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Departments of Biochemistry, Molecular Biology, and Cell Biology (W.C., K.E.M.) and Neurobiology and Physiology (T.K.W., K.E.M.), Center for Reproductive Science, Northwestern University, Evanston, Illinois 60208
Address all correspondence and requests for reprints to: Dr. Kelly E. Mayo, Department of Biochemistry, Molecular Biology, and Cell Biology, 2153 North Campus Drive, Northwestern University, Evanston, Illinois 60208. E-mail: k-mayo{at}nwu.edu
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Activins are formed by the combinatorial assembly of two closely
related subunits, ßA and
ßB, generating three dimeric isoforms, activin
A (ßAßA), activin B
(ßBßB), and activin AB
(ßAßB). Although
originally identified as a gonadal peptide hormone capable of
stimulating the release of FSH from the pituitary, activin has a wide
variety of biological functions, including the regulation of cell
proliferation and differentiation, and induction of mesoderm tissues in
amphibian development (7). Several lines of evidence suggest an
important role for activin in the liver. Injection of
[125I]activin A into rats has identified the
liver as a major target of activin binding or clearance (8). In inhibin
-subunit-deficient mice, activin levels are elevated, leading
to a severe cancer cachexia-like wasting syndrome that includes
hepatocellular death in the liver (9). Infusion of recombinant activin
A in mice and rats causes a marked reduction in liver mass resulting
from extensive cell death (6, 10). Activin also causes cell death in
primary hepatocyte cultures, and this effect can be blocked by
follistatin, an activin-binding protein, indicating that it is a
specific biological response to activin (6). The hepatocyte death
induced by activin exhibits characteristic nuclear and cytoplasmic
features of apoptosis (6). Similarly, activin has been shown to induce
apoptosis in other cell types, including mouse B cell hybridomas and
mouse and human myeloma cells (11, 12). The activin-related protein
TGFß also causes apoptosis in normal and transformed hepatocytes (13, 14). Transgenic mice overexpressing TGFß1 selectively develop hepatic
fibrosis and exhibit apoptosis of hepatocytes (15).
Members of the TGFß family of proteins, including TGFßs, activins and bone morphogenic proteins (BMPs), exert their biological actions through two types of cell surface receptors, designated type I and type II receptors, both of which are serine/threonine protein kinases (16, 17). Ligand binding to a type II receptor, which is a constitutively active kinase, recruits a type I receptor into the complex (18). The type I receptor is phosphorylated and activated by the type II receptor and propagates the signal to downstream proteins. Two type II receptors, ActRII (19) and ActRIIB (20), bind activin with high affinity and selectivity. The type I receptor ActRIB (also known as ALK-4) (21), selectively mediates activin signaling (22, 23). An additional type I receptor, ActRI (also known as ALK-2), associates with activin type II receptors to form heteromeric complexes after activin binding to the type II receptors (24, 25). However, ActRI has recently been shown to coimmunoprecipitate with a BMP type II receptor (26) and mediate the effects of BMP (27), and thus may be a common type I receptor shared by the activin and BMP signaling pathways.
The Smad family of proteins plays key roles in transducing signals from cell surface serine/threonine kinase receptors for TGFß superfamily proteins to nuclear target genes (28, 29, 30). To date, eight vertebrate Smad proteins have been identified, and these can be grouped into three classes. The first class is the receptor-regulated Smads. Among them, Smad1, Smad5, and Smad8 mediate BMP signaling, whereas Smad2 and Smad3 transduce activin and TGFß signals (28). These receptor-regulated Smads transiently interact with and become phosphorylated by specific activated type I receptors (31, 32, 33). Once phosphorylated, they associate with a common partner, Smad4 (also known as DPC4), which is the only known member of the second class of Smad proteins in vertebrates (34). The Smad complex then translocates to the nucleus and regulates transcription of target genes through interaction with specific DNA sequences and other DNA-binding proteins. For example, the Smad2/Smad4 complex has been shown to associate with the winged helix transcription factor FAST (35, 36, 37) and thereby interact with activin-responsive elements. Smad6 and Smad7 form the third class of Smad proteins, the inhibitory Smads. They act as inhibitors of BMP, activin, and TGFß signaling by competing with the receptor-regulated Smads for binding to the type I receptors or by competing with Smad4 for binding to the receptor-regulated Smads (38, 39, 40).
