with 2 nM
activin A for 16 h, and luciferase activity was measured in cell
lysates. Luciferase activity was normalized to ß-galactosidase
activity, and luciferase activity in the cell lysates in the absence of
exogenous Smad2 and FAST1 was set to 1. Data were expressed as the
mean ± SD of triplicates from a
representative experiment.
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Smad4 Requirement for Constitutive Smad2 Activity
To determine whether the constitutively active mutants of
Smad2 needed Smad4 to manifest their activity, we conducted
transcriptional activation assays in Smad4-deficient SW480.7 cells (29, 30). AR3-Lux expression did not respond to activin stimulation (data
not shown), although the reason is not clear. Therefore, we used TGFß
as a ligand, because TGFß as well as activin could activate this
reporter gene transcription (9, 27). In the absence of Smad4, neither
WT-Smad2 nor Smad22E transfection affected AR3-Lux expression (Fig. 2
). Expression of full-length Smad4
increased AR3 expression in response to TGFß stimulation by WT-Smad2
transfection. In the presence of Smad4, Smad22E transfection caused
enhanced basal expression (Fig. 2
). In contrast, a Smad4
construct with a small carboxyl-terminal truncation,
i.e. deletion of amino acids 516552 [Smad4(1515)]
failed to increase either basal or TGFß-induced luciferase activity
in the presence of either WT-Smad2 or Smad22E. These findings suggest
that Smad4 is essential for Smad22E-induced AR3 transcription.

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Figure 2. Role of Smad4 for Smad2-Induced AR3-Lux
Transcription
Smad4-deficient SW480.7 cells were transiently transfected with the
AR3-Lux reporter, ß-galactosidase, FAST1, and WT-Smad2 or Smad22E
together with Smad4 expression constructs as indicated. After 24 h
of transfection, cells were treated with 100 pM
TGFß1 for 16 h, and luciferase activity was measured
in cell lysates. Luciferase activity was normalized to
ß-galactosidase activity, and luciferase activity in the cell lysates
in the absence of exogenous Smad2 and FAST1 was set to 1. Data were
expressed as the mean ± SD of triplicates from a
representative experiment.
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Binding of Smad2 to Smad4
The reporter assays described above revealed that Smad4 was
required for enhanced basal transcriptional activity of Smad22E.
Phosphorylated Smad2 is proposed to associate with Smad4 for
transcriptional activation (3, 11). To test that hypothesis, we
characterized the Smad2:Smad4 association by two assays, a
glutathione-S-transferase (GST)-pull down assay and
fluorescent analysis.
The GST-pull down assay revealed that more in vitro
translated 35S-Smad22E bound to GST-Smad4 than
did 35S-WT-Smad2 (Fig. 3B
). No Smad binding was detected after
incubation with GST beads alone. These results suggested that the
Smad22E:Smad4 interaction was higher affinity than the WT-Smad2:Smad4
interaction.

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Figure 3. Association of Smad2 with Smad4 by GST-Pull Down
Assay
Association of in vitro translated Smad2 and GST-Smad4.
A, GST or GST-Smad4 proteins were expressed in BL21 and
affinity-purified by glutathione-Sepharose 4B beads. B,
35S-Labeled WT-Smad2 or Smad22E protein was first
incubated with GST beads preadsorbed with BSA. The flow-through
fraction was then reacted with GST beads or GST-Smad4 beads that were
preadsorbed with BSA. After the beads were washed, the proteins were
resolved by SDS-PAGE, and the 35S-labeled bands were
detected by autoradiography.
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Association of Smad2 with Smad4 was quantified by fluorescent analysis
using dansylated Smad4 and purified Smad2. For this purpose, Smad2 and
Smad4 proteins expressed in Escherichia. coli were purified
to near homogeneity (Fig. 4A
). Maximum
fluorescence intensity of dansylated Smad4 increased in the presence of
Smad2 in a dose-dependent manner (Fig. 4
, B and C), suggesting the
detection of Smad2:Smad4 complexes. In contrast, no increase in the
fluorescence intensity was observed in the presence of an unrelated
protein, BSA (data not shown). When differences of maximum fluorescence
intensity between dansylated Smad4 in the presence of Smad2 and that in
the absence of Smad2 were plotted as a function of free Smad2
concentration, a one-site binding model provided the best fit for both
the WT-Smad2:Smad4 data and the Smad22E:Smad4 data. The average
Kd values of three independent trials were
270 ± 66 nM (mean ±
SE) for WT-Smad2:Smad4 and 79 ± 18
nM for Smad22E:Smad4; these
Kds were statistically different
(P < 0.05), indicating tighter binding of Smad4 to
Smad22E as compared with WT-Smad2.

