Cyclin D1 over-expression correlates with ß-catenin activation, but not with H-ras mutations, and phosphorylation of Akt, GSK3ß and ERK1/2 in mouse hepatic carcinogenesis

Junichi Gotoh, Masahiko Obata, Masumi Yoshie, Shinichi Kasai1 and Katsuhiro Ogawa2

Department of Pathology and
1 Department of Surgery, Asahikawa Medical College, 2-1-1-1 East, Midorigaoka, Asahikawa 078-8510, Japan


    Abstract
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 Abstract
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 Materials and methods
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 Discussion
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Mutational activation of ß-catenin and cyclin D1 over-expression are a frequent change in mouse hepatic tumors. Although activated ß-catenin may bind to T cell factor (TCF) family members and transcriptionally activate the cyclin D1 gene, either ß-catenin or cyclin D1 may be activated by various pathways independently of ß-catenin mutations. In this study, we investigated ß-catenin activation and mutations, cyclin D1 expression, H-ras mutations and phosphorylation of extracellular signal regulated protein kinases 1/2 (ERK1/2), Akt and glycogen synthetase kinase 3ß (GSK3ß) in mouse hepatic carcinogenesis. Nuclear/cytoplasmic staining of ß-catenin, a sign of ß-catenin activation, was frequently observed in association with the high nuclear cyclin D1 labeling index in the hepatic tumors at the late stage of carcinogenesis. The ß-catenin activation was further suggested by the fact that all hepatocellular carcinoma (HCC) cell lines examined showed the nuclear ß-catenin/TCF4 complex together with cyclin D1 over-expression. However, the fact that only 31.8% (7/22) of the lesions with the nuclear/cytoplasmic ß-catenin staining showed ß-catenin mutations indicated that ß-catenin was activated not only by its own mutations but also by other reason(s). On the other hand, there was no correlation between the ß-catenin/cyclin D1 activation and the H-ras mutations or phosphorylation of Akt, GSK3ß and ERK1/2, although GSK3ß was frequently over-expressed in the tumors. These results indicate that, although ß-catenin and cyclin D1 activation are well correlated, the Akt/GSK3ß and ras/ERK1/2 pathways may not play a major role in the ß-catenin/cyclin D1 activation.

Abbreviations: DEN, diethylnitrosamine; ERK, extracellular signal regulated kinase; GSK3ß, glycogen synthetase kinase-3ß; HCC, hepatocellular carcinoma; Igf, insulin like growth factor; L.I., labeling index; PI3K, phosphatidyl inositoside 3-phosphate kinase; SSCP, single strand conformation polymorphism; TCF, T cell factor.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The hepatocellular carcinoma (HCC) is one of the most prevalent human malignancies in the world and a major cause of death in many countries (1). Tumors usually develop in livers that are chronically injured by hepatitis B or C viruses, alcohol or other hepatotoxic substances such as aflatoxin B1. The molecular mechanisms that transform normal hepatocytes to neoplastic cells, however, are poorly understood.

Recently, mutations of ß-catenin have been reported to be frequent in both human and rodent HCCs, suggesting a role for ß-catenin in neoplastic progression of hepatic cells (29). They usually involve the N-terminal glycogen synthetase kinase 3ß (GSK3ß) phosphorylation sites and may therefore result in resistance to the ß-TrCP F box protein-mediated ubiquitination-proteasome degradation of ß-catenin protein, hence causing its accumulation and transfer to nuclei, where it binds to a T cell factor (TCF) family members, and may up-regulate various genes of which expression impacts on processes such as cell proliferation, apoptosis, cellular communication, angiogenesis, invasion and metastasis (10,11).

While cyclin D1 is frequently over-expressed in HCC, in only a proportion of the cases it is due to amplification of the cyclin D1 gene, and other mechanisms remain to be clarified (12). Cyclin D1 activates cyclin-dependent-kinase 4/6 that phosphorylates