Expression of activins C and E induces apoptosis in human and rat hepatoma cells

Susanne Vejda*, Natascha Erlach*, Barbara Peter, Claudia Drucker, Walter Rossmanith2, Jens Pohl1, Rolf Schulte-Hermann and Michael Grusch3

Institute of Cancer Research, University of Vienna, Borschkegasse 8a, A-1090 Vienna, Austria and 1 BIOPHARM GmbH, Czernyring 22, D-69115 Heidelberg, Germany


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Activins C and E (homodimers of the ßC and ßE subunits), which are almost exclusively expressed in the liver, are members of the transforming growth factor ß (TGFß) superfamily of growth factors. We examined their expression in three different hepatoma cell lines and found that, compared with normal liver or primary hepatocytes, human hepatoblastoma (HepG2), human hepatocellular carcinoma (Hep3B) and rat hepatoma (H4IIEC3) cells have either completely lost or drastically reduced the expression of activins C and E. In order to elucidate the biological function of these proteins we transiently transfected HepG2, Hep3B and H4IIEC3 cell lines with rat activin ßC or ßE cDNA to study the consequences of restoring activin expression in hepatoma cells. Transfection with activin ßA, a known inhibitor of hepatic DNA synthesis and inducer of apoptosis, served as a positive control. We found that transfection of the three cell lines with activin ßC or ßE, as well as with activin ßA, reduced the increase in cell number by up to 40% compared with cells transfected with a control plasmid. Co-culture with a CHO cell clone secreting activin C also inhibited HepG2 cell multiplication. Furthermore, the three hepatoma cell lines studied showed an enhanced rate of apoptosis and elevated levels of active caspases in response to activin transfection. These results indicate that activins C and E share the potential to induce apoptosis in liver derived cell lines with activin A and TGFß1.

Abbreviations: EGFP, enhanced green fluorescent protein; FCS, fetal calf serum; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; IRES, internal ribosomal entry site; MEM, minimal essential medium; MTT, methylthiazoltetrazolium; MTX, methotrexate; PBS, phosphate-buffered saline; RPL6, ribosomal protein L6; TGFß, transforming growth factor ß


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Activins belong to the transforming growth factor ß (TGFß) superfamily of growth factors (1). This family includes a number of factors regulating cell proliferation and differentiation. To date, four mammalian activin subunits have been described, activins ßA, ßB, ßC and ßE (29). Mature proteins are composed of two ß subunits. Recently, we and another group demonstrated that homodimers (activins AA, BB, CC and EE) as well as heterodimers (activins AB, AC, AE and CE) are formed from the ß subunits (9,10). Activin A has been implicated in liver growth regulation by its capability to inhibit mitogen-induced DNA synthesis (11) and to induce apoptosis in vivo and in vitro (12,13). In addition, the activin-related protein TGFß1 inhibits DNA synthesis and causes apoptosis in normal and transformed hepatocytes (1417). Therefore, activin A and TGFß1 are supposed to act as negative regulators of liver growth.

The activin ßC gene was cloned and identified in human (3), mouse (5) and rat (9). The cDNA sequence of the activin ßE subunit has been reported for mouse (4), rat (7,9) and human (8). The two subunits share 82 and 61% amino acid sequence similarity of the mature peptides from rat and mouse, respectively, and they are thought to be a subset of related sequences (4,9). We found activin subunits ßC and ßE expressed almost exclusively in the liver, in hepatocytes (9). The activin ßC protein was localized to human liver and prostate (10). As in the rat, activin ßE mRNA is predominantly expressed in the liver in humans, with very low levels also detected in heart, testis, peripheral blood leukocytes, placenta and skeletal muscle (8). It has been postulated that activin C may act as a liver chalone, because, following partial hepatectomy, transient down-regulation of activin ßC mRNA was observed (1820). Decreased expression of activin C would reduce its inhibitory effect on cell proliferation to allow liver regeneration.