In the present study we observed that activin A induces apoptosis in a human hepatoma cell line, HepG2 cells. We use this model system to investigate the involvement of known activin signaling components, including the four activin receptors and two of the Smad proteins, in the activin A-induced apoptotic pathway. We demonstrate by blocking receptor action using a dominant negative approach that activin receptors are essential for mediating the apoptotic response. Furthermore, we show that overexpression of activin receptors and Smad proteins can mimic activin action and induce apoptosis in HepG2 cells. Our data suggest that these signaling molecules play a critical role in mediating apoptotic cell death induced by activin A in HepG2 cells.
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Materials and Methods |
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The cDNA clones for pCMV5-Flag-Smad2 and pCMV5-Flag-Smad4 were provided by Dr. Jeffrey Wrana (Hospital for Sick Children, Toronto, Canada) (32, 41). For luciferase assays, Flag-Smad2 and Flag-Smad4 were subcloned into the pcDNA3 vector. For fluorescence analysis, Smad2 and Smad4 were subcloned into the C-terminus of green fluorescence protein (GFP) in the pEGFP vector (CLONTECH Laboratories, Inc., Palo Alto, CA). To generate the Smad2 mutant (Smad2-DN), Smad2 was truncated 30 bp before the stop codon, deleting the last 10 amino acids, including the SSMS motif at the C-terminus of the protein.
Cell culture and transfection
HepG2 cells were maintained in DMEM (Sigma, St.
Louis, MO) supplemented with 2 mM sodium pyruvate and 10%
FCS (Life Technologies, Inc./BRL, Grand Island, NY) at 5%
CO2 in a 37 C incubator. For luciferase assays
and fluorescence microscopy, cells were transfected with cationic
liposomes prepared as previously described (42). The plasmid DNAs were
preincubated with liposomes for 30 min in Opti-MEM I medium (Life Technologies, Inc./BRL) before the DNA/liposome mixture was
added to the cells. To generate stable cell lines, constructs encoding
ActRIIB-DN and a neomycin-resistant selectable marker were transfected
into HepG2 cells using lipofectin (Life Technologies, Inc./BRL). Stable transfectants were selected in 450 µg/ml
geneticin (Mediatech, Herndon, VA) and identified by RNA blot analysis.
Positive clones were expanded and maintained in geneticin.
Luciferase assay
HepG2 cells cultured in 12-well plates were transiently
cotransfected with p3TP-Lux (provided by Dr. Joan Massague, Memorial
Sloan-Kettering Cancer Center, New York, NY) and the indicated
expression constructs for 6 h. pcDNA3 vector DNA was used to keep
the total amount of DNA in all samples constant. After 6 h, the
cells were allowed to recover in fresh growth medium for 18 h and
were treated with or without 1 nM activin A in DMEM
containing 0.2% FBS for 20 h. Cells were washed with PBS twice
and lysed in 150 µl lysis buffer [25 mM HEPES (pH 7.8),
15 mM MgSO4, 0.5 mM EGTA,
1 mM dithiothreitol, and 0.2% Triton X-100]. Four hundred
microliters of assay buffer (lysis buffer minus Triton X-100 and with
2.5 mM ATP and 1 µg/ml BSA) and 100 µl 1 mM
luciferin (sodium salt, Analytical Bioluminescence, San Diego, CA) were
added to 100 µl cell lysate, and emitted luminescence over 10
sec was measured using a Monolight 2010 Luminometer (Analytical
Bioluminescence). Luciferase activity was normalized to the amount of
protein in each extract, determined using a Bio-Rad Laboratories, Inc., protein assay (Richmond, CA). Each experiment was repeated
at least three times.