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Figure 4. Association of Smad2 with Smad4 by Fluorescence
Analysis
A, Purified Smad2 and Smad4 proteins for fluorescence analysis.
Protein expressed in E. coli as GST fusion protein was
purified to near homogeneity. The proteins were loaded on a 10%
SDS-polyacrylamide gel and stained by Coomassie brilliant blue R-250.
B, Emission spectrum of dansylated Smad4 in the presence of purified
Smad2. Purified Smad4 was labeled with dansyl aziridine. Dansylated
Smad4 was mixed with purified Smad2. Excitation was performed at 340
nm, and emission was scanned from 430 nm to 570 nm. C, Titration curve
with Smad2 for differences of maximum fluorescence intensity between
dansylated Smad4 in the absence of Smad2 and that in the presence of
Smad2. To determine the dissociation constant of the Smad2:Smad4
complex, after calculating free Smad2 concentration, one-site binding
model, y = a x
x/(Kd + x), was applied. The
figure shows a plot of the fluorescence responses against total Smad2
concentration, and each point shows average and SE of
three independent experiments; to compare data among experiments,
y axis data were normalized to show the percentage of
the maximum response in each experiment.
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Subcellular Localization of Smad2
A number of qualitative immunocytochemical analyses have indicated
that receptor-regulated Smads accumulated in the nucleus in response to
TGFß or BMP stimulation (9, 31, 32, 33), suggesting that phosphorylation
of the serine residues at the carboxyl terminus caused nuclear
translocation. We examined the distribution of unactivated and
activated Smad2 in L17 cells by subcellular fractionation and
subsequent immunoblot analysis. Subcellular distribution of lactate
dehydrogenase (LDH) as a cytosolic marker and DNA as a nuclear
marker confirmed that cross-contamination of cytosol with nucleus was
minimal (Fig. 5
, A and C).

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Figure 5. Subcellular Distribution of Smad2 and Smad4 in L17
Cells
L17 cells were transiently transfected with HA-WT-Smad2 or HA-Smad22E
with or without Flag-Smad4 expression constructs. After 48 h of
transfection, cells were treated with or without 2 nM
activin A for 45 min. Cell suspensions were homogenized in 0.25
M sucrose buffer and cytosol was separated from nuclei
by centrifugation. A and C, Cross-contamination of each fraction. LDH
activity was measured as a cytosol marker, whereas DNA was measured as
a nuclear marker. B and D, Twenty-microgram proteins were subjected to
Western blot using anti-HA or anti-Flag antibody to detect expressed
Smad2 or Smad4.
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The amount of WT-Smad2 was higher in cytosol than in nucleus (Fig. 5B
).
The amount of Smad22E in cytosol was also higher than in nucleus;
however, there was relatively more Smad22E in the nuclear fraction
than WT-Smad2. This effect was independent of exogenous Smad4
expression (Fig. 5B
). A tendency of higher nuclear localization of
Smad22E was also detected by immunocytochemical analyses (data
not shown). Unlike Smad2, exogenously expressed Smad4 was evenly
distributed within the cell, and the distribution was not changed by
Smad22E expression (Fig. 5B
). Activin stimulation had quite minimal
effect on Smad2 subcellular distribution both in cells transfected with
WT-Smad2 and in cells transfected with Smad22E (Fig. 5D
). No
remarkable changes in subcellular distribution of Smad2 were observed
by immunocytochemical analyses, although Smad2 lacking the
amino-terminal region [Smad2(264467)] was constitutively localized
in nucleus (data not shown), consistent with the previous results (34, 35).
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DISCUSSION
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In this study we showed that mimicking receptormediated
phosphorylation of Smad2, by mutation of the phosphorylated serines to
acidic amino acids, enhanced Smad2s basal transcriptional activity.
Using this constitutively activated Smad2 mutant (Smad22E), we
explored the molecular mechanisms linking phosphorylation to changes in
function. In vitro binding assays revealed that Smad22E
bound more tightly to Smad4 than did WT-Smad2. In addition, there was
relatively more Smad22E in the nucleus than WT-Smad2. Together, those
two effects likely account for the enhanced basal activity of
Smad22E, consistent with data from qualitative studies suggesting
that phosphorylation-induced Smad2 complex formation with Smad4 and
nuclear translocation are related to transcriptional activation (2, 3).