Lau et al. demonstrated that activin ßE mRNA increased rapidly and decreased to near basal levels by 48 h following partial hepatectomy (20). A rapid increase in mRNA was also reported in mice treated with lipopolysaccharide, which correlates with a pattern of acute phase response in the liver (7). Mice deficient in the activin ßC gene, activin ßE gene or in both genes appeared grossly normal. The animals were viable, survived to adulthood and liver regeneration following partial hepatectomy proceeded similar to that of wild-type mice (20).

Up to now, the biological roles of activins C and E have not been elucidated. Since the two factors display a liver-specific expression pattern and belong to the TGFß superfamily of growth factors, one may envision that activins C and E could also play roles in the sophisticated regulatory network that maintains a constant liver mass. In line with this hypothesis we have recently shown that overexpression of activin ßC or ßE in mouse liver can inhibit regenerative DNA synthesis (21).

To investigate potential functions of the liver-specific activins in the context of tumor cell growth, we initially examined the expression of activin subunits in logarithmically growing hepatoma cell lines in comparison with liver and primary hepatocytes. Since the human hepatoblastoma cell line HepG2, the rat hepatoma cell line H4IIEC3 and the human hepatocellular carcinoma cell line Hep3B all had either completely lost or drastically reduced expression of activins, we subsequently investigated the effects of ectopically expressing activin A (as a positive control), C or E in these cell lines. Transient transfection of activin ßA, ßC and ßE cDNA delayed the increase in cell number of all tested cell lines as determined by the methylthiazoltetrazolium (MTT) cell proliferation assay and cell counting. Furthermore, we demonstrated that these activins induce caspase activation and increase the rate of apoptosis in HepG2, H4IIEC3 and Hep3B cells.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cell culture
HepG2 (human, hepatoblastoma) cells were maintained in minimal essential medium (MEM) (Invitrogen) supplemented with 1 mM sodium pyruvate, 1% non-essential amino acids (Biochrom Kg) and 10% fetal calf serum (FCS). Hep3B (human hepatocellular carcinoma) and H4IIEC3 (rat hepatoma) were grown in RPMI-1640 (Invitrogen) supplemented with 10% FCS at 37°C in a humidified atmosphere containing 5% CO2. Primary rat hepatocytes were isolated and cultured as previously described (22) in accordance with the Austrian guidelines for animal care and protection.

Plasmids
The complete coding sequences of the rat activin subunits ßA (1275 bp) (23), ßE (1053 bp) (9) and ßC (1056 bp) (9) were cloned into either pTracer-CMV (ßA and ßE) or pcDNA3 (ßC) (Invitrogen). The sequences preceding the initiation codon were changed to a Kozak consensus sequence (24) by PCR mutagenesis in each case. Plasmid DNA was purified using a plasmid maxi kit (Qiagen). In addition, the complete coding sequences of the rat activin subunits were cloned into the multiple cloning site of pIRESe-EGFP. This vector contains the internal ribosomal entry site (IRES) of encephalomyocarditis virus between the MCS and the enhanced green fluorescent protein (EGFP) coding regions. This permits both the gene of interest (cloned into the MCS) and the EGFP gene to be translated from a single bicistronic mRNA. Empty vectors as well as a plasmid containing ß-galactosidase cDNA (pCMV-lacZ) were used as control plasmids.

Transient transfection of cells
HepG2, Hep3B and H4IIEC3 cells were seeded into either 96-well (4000 cells/well), 24-well (10 000 cells/well) or 12-well (20 000 cells/well) tissue culture plates (Greiner). Twenty-four hours after seeding DNA transfection using the FuGENE 6 Transfection Reagent (Roche Diagnostics Corp., Roche Molecular Biochemicals) was performed according to the instructions of the manufacturer. Twenty-four hours after transfection the medium was renewed. Plasmids containing both the activin cDNA and EGFP, activin cDNA alone, lacZ cDNA or plasmids alone were used for transient transfection of HepG2, Hep3B and H4IIEC3 cells. The EGFP protein served as a positive control to monitor uptake of plasmids.