Messenger RNA (mRNA) measurements
For RT-PCR, 5 µg total RNA were reverse transcribed into cDNA
using random hexameric oligonucleotides. Aliquots of the cDNA were then
amplified by PCR using an annealing temperature of 62 C with the
incorporation of [32P]deoxy-CTP (Amersham Pharmacia Biotech, Arlington Heights, IL). The PCR products were
separated by electrophoresis on a 6.5% polyacrylamide gel and
visualized by autoradiography. The sequence-specific primers used and
the expected sizes of the fragments amplified are shown in Table 1
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Fluorescence microscopy
HepG2 cells cultured on 15-mm square glass coverslips were
transfected with the indicated constructs. After recovery in fresh
medium for 24 h, cells were fixed with 4% paraformaldehyde and
mounted in Mowiol 4–88 (Calbiochem, La Jolla, CA)
mounting medium. Fluorescence was observed using a x63 oil immersion
objective on a Carl Zeiss Axiophot microscope (Carl Zeiss, Oberkochen, Germany).
In situ detection of apoptotic cells
Apoptosis was examined in situ by terminal
deoxynucleotidyltransferase-mediated deoxy-UTP nick end labeling
(TUNEL) using an ApopTag detection kit (Oncor, Gaithersburg, MD).
Briefly, HepG2 cells cultured on glass coverslips were fixed in 4%
formalin (Sigma) at room temperature for 10 min,
permeabilized with 0.5% Triton X-100 for 3 min, and stored in 70%
ethanol at -20 C for up to 3 days. Cells were then rehydrated in PBS,
transferred to a humidified chamber, and covered with equilibration
buffer for 5 min, which was replaced with the working strength terminal
deoxynucleotide transferase enzyme solution. After incubation at 37 C
for 1.5 h, the coverslips were submerged in the working strength
stop/wash buffer for 10 min and washed in PBS. dRhodamine-conjugated
antidigoxigenin antibody was applied to the cells for 1 h. After
the PBS wash, cells were mounted and visualized using a x40 or x63
oil immersion objective on a Carl Zeiss LSM410 confocal
microscope (fitted with an argon laser with a band at 488 nm for
fluorescein isothiocyanate (FITC) and an He-Ne laser with a band at 543
nm for rhodamine). GFP fusion proteins were detected using an FITC
filter set. Positive TUNEL staining was detected using a rhodamine
filter set. Fields in each of the four quadrants of each slide were
randomly chosen before visual inspection, and at least 300 cells/slide
were counted. Apoptotic cells were expressed as a percentage of the
total cells. Each experiment was repeated at least three times.
Statistical analysis
All values are expressed as the mean ± SD.
Student’s t test was used to evaluate differences between
the control samples and activin-treated samples or between samples
transfected with vector alone and samples transfected with the
different receptor and Smad constructs. P < 0.05 was
considered statistically significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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. Quantification of the TUNEL assays
revealed that activin A induced a dose-dependent increase in the number
of apoptotic cells, with a maximal effect observed between 1–10
nM activin A (Fig. 1B). The stimulatory effect of activin A
on apoptosis was antagonized by cotreatment with the activin-binding
protein follistatin at a 20-fold excess molar ratio (Fig. 1C). Inhibin,
which antagonizes the effects of activin in many systems, only
partially reduced activin A-induced apoptosis in HepG2 cells. Finally,
the related ligand TGFß induced apoptosis to a similar extent as did
activin A in HepG2 cells.
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. Neither the activin
ßA-subunit nor the activin
ßB-subunit mRNAs are expressed in HepG2 cells,
although they are readily detected in a control tissue known to express
activin, the ovary (Fig. 2A). In
contrast, mRNA for the activin-binding protein follistatin was detected
at a low level in HepG2 cells (Fig. 2A). Activin signaling is mediated
by at least four receptors (ActRI, -IB, -II, and -IIB) and several Smad
proteins, including Smad2 and Smad4 (17, 28, 29, 30). Figure 2B shows that
all of these signaling components are expressed in HepG2 cells,
although their relative abundance at the mRNA level varies quite
substantially.
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).