The reporter gene assays indicated that Smad4 was essential for
Smad2-mediated signaling (Fig. 2
). The findings were consistent with
the previous reports; Smad2 and Smad4 synergized on reporter gene
transcription (8, 9, 33), but Smad4 lacking the carboxyl terminus
activated neither 3TP-Lux (8) nor AR3 gene transcription (9). Because
full-length Smad4 but not the truncated Smad4 could form complexes with
Smad2 in response to agonist stimulation (36), the current concept that
Smad2:Smad4 complexes are the active signaling form has been proposed
(2, 3, 11). The tighter binding of Smad22E to Smad4, as determined by
in vitro binding assays, could thus at least partially
explain the elevated basal transcriptional activity. The
Kd for the Smad22E:Smad4 interaction (
80
nM) is in the same range as that determined for
other transcription factor pairs, including nuclear factor
B (NF
B)
p50-p50 homodimer [
1 µM (37)] and Jun:Fos
heterodimers [23110 nM (38, 39)]. A change in
affinity due to Smad2 phosphorylation could thus conceivably alter the
amount of active complex.
A greater percentage of Smad22E was localized in the nucleus than
WT-Smad2, and exogenous Smad4 expression had no effect on the
subcellular distribution. These results were consistent with the
immunocytochemical observation on nuclear translocation of
receptor-regulated Smads by agonist stimulation even in Smad4-deficient
cells (9). Because Smad4 was necessary for Smad22E-mediated AR3
transcription, nuclear translocation of Smad2 would not be sufficient
for the signaling.
Currently, intracellular signaling of activin is indistinguishable from
that of TGFß; Smad2 was phosphorylated in response to stimulation
by activin as well as TGFß with a peak at 60 min after
ligand addition (8, 35), a time at which strongest formation
of Smad2:Smad4 complexes was also seen (40). However, nuclear
translocation of Smad2 by activin stimulation was less evident than
that of Smad3 by activin stimulation or that of Smad2 by TGFß
stimulation (41). Although our results that activin had quite minimal
effect on subcellular distribution of Smad2 might result from transient
overexpression of Smad2, there might be additional activin-regulated
steps for Smad2-mediated signaling independent of subcellular
distribution.
Mutation of the carboxyl-terminal serines of Smad2 to glutamate
resulted in elevated basal expression of AR3-Lux; however, no effect
was observed for two other reporter genes, 3TP-Lux and SBE4-Lux. There
are several possible explanations for that finding. 1) Smad2 may
participate in transcriptional activation of AR3-Lux, but not in the
activation of the other promoters. The activin-responsive factor that
bound the Mix.2 promoter element in response to activin
stimulation in Xenopus embryos (42) contained Smad2, and
Smad2 was essential for formation of the complex (43, 44). In contrast,
Smad2 did not bind DNA elements from either 3TP-Lux (45) or SBE4-Lux
(26). Furthermore, Smad2 had little effect on basal expression of
either of those reporter genes (5, 22, 23, 33, 45, 46). 2)
Transcription of 3TP-Lux and SBE4-Lux could require additional
ligand-induced events, as yet undefined, whereas for AR3-Lux
transcription Smad2 carboxyl-terminal phosphorylation could be
sufficient. 3) The nuclear concentration of Smad2 required for
transcriptional activation of AR3-Lux may be lower than that needed for
3TP-Lux and SBE4-Lux. The EC50 for activin
induction of AR3-Lux is lower than that for the other reporter genes
(M. Bayram and L. S. Mathews, unpublished), suggesting that
AR3-Lux is a more sensitive reporter for Smad-mediated signals.
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MATERIALS AND METHODS
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cDNA Constructs
The following plasmids were provided: 3TP-Lux (24) by Dr. J.
Massagué, SBE4-Lux (26) by Dr. B. Vogelstein, AR3-Lux (25) and
Xenopus FAST1 cDNA (43) by Dr. M. Whitman, carboxyl-terminal
Flag-tagged human Smad cDNAs (30) by Dr. R. Derynck, Xenopus
Smad2 cDNA (47) by Dr. J. M. Graff, HA-pcDNA3 and Flag-pcDNA3
expression vector (48) by Drs. N. Inohara and T. Koseki, human Smad4
subcloned into the vector pGEX-KG using BamHI and
EcoRI sites by E. Tang and Dr. K.-L. Guan.