Isolation of total RNA and RNase protection assay
HepG2, H4IIEC3 and Hep3B cells were directly lysed on the dishes using TRIzol Reagent (Life Technologies) and total RNA was isolated according to the instructions of the manufacturer. RNA was dissolved in 3 mM EDTA and the concentration was determined photometrically. RNA probes complementary to the respective cDNA sequences of rat activins ßA (235 bp), ßC (196 bp) and ßE (180 bp) and rat glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (101 bp) were used as previously described (9). To analyze the expression of human activins, RNA probes for human activin ßA (367 bp corresponding to nt 159–526 of the sequence with Genbank accession no. NM_002192), activin ßC (179 bp corresponding to nt 991–1170 of NM_005538), activin ßE (248 bp corresponding to nt 1042–1290 of NM_031479), human ribosomal protein L6 (RPL6) (132 bp corresponding to nt 363–495 of NM_000970) and human GAPDH (106 bp corresponding to nt 714–820 of NM_002046) were used. The housekeeping genes GAPDH and RPL6 were used as a control for sample loading and to compare expression levels. RNase protection assays were performed as previously described (25), with the following minor modifications: hybridizations were carried out at 51°C; RNase A and RNase T1 were used at 15 and 1 µg/ml, respectively. Dried gels were analyzed with a PhosphorImager and ImageQuant software (Molecular Dynamics).

SDS–polyacrylamide gel electrophoresis (SDS–PAGE) and western blotting
Hep3B cells were transfected with either activin ßA or empty vector and shifted to serum-free medium 24 h later. Another 72 h later proteins were precipitated from media supernatants with 1 vol of acetone, dissolved in loading buffer (7 M urea, 60 mM Tris–HCl pH 6.8, 2% SDS, 100 mM DTT, 0.01% bromphenol blue) and separated by 15% SDS–PAGE. Proteins were subsequently transferred to Hybond-P membranes (Amersham Pharmacia Biotech). Non-fat dried milk at 5% in TBST (50 mM Tris–HCl pH 7.5, 150 mM NaCl, 0.1% Tween 20) was used to block non-specific binding. The activin ßA antibody (kindly provided by W.Vale; 26) was used at 0.4 µg/ml in TBST containing 1% non-fat dried milk. Visualization was performed with an enhanced chemiluminescence (ECL Plus) detection system (Amersham Pharmacia Biotech).

Determination of cell number
As an indicator of cell number we used the MTT assay, which is based on the reduction of soluble yellow MTT tetrazolium salt to a blue insoluble MTT formazan product by mitochondrial succinic dehydrogenase (27). The amount of formazan product is proportional to the number of viable cells. MTT was added to each well to a final concentration of 0.5 mg/ml and incubation continued for an additional 2 h. Reduced MTT was dissolved in dimethyl sulfoxide and measured spectrophotometrically in a dual beam microtiter plate reader at 562 nm with a 620 nm reference. Experiments were performed in quintuplicate wells, repeated at least four times and the values are expressed as fold of control (cultures transfected with plasmid alone).

For counting of H4IIEC3 cells, they were detached with trypsin and their number determined in an automatic cell counter (CASY1-Model TTC; Schaerfe System).

Generation of activin C expressing CHO cells
The full-length cDNA of human activin ßC (3) was cloned into pABStopXS (28) containing a SV40 promoter, as well as a CMV enhancer, resulting in a plasmid named pAB 121. Plasmid pAB 121, as well as a plasmid pSVO dhfr have been co-transfected into CHO dhfr cells (29) at a ratio of 5:1 using a commercially available lipofection kit (Gibco). Upon transfection, cells were cultivated in the presence of increasing concentrations of methotrexate (MTX), ranging from 0.0005 to 6.0 µM MTX after 4 months, in order to amplify the transgene and to increase expression (30). Expression of activin C was verified by western blot. As a negative control, the empty plasmid pAB StopXS has been used instead of pAB 121.

HepG2–CHO co-culture
Tissue culture inserts (0.02 µm anopore membrane; Nunc) with either 150 000 activin C expressing or mock transfected CHO cells were placed in 6-well plates containing 40 000 HepG2 cells/well in MEM supplemented with 1 mM pyruvate, 1% non-essential amino acids and 10% FCS. Three days later part of the growth medium was renewed and on day 7 MTT assays were performed as described above. The experiment was repeated three times and the values are expressed as fold of control (co-culture with mock transfected CHO cells).