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). Expression of ActRIIB
alone increased basal activity 5-fold compared with that in control
cells. In activin-treated cells, neither ActRI, ActRII, nor ActRIIB
alone had an effect on reporter gene activity, whereas ActRIB increased
luciferase activity 2.5-fold compared with that in activin-treated
control cells. Strikingly, coexpression of one of the type I receptors
together with one of the type II receptors resulted in a synergistic
activation of the p3TP-Lux reporter gene. Even in the absence of
activin, basal activity in these cells reached levels as high as those
in activin-treated control cells. Activin treatment further increased
reporter gene activity in these cells. Coexpression of ActRIB and
ActRIIB gave the strongest ligand-independent activation of the
reporter gene, implying constitutive activation of the activin
signaling pathway by the combination of these two overexpressed
receptors.
To investigate further the involvement of activin type II receptors in
activin signaling, we overexpressed a truncated form of ActRIIB
(ActRIIB-DN) to interrupt activin signaling. Similar mutants have been
successfully used by others to inhibit activin signaling through a
dominant negative action (43, 44). Overexpression of ActRIIB-DN blocked
activin-induced transcriptional activation of the reporter gene in
HepG2 cells. Transfected wild-type ActRIIB was able to overcome the
inhibitory effect of the ActRIIB-DN mutant and restore the
transcriptional activation of the reporter gene in a dose-dependent
manner (Fig. 3B).
The involvement of Smad2 and Smad4 in activin signaling in HepG2 cells
was also investigated by examining transcriptional activation of the
p3TP-Lux reporter gene. Expression of either Smad2 or Smad4 alone had
little effect on reporter gene activity (Fig. 4A). However, coexpression of Smad2 along
with Smad4 resulted in a 70-fold increase in basal activity compared
with that in control cells. Reporter gene activity was further
increased 3-fold after activin treatment of the cells. Consistent with
our previous results, transfection of ActRIIB and ActRIB led to a
ligand-independent activation of the reporter gene (Fig. 4B). This
activity could be further increased when Smad2 or Smad4 was coexpressed
with the activin receptors. Maximal activation of the 3TP-Lux reporter
gene was obtained when all four proteins (ActRIB, ActRIIB, Smad2, and
Smad4) were coexpressed in the cells.
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). More importantly, Smad2-DN abolished the constitutive
activation of the reporter gene by activin receptors ActRIB and ActRIIB
(Fig. 4B).
Nuclear translocation of Smad2 and Smad4 proteins
The subcellular localization of Smad2 and Smad4 was examined after
activation of the activin signaling pathway in HepG2 cells. The Smad2
and Smad4 proteins were fused to the C-terminus of GFP, allowing direct
visualization of the fusion proteins by fluorescence microscopy.
Addition of the GFP did not disrupt the functional properties of the
Smad proteins, as determined by their ability to activate the p3TP-Lux
reporter gene in transfected HepG2 cells treated with activin (data not
shown). When expressed alone, GFP-Smad2 was expressed diffusely
throughout the cells. Activation of the activin signaling pathway by
coexpression of activin receptors ActRIB and ActRIIB caused Smad2 to
accumulate in the nucleus of 95% of the transfected cells (Fig. 5). Similar nuclear translocation of
GFP-Smad2 was also observed in cells cotransfected with combinations of
receptors ActRIB and ActRII, ActRI and ActRIIB, and ActRI and ActRII,
although the effect was not as robust (data not shown), consistent with
the transcriptional response data. In contrast, GFP-Smad4 was detected
predominantly in the cytoplasm regardless of receptor coexpression. In
cells cotransfected with Smad2 and the activin receptors, GFP-Smad4
translocated into the nucleus, although only in a subset of transfected
cells (
20%). Coexpression of activin receptors did not have any
effect on localization of the transfected GFP protein, which was found
throughout the cell (data not shown).