The human Smad2 mutants for expression vectors were constructed by a
one-step PCR method. The human Smad2 and human Smad4 cDNAs were
subcloned into the EcoRI and XbaI sites of
hemagglutinin (HA)-pcDNA3 to produce N-terminal HA-tagged proteins. For
GST-Smad2 protein expression, Smad2 cDNA was subcloned into the vector
pGEX-2T using SmaI and EcoRI sites.
Cell Culture and cDNA Transfection
The L17 cells, a derivative of the mink lung epithelial cell
line (Mv1Lu) provided by Dr. J. Massagué, were cultured and
transfected as described previously (49). SW480.7 colon carcinoma cell
line was provided by Dr. E. J. Stanbridge (29). The cells were
cultured in DMEM with 10% FBS, 100 U/ml penicillin, and 100 µg/ml
streptomycin. For transient transfection, cells in 24-well plates or in
10-cm cell culture dishes were transfected by diethylaminoethyl-dextran
method.
Reporter Assay
Luciferase assays were basically conducted as described
previously (50). L17 cells and SW480.7 cells were transiently
transfected with various Smad constructs and FAST1 together with a
reporter construct (3TP-Lux, AR3-Lux or SBE4-Lux) and a plasmid
expressing ß-galactosidase (pCMV-ßGal). In each experiment, equal
amounts of DNA were transfected, which was achieved by adjusting empty
vector (pcDNA1 and pcDNA3). For basal induction experiments, L17 cells
were harvested at 40 h after transfection. For ligand stimulation
experiments, at 24 h after transfection, cells were treated with 2
nM activin A (provided by Dr. T. K. Woodruff and by
the NIH Hormone Distribution Program, NIDDK) or 100 pM
TGFß1 (Becton Dickinson and Co.,
Franklin Lakes, NJ) for 16 h. Luciferase activity was normalized
to ß-galactosidase activity, and luciferase activity in the cell
lysates in the absence of exogenous Smads, FAST1, and ligand was set to
1.
GST-Pull Down Assay
The GST and GST-fused Smad4 proteins were expressed in E.
coli and purified by use of glutathione-Sepharose beads
(Amersham Pharmacia Biotech. Arlington Heights, IL)
according to the manufacturers protocol. Purified GST or GST-Smad4
bound to glutathione-Sepharose beads was preadsorbed with 0.5 mg/ml
BSA, 1 mM EDTA to prevent nonspecific binding
with beads for 3 h at 4 C. 35S-labeled
WT-Smad2 or Smad22E proteins, which were translated in
vitro by use of the TNT rabbit reticulocyte lysate kit
(Promega Corp., Madison, WI), were then loaded on a
GST-Sepharose column preequilibrated with binding buffer [50
mM Tris-HCl (pH 7.4), 120
mM NaCl, 2 mM EDTA, 0.1%
NP-40, 1 mM phenylmethylsulfonyl fluoride
(PMSF)]) for 1 h at 4 C. The flow-through was loaded on a
glutathione-Sepharose column bound to 3 µg of GST or GST-Smad4 and
incubated for 1.5 h at 4 C. The columns were washed four times
with binding buffer. Specifically bound proteins dissolved in SDS-PAGE
sample buffer were separated by SDS-PAGE (10% gel) and visualized by
fluorography.
Fluorescence Analysis
For detection of Smad binding by fluorescence analyses, Smad
proteins were highly purified from E. coli expressed as
GST-fused proteins (90% < purity). The purified Smad proteins had
appropriate in vitro biological activities; Smad2 was
phosphorylated by TGFß receptor complexes and bound to calmodulin in
a calcium-dependent manner, and Smad4 protein bound to SBE4 (M. Funaba
and L. S. Mathews, in preparation). A preliminary trial showed
that Xenopus Smad2 expression construct yielded more protein
than human Smad2 expression construct. Therefore, we
expressed Xenopus Smad2 protein. Because of high
homology of human Smad2 with Xenopus Smad2 (98% identity),
these two proteins are expected to behave similarly on binding with
human Smad4.