Scoring of apoptosis and detection of caspase activation
Twenty-four hours after transfection of the cells with either activin ßA, ßC or ßE cDNA or plasmid alone in 24-well plates, they were washed twice in phosphate-buffered saline (PBS), fixed for 15 min in 2% formaldehyde/PBS at room temperature, permeabilized with PBS/0.2% Tween 20 for 1 min and stained in 1 µg/ml Hoechst 33258 (Calbiochem) for 2 min. The percentage of cells displaying typical apoptotic nuclear morphology (crescent shaped condensed chromatin lining nuclear periphery; apoptotic bodies) (16,31), referred to as the apoptotic index, was then assessed using a fluorescence microscope. Four experiments in duplicate (two wells) were performed and between 300 and 800 nuclei per well were counted.

Caspase activity was detected with the CaspACETM FITC-VAD-FMK in situ marker (Promega). Twenty-four hours after transfection the marker was added to Hep3B cells to a final concentration of 10 µM and cells were incubated for 45 min. Then they were washed twice with fresh medium and assayed for FITC fluorescence under a fluorescence microscope. To compare the pattern of apoptotic chromatin changes with that of caspase activation, Hep3B cells were incubated with 5 µg/ml Hoechst 33258 and 10 µM FITC-VAD-FMK marker simultaneously and the same microscope frame was photographed once with filters for FITC and then with filters for Hoechst 33258.

Measurement of DNA synthesis rates
HepG2 cells (200 000 cells/dish) were seeded into 2.5 cm Nunc dishes and transfected as described above. Twenty-four hours later [3H]thymidine solution (0.5 µCi/ml) was added for 2 h. Fixation, processing for autoradiography and staining were done as described (32). Experiments were performed in duplicate, repeated three times and 1000 Hoechst stained nuclei per sample were counted.

Statistical analysis
All values are expressed as means ± SD. Student's t-test was used to evaluate differences between samples transfected with vector alone and samples transfected with activin cDNA.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Endogenous activin ßA, ßC and ßE subunits in human and rat hepatoma cells
In sharp contrast to normal human liver and primary rat hepatocytes, human and rat hepatoma and hepatocellular carcinoma cell lines express very little or no endogenous activin ß subunits (Figure 1). In the human hepatoma cell line HepG2 small amounts of activin ßE and ßC were found, whereas activin ßA expression was undetectable. In the hepatocellular carcinoma cell line Hep3B none of the activin subunits could be detected. The rat hepatoma cell line H4IIEC3 expresses activin ßE mRNA, albeit at several-fold lower levels than primary rat hepatocytes. Activin ßA and ßC were at undetectable levels. We have previously reported the absence of activin ßB, the fourth mammalian activin ß subunit hitherto described, from the rat liver (9).



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Fig. 1. (A) RNA expression of endogenous human activin subunits in HepG2 human hepatoblastoma and Hep3B human hepatocellular carcinoma cells as compared with two samples of human liver. Aliquots of 50 µg of total RNA were hybridized to riboprobes for human activin ßA, ßC and ßE and human ribosomal protein L6. (B) RNA expression of endogenous rat activin subunits in H4IIEC3 rat hepatoma cells and in primary rat hepatocytes. Aliquots of 10 µg of total RNA were hybridized to riboprobes for rat activin ßA, ßC and ßE and rat GAPDH.