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). To determine
whether this mutant could also disrupt the activin-induced apoptotic
response, stable cell lines expressing ActRIIB-DN were established
(Fig. 6A). Cell lines 2 and 29 express
ActRIIB-DN RNA at high levels, whereas lines 35 and 52 do not express
detectable levels of ActRIIB-DN RNA. Because of its low abundance,
endogenous ActRIIB was below the level of detection in this RNA blot
analysis. Cell lines 35 and 52 maintained normal responsiveness to
activin, as shown by the transcriptional activation and apoptotic cell
death assays (Fig. 6, B and C). In ActRIIB-DN high expression cell
lines 2 and 29, activin-induced activation of the reporter gene was
abolished (Fig. 6B), consistent with our earlier results using
transient assays. TUNEL staining indicated that these cell lines no
longer exhibited an apoptotic response to activin (Fig. 6C). Activation
of the reporter gene could be rescued, and the apoptotic response could
be partially restored in these cell lines by overexpressing the
wild-type ActRIIB receptor (Fig. 6, B and C) or the ActRII receptor
(data not shown).
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).
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). Cotransfection of the Smad2-DN construct blocked the
stimulatory effect of cotransfected Smad2 and Smad4 on apoptosis.
Consistent with the function of the Smad proteins being downstream of
the activin receptor, the Smad2-DN construct also blocked HepG2 cell
apoptosis induced by either activin or cotransfected activin receptors
IB and IIB (Fig. 7).
To establish at a cellular level the relationship between Smad protein
expression and apoptosis, GFP-Smad fusion proteins were used to
visualize directly cells that were overexpressing the Smad proteins. In
cells transfected with GFP-Smad2 alone, GFP was colocalized with
positive TUNEL staining (Fig. 8). Similar
colocalization was observed in cells transfected with GFP-Smad4 or
GFP-Smad2 plus Smad4. Smad2-DN was able to suppress apoptotic cell
death induced by Smad2 or Smad4, indicating that apoptosis in Smad2- or
Smad4-overexpressing cells is a direct consequence of Smad2 or Smad4
activity in these cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Binding of activin to a type II receptor is the first step in the activin signaling pathway. In our studies, expression of the ActRIIB-DN mutant receptor that lacks the cytoplasmic domain resulted in a loss of activin A-stimulated transcriptional activation and cellular apoptosis. This is presumably because ActRIIB-DN is still competent for ligand binding and interaction with a type I receptor, but is unable to phosphorylate and activate the type I receptor (47, 48), and therefore prevents signal transduction. Activin responsiveness could be restored by the coexpression of exogenous wild-type Act II or ActRIIB, suggesting that both isoforms of the activin type II receptor are capable of binding the type I receptor and forming a functional receptor complex that transduces activin signals in HepG2 cells. Interestingly, ActRIIB-DN is less efficient at suppressing signaling in the presence of coexpressed ActRIB. This raises the possibility that endogenous ActRIB is being trapped by the mutant type II receptor, in agreement with studies showing an interaction between an analogous deletion mutant of the type II TGFß receptor and its type I receptor (48). However, in other studies a kinase-deficient ActRIIB receptor failed to block the activity of a constitutively active ActRIB, arguing against trapping of the type I receptor in this system (23).
Consistent with earlier overexpression studies on TGFß and activin receptors (18, 23), coexpression of both a type I and a type II receptor activated the activin signaling pathway even in the absence of the ligand, indicating there is a ligand-independent interaction between the two receptors. Biochemical analysis has confirmed that in the absence of activin, overexpressed ActRIB and ActRIIB are able to form a stable heteromeric complex (our unpublished results) (27).
Overexpression of either Smad2 or Smad4 caused HepG2 cells to undergo apoptosis. Although the transfected GFP-Smad4 fusion protein showed predominantly a cytoplasmic localization, even when cotransfected with receptors ActRIB and ActRIIB, it is likely that some fraction of the Smad4 does translocate to the nucleus and is sufficient to mediate an apoptotic signal. Using an inducible nuclear translocation expression system, it was reported that overexpression of Smad4 in the nucleus was able to induce cellular apoptosis, and a tumor-derived mutation affecting the DNA binding of Smad4 caused a significant decrease in apoptotic cell numbers (49). Taken together, these data suggest that nuclear Smad4 is able to induce apoptosis, an activity that may contribute to its tumor-suppressive role.