The purified Smad4 protein (8 µM) was labeled with 1 µl
of 200 mM dansyl aziridine (Molecular Probes, Inc., Eugene, OR) at the thiol sites for 2 h at room
temperature. One microliter of 1 M DTT was then added to
consume excess thiol-reactive reagent. The conjugate and free
thiol-reactive reagents were separated by use of a Sephadex G-25 spin
column (Roche Molecular Biochemicals,
Indianapolis, IN). Fluorescence measurements were performed
by using 40 nM dansylated Smad4 in 20 mM HEPES
(pH 7.5), 200 mM NaCl, 1 mM
CaCl2, and 1 mM DTT in the presence
of various concentrations of Smad2. The measurements were taken with an
FluoreMax-2 (Instruments SA Inc., Edison, NJ) with excitation
wavelength set at 340 nm and band width of 10 nm for both excitation
and emission wavelengths. Emission scans were recorded between 430 and
570 nm and fluorescent intensity of maximum emission was measured. To
determine the dissociation constant (Kd),
difference of maximum intensity at each concentration of Smad2 from
that of Smad4 without Smad2 was plotted against the free Smad2
concentration. A one-site binding model was applied (y
= a x x/(Kd + x)
where x is the concentration of free Smad2 and y
is the difference between the fluorescence intensity in the presence
and absence of Smad2) and the Kd was calculated
by use of Graph Pad PRISM (GraphPad Software, Inc., San
Diego, CA). Free Smad2 concentration was estimated from the following
equation: free Smad2 (nM) = total Smad2
(nM) - F/F
x 40
where F and F
are the difference of the
fluorescence intensity at the Smad2 concentration and at the highest
Smad2 concentration, respectively. Binding experiments were conducted
three times using three different lots of proteins. The comparison of
Kd values between WT-Smad2:Smad4 and
Smad22E:Smad4 was evaluated by Students t test, and
differences of P < 0.05 were considered statistically
significant.
Subcellular Fractionation
L17 cells were transiently transfected with HA-Smad2 and
Smad4-Flag expression constructs. For activin stimulation experiments,
at 48 h after transfection, cells were treated with 2
nM activin A for 45 min. The entire subcellular
fractionation procedure was performed at 4 C. Cells were washed with
HEPES dissociation buffer three times and suspended in isotonic buffer
[20 mM Tris-HCl (pH 7.4), 0.25 M sucrose, 1
mM EDTA, 1 mM
Na3VO4, 1 mM
PMSF, 1% aprotinin], followed by homogenization with a Dounce
homogenizer (25 strokes) and a Potter-Elvehjem homogenizer (15
strokes). After centrifugation at 900 x g for 7 min,
the supernatant was further centrifuged at 100,000 x g
for 1 h. The supernatant was referred to as a cytosolic fraction.
The pellet after the first centrifugation was washed with isotonic
buffer and resuspended in isotonic buffer. After ultrasonication, the
resuspension was centrifuged at 100,000 x g for 30
min. The supernatant was referred to as a nuclear fraction. Protein
concentrations from each fraction were measured by the bicinchoninic
acid method (51) after concentration by the sodium
deoxycholate-trichloroacetic acid method (52). Protein contents
in cytosolic fractions and in nuclear fractions were comparable among
cell treatments.
Five percent of the protein in each fraction was subjected to Western
blot analyses for examining expression of Smad2 and Smad4. To check
cross-contamination during homogenization, LDH activity as a
cytosolic marker and DNA content as a nuclear marker were measured by
the methods of Storrie and Madden (53) and Labarca and Paigen (54),
respectively.
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ACKNOWLEDGMENTS
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We thank Dr. Teresa Woodruff and NIH Hormone Distribution
Program, NIDDK, for providing activin A, and Drs. Rik Derynck, Jon
Graff, KunLiang Guan, Naohiro Inohara, Takeyoshi Koseki, Joan
Massagué, Eric Stanbridge, Eric Tang, Bert Vogelstein, and
Malcolm Whitman for providing plasmids and cell line. We also thank
Cole Zimmerman and Taju Kariapper for stimulating discussion and Akira
Abe for suggestions for fluorescence analysis.
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FOOTNOTES
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Address requests for reprints to: Masayuki Funaba, Azabu University School of Veterinary Medicine, 117-71 Fuchinobe, Sagamihara 229-8501, Japan. E-mail: funaba{at}azabu-u.ac.jp
This work was supported in part by American Cancer Society Grant
RPG-98352-01-TBE (to L.S.M.).
Received for publication February 8, 2000.
Revision received June 16, 2000.
Accepted for publication July 11, 2000.
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