 
Transient transfection of activin ßA, ßC and ßE cDNA in cultured human and rat hepatoma cells
In order to gain insight into the function of the activin subunits ßC and ßE, transfection studies with HepG2, H4IIEC3 and Hep3B cells were performed. Cells were transiently transfected with plasmids containing rat activins ßA, ßC and ßE cDNA as described in Materials and methods. In order to monitor activin expression after transfection of cDNA, total RNA was isolated from transfected HepG2, Hep3B and H4IIEC3 cells 24 h after transfection and RNase protection assays were performed. As shown in Figure 2, activin subunits ßA, ßC and ßE were highly expressed at a similar level in HepG2 and Hep3B cells using 0.5 or 5 µg total RNA, respectively. Since plasmids containing rat activin cDNA were used to transfect the cells, only the ectopically expressed activin subunits were detected in the RNase protection assays. However, in the rat cell line H4IIEC3 endogenous activin ßE mRNA was detected in addition to the forced expression of activin subunits, and only moderate levels of ectopic expression could be achieved. Uptake of plasmids in all cell lines was also controlled by examining the expression of EGFP protein. Transfection efficiencies of ~30% were achieved in HepG2 and Hep3B cells and slightly less in H4IIEC3 cells.



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Fig. 2. RNA expression levels of the rat activin subunits in (A) HepG2, (B) H4IIEC3 and (C) Hep3B cells transfected with rat activin cDNA. Seventy-two hours after transfection total RNA was isolated and 0.5 (HepG2 cells) or 5 µg (H4IIEC3 and Hep3B cells) of total RNA were analyzed by RNase protection assay using riboprobes hybridizing to rat activin ßA, ßC and ßE and human or rat GAPDH. The riboprobes were labeled to similar specific activity. pActivinßA, RNA of cells transfected with rat activin ßA cDNA; pActivinßC, RNA of cells transfected with rat activin ßC cDNA; pActivinßE, RNA of cells transfected with rat activin ßE cDNA; mock, RNA of control transfected cells.

 
To ensure that ectopically expressed activin is translated into protein and subsequently secreted into the medium, we checked for the presence of activin A in media supernatants of Hep3B cells transfected with either activin ßA or a vector control. Western blotting with anti-activin A antibody detected activin A in the supernatants of activin A transfected cells but not in untransfected or mock transfected cells (Figure 3).



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Fig. 3. Detection of activin A monomer by western blotting. Proteins of conditioned medium produced by activin ßA transfected Hep3B cells were precipitated and subjected to 15% SDS–PAGE under reducing conditions. As a negative control medium from untransfected (lane 1) and mock transfected (lane 2) Hep3B cells were treated alike. The expected size for monomeric activin A is 14 kDa.

 
Reduced growth of HepG2, Hep3B and H4IIEC3 cells in response to forced expression of activin A, C and E
To study the effects of activin expression on hepatoma cell lines, they were transfected with activin cDNA and growth properties of these cells were determined using the MTT assay. In this assay the metabolic activity of cells is used as a parameter for cell number. First, growth of transfected cells was analyzed 24, 48, 72 and 96 h after transfection (data not shown). As the most pronounced effect on cell growth was observed 72 h after transfection, this time point was selected for further MTT assays. The number of HepG2, H4IIEC3 and Hep3B cells was significantly decreased after transfection with activin ßC and ßE cDNA compared with cultures transfected with the empty vector, indicating a reduced growth rate in the presence of either activin C or E (Figure 4). The loss of MTT reducing activity was ~30–40% when cells were transfected with either activin ßC or ßE cDNA. Longer incubation times did not increase the effect (data not shown). In HepG2 and Hep3B cells transfection with activin ßA cDNA served as a positive control, because it is well known that activin A negatively affects the growth rate of these cells (33,34). Indeed, transfection of activin ßA cDNA resulted in growth inhibition in HepG2 and Hep3B cells. Furthermore, transfection with activin ßA was also inhibitory in H4IIEC3 cells. Of the three activins studied, activin ßA transfection induced the most conspicuous effect. Loss of MTT reducing activity of HepG2 and Hep3B cells was 30–50%, while that of H4IIEC3 cells reached as high as 75%. To demonstrate the specificity of the effects the unrelated ß-galactosidase protein was expressed in HepG2 cells, but had no effect on HepG2 cell number (Figure 4A). In order to determine whether the effect seen in the MTT assay was indeed due to reduced cell numbers (and not to reduced metabolic activity of equal numbers of cells), H4IIEC3 cells were counted using an automatic cell counter. As shown in Figure 4D, these experiments confirmed the results obtained by the MTT assays. Cell number was significantly reduced when cells were transfected with activin ß cDNA. Again, the effect on cell growth was weaker in the presence of activin ßC and ßE cDNA than after transfection with activin ßA cDNA.