A critical role for Smad proteins in the pathway leading to apoptosis is further indicated by the results with the Smad2-DN mutant, which acts as a dominant negative regulator of activin A-dependent gene transcription and cellular apoptosis. This protein lacks the 10 amino acids at the C-terminus of the wild-type protein and hence is unable to be phosphorylated by the activated type I receptors. Phosphorylation of Smad2 is important for its association with Smad4 and subsequent nuclear translocation (45, 46). The ability of Smad2-DN to block activin, activin receptor, and Smad protein-mediated apoptosis suggests that Smad-dependent signaling to the nucleus is important for activin A-stimulated apoptosis. The Smad proteins are also important mediators of TGFß signaling (28, 29, 30). TGFß has been shown to induce apoptotic cell death in liver cells (13, 14), and we observed an apoptotic response to TGFß in HepG2 cells. It is likely that the Smad pathway is also involved in TGFß-induced liver cell apoptosis, providing for cross-talk between the activin- and TGFß pathways in regulating liver cell function.
Our identification of Smad proteins as intracellular mediators of activin-induced apoptosis provides a potential point for cross-talk between activin signaling and other signaling pathways involved in the regulation of apoptosis. One such pathway might involve Ca2+ signaling. Recently, calmodulin (CaM) has been shown to physically interact with Smad proteins in a calcium-dependent manner (50). CaM is an important intracellular receptor for calcium ions (51), and alterations in intracellular calcium appear to be essential for apoptosis. Calcium is required for activation of the latent Ca2+-dependent endonuclease that degrades internucleosomal DNA, a hallmark of apoptosis (51). Ca2+-bound CaM also activates a serine-threonine phosphatase, calcineurin, which has been demonstrated to induce apoptosis in the presence of calcium and the absence of growth factors (52).
The final stage of the apoptotic process, called execution, occurs through the activation and proteolytic function of a family of cysteine proteases, the caspases (53). Studies on the FaO rat and Hep3B human hepatoma cell lines provide evidence that the activity of the caspase-3 subfamily is stimulated in TGFß1-induced apoptosis (54, 55). In future studies it will be important to determine whether these same caspases are involved in activin-induced apoptosis and to establish how they are activated by TGFß family proteins. The inhibitory Smads, Smad6 and Smad7, have been shown to negatively regulate TGFß/activin signaling (38, 39, 40). In mouse B hybridoma HS-72 cells, Smad7 was induced by activin A and the ectopic expression of Smad7 suppressed activin A-induced apoptosis (56). It will also be important to determine whether Smad7 is induced by activin A and antagonizes activin A-induced apoptosis in HepG2 cells.
In summary, our data show that activin receptors and Smad proteins are expressed in HepG2 cells and are functional in an activin signaling pathway, leading to the activation of gene expression and cellular apoptosis. Furthermore, we show that activin receptors and Smad proteins are essential for mediating the apoptotic response induced by activin and that overexpression of activin receptors or Smad proteins can mimic activin action to induce apoptosis in HepG2 cells. These studies provide a framework for understanding interactions between the Smad protein signaling pathway and the pathways leading to cellular apoptosis and for understanding the complex roles of activin in modulating liver cell function.
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Acknowledgments |
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Footnotes |
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Received July 15, 1999.
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G. H. Su, R. Bansal, K. M. Murphy, E. Montgomery, C. J. Yeo, R. H. Hruban, and S. E. Kern ACVR1B (ALK4, activin receptor type 1B) gene mutations in pancreatic carcinoma PNAS, March 13, 2001; 98(6): 3254 - 3257. [Abstract] [Full Text] [PDF]
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 A. L. Lau, T. R. Kumar, K. Nishimori, J. Bonadio, and M. M. Matzuk Activin beta C and beta E Genes Are Not Essential for Mouse Liver Growth, Differentiation, and Regeneration Mol. Cell. Biol., August 15, 2000; 20(16): 6127 - 6137. [Abstract] [Full Text]
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