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Fig. 4. Determination of the number of viable (A) HepG2, (B) H4IIEC3 and (C) Hep3B cells after transfection of activin ßA, ßC and ßE cDNAs, lacZ cDNA or plasmid alone by MTT assay 72 h after transfection. Data are expressed relative to cells transfected with plasmid alone. Bars represent the mean ± SD of 4–10 independent experiments (n). Significance levels: *P < 0.05, **P < 0.01, ***P < 0.001 (Student's t-test). (D) Number of H4IIEC3 cells was counted 72 h after transfection of activin ßA, ßC and ßE cDNA or plasmid alone using an automatic cell counter. A representative experiment is shown. (E) Determination of the number of viable HepG2 cells by MTT assay after 7 days co-culture with activin ßC transfected CHO cell clones versus mock transfected clones as controls.

 
Reduced HepG2 cell number caused by co-culture with activin C expressing CHO cells
To rule out potential artefacts caused by transient overexpression, HepG2 cells were maintained for 1 week in co-culture with a CHO cell clone stably overexpressing human activin ßC. CHO cells were kept in inserts in the medium of HepG2 cells, separated by a porous membrane. Under these conditions activin C expressing CHO cells reduced HepG2 cell number by 25% compared with the control, i.e. mock transfected CHO cells (Figure 4E). These results strongly suggest that secreted activin C protein inhibits cell multiplication of HepG2 cells.

Enhanced rate of apoptosis caused by transfection of hepatoma cells with activin subunits
Since activin A and TGFß1 are well known inducers of apoptosis in hepatic cells, we investigated whether the reduced cell numbers after transfection of HepG2, H4IIEC3 and Hep3B cells with activin ßA, ßC and ßE cDNA are a consequence of an increased rate of cell death. Therefore, cells were transfected with activin cDNA and 24 h later they were fixed and the cell nuclei were stained with Hoechst 33258. The 24 h time point was chosen for apoptosis detection because in initial studies at later time points the analysis was impeded by the debris from apoptotic cells, which had subsequently undergone secondary necrosis. As shown in Figure 5D, H4IIEC3 cells displayed typical features of apoptotic cell death. These pictures of nuclei were similar in HepG2 (not shown) and Hep3B (Figure 6B) cells. The background level of apoptotic cell death due to the transfection of empty plasmid was low (between 1 and 4% on average). Compared with control cells, transfection of activin ßA, ßC and ßE cDNA resulted in significantly higher percentages of condensed and fragmented nuclei in all three tested cell lines, as shown in Figure 5A–C. Again, activin ßA transfection resulted in the most pronounced effect in HepG2 and H4IIEC3 cells. Hep3B cells were the cell line most sensitive to apoptotic cell death after transfection with activin ßE and ßC.



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Fig. 5. Effect of ectopic expression of activins on rate of apoptosis in (A) HepG2, (B) H4IIEC3 and (C) Hep3B cells. Twenty-four hours after transfection with activin cDNA or plasmid alone, cells were fixed, stained with Hoechst 33258 and normal and apoptotic nuclei were counted. Four independent experiments (n = 4) were performed in duplicate for each transfection using activin ßA, ßC or ßE cDNA or plasmid alone and counting at least 300 cells per well. The percentage of apoptotic nuclei is shown as mean ± SD of four experiments. (D) Example of Hoechst stained nuclei of H4IIEC3 cells. Cells were transfected and stained as described above. Arrows indicate apoptotic nuclei.

 


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Fig. 6. Comparison between apoptosis detection with Hoechst 33258 and labeling of active caspases. (A) Hoechst 33258 and the FITC-VAD-FMK marker were added simultaneously to Hep3B cells 24 h after transfection with activin cDNAs or empty vector and the percentages of apoptotic nuclei (Hoechst) and of FITC labeled cells (caspACE) were determined. The experiment was done in duplicate and at least 300 cells per well were counted. (B) Micrographs of Hep3B cells transfected with activin ßC and stained as described above. The same microscope frame was photographed with filter sets for Hoechst 33258 (left) and FITC (right). Note that cells with typical condensed apoptotic nuclei (arrows in left panel) are also strongly positive in the caspase assay (arrows in right panel).

 
To further confirm the induction of apoptosis, we investigated the activation of caspases in Hep3B cells, which had shown the most pronounced effect with regard to the appearance of apoptotic nuclei following activin transfection. Activated caspases have been shown to be important executioners of apoptosis and are responsible for many of the morphological and biochemical changes associated with this form of cell death (35). Binding of the FITC-VAD-FMK marker to the catalytic center of activated caspases allows the identification of caspase activation in single cells (36). Transfection with activin ßA, ßC or ßE resulted in an increase in FITC labeled Hep3B cells, suggesting that activin expression induces caspase activation and consequently apoptosis (Figure 6). Moreover, double labeling of cells with FITC and Hoechst 33258 showed good agreement between caspase activation and apoptotic chromatin changes.

DNA synthesis rates in HepG2 cells after transfection with activin subunits
Activin A has been shown to inhibit DNA synthesis rates in rat hepatocytes (11). Therefore, we determined whether transfection of HepG2 cells with activin cDNAs results in decreased rates of cells undergoing DNA synthesis. For this purpose we used the same conditions under which pro-apoptotic activity was demonstrated. To eliminate a potential bias due to DNA of apoptotic cells, autoradiography was chosen and apoptotic nuclei were not included in the count. Under these conditions supplementation with [3H]thymidine revealed no decrease in labeled nuclei in activin transfected compared with mock transfected cells (Table I).


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Table I. Determination of the DNA synthesis rates of HepG2 cells 24 h after transfection with the respective activin ß subunits, the lacZ gene or empty vector

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Up to now, the biological functions of activins C and E have not been elucidated. Activin C was suggested to act as a liver chalone (18). It has been proposed that liver mass is regulated by an inhibitory factor produced within the liver. This putative factor, designated liver chalone (37), inhibits proliferation of hepatocytes. Expression studies in humans, rats and mice revealed the liver as the major source of activin ßC and ßE transcripts (39,18). On the basis of structural homology, activins belong to the TGFß superfamily of cytokines. Among these, TGFß1 and activin A inhibit mitogen-induced DNA synthesis (17) and induce apoptosis in hepatocytes in vivo and in vitro (1215). Despite their high structural similarity (9), it has not been clarified whether activins C and E have similar potential.

We examined the expression of activins A, C and E in hepatoma cell lines and found that in contrast to normal human liver or primary rat hepatocytes (and normal rat liver; not shown), hepatoma cells have partially or completely lost activin expression. Moreover, we could demonstrate that transient transfection with activin cDNA significantly decreased the number of HepG2, Hep3B and H4IIEC3 cells compared with control cultures. This effect was observed using two different detection systems in three different cell lines. Cell number and metabolic activity of cells were determined. The MTT assay is based on metabolic conversion of a defined substrate and the amount of the product is proportional to the number of viable cells. Therefore, this assay can be employed to measure a differential increase in cell number. An inhibitory effect of activin A on hepatic cell proliferation has been described by several authors (33,38,39). We demonstrate for the first time that, like activin A, activins C and E also reduce the increase in hepatoma cells and enhance the rate of apoptosis. This was demonstrated by the classical method of observing apoptotic changes in nuclear morphology and confirmed by detection of activated caspases.

When both detection methods were compared we observed that the great majority of cells were either positive or negative in both assays. Nevertheless, some cells displayed activated caspases but not apoptotic nuclei and vice versa. While the first case can easily be explained by the fact that changes in nuclear morphology occur at least in part as a consequence of caspase activation, the latter case probably represents cells in a late stage of cell death when activated caspases are no longer detectable.

The increase in apoptosis rates following expression of activins C and E, although weaker than with activin A in two of the three tested cell lines, was highly significant when compared with mock transfected controls. In HepG2 and Hep3B cells all three activin subunits were expressed to high levels, indicating that the transfection efficiency of the plasmids was similar. Transfection efficiency in the rat hepatoma cells was generally lower than that in the other tested cell lines, yet they showed the most dramatic effect in response to forced expression of activin ßA. Since H4IIEC3 cells produce considerable amounts of endogenous activin ßE RNA, we achieved only a moderate increase in total activin ßE RNA levels after transfection. Nevertheless, activin ßE transfected H4IIEC3 cells reproducibly displayed an increase in apoptosis rates compared with vector controls. A possible explanation may be that although the overall increase in activin production was low due to a lower transfection efficiency than in the other two cell lines, the local increase in the vicinity of transfected cells may still have sufficed for apoptosis induction. Due to the lack of an appropriate quantitative assay, the amounts of activins C and E produced by cells transfected with activin cDNA could not be determined. A comparison of the mRNA expression levels of activins with that of the housekeeping gene GAPDH suggests, however, that the expression levels achieved by forced expression are within a similar range to those found in primary rat hepatocytes. By precipitating activin A from the medium supernatant of transfected cells we have demonstrated that activin A protein is produced and secreted by transfected cells. Due to the lack of equally sensitive antibodies for activins C and E we could not detect activin E in media supernatants of transfected hepatoma cells and could detect activin C only in stably transfected CHO cells (not shown). However, we have previously demonstrated, with a 2-dimensional polyacrylamide gel electrophoresis approach, that all three activins are produced and secreted following transfection of the same plasmids as were used in this study into 293T embryonic kidney cells, which show 2- to 3-fold higher transfection efficiencies (9). While the lack of effect of the unrelated protein ß-galactosidase and the effectiveness of activin C in the co-culture experiment demonstrate the specificity of the activin effects, quantitative dose–effect determination will have to await the availability of bioactive recombinant activins C and E.

As mentioned above, several members of the TGFß superfamily of growth factors have been implicated in the control of the birth–death balance of liver cells. TGFß1 is a potent inhibitor of hepatocyte proliferation and inducer of apoptosis. The concentration of activin A required to induce apoptosis to the same extent was reported to be 10-fold higher compared with TGFß1 and the factor must be continuously expressed for 24 h for maximal response (12). While knockout mice for activin ßC, ßE or both genes show no apparent phenotype (20), probably due to compensation of their function by other family members, we have recently shown inhibition of regenerative DNA synthesis in mice overexpressing activin ßC or ßE in the liver (21). Our data suggest that in HepG2 cells induction of apoptosis rather than a decrease in the percentage of cells synthesizing DNA is the predominant effect of activin action. A pro-apoptotic effect of activins C and E would be consistent with our observation that all three of the investigated hepatoma and hepatocellular carcinoma cell lines show either no or greatly reduced expression of activins ßC and ßE (and of activin ßA) when compared with liver or primary hepatocytes. This suggests that the cells dispose of the growth suppressing effect of activins during the development of malignancy. Along similar lines, we have recently reported that follistatin, which binds to and inactivates activin A (4042) and also binds activin E (8), is up-regulated in a high proportion of hepatic tumors in rats and mice (43). These findings suggest that liver tumor cells counteract the growth inhibitory effects of activins by reducing their synthesis and/or by producing the activin inhibitor follistatin. In conclusion, activins may be important factors for the maintenance of homeostasis of cell number in the liver.


    Notes
 
2 Present address: Neuromuscular Research Department, Institute of Anatomy, University of Vienna, Währinger Strasse 13, A-1090 Vienna, Austria Back

3 To whom correspondence should be addressed. Tel: +431 4277 65144; Fax: +431 4277 65159; Email: michael.grusch{at}univie.ac.at Back

* These two authors contributed equally to first authorship. Back


    Acknowledgments
 
This work was supported in part by a grant from the Herzfelder'sche Familienstiftung to M.G. We wish to thank Paul Breit for his assistance in preparing the figures.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received January 15, 2003; revised August 12, 2003; accepted August 14, 2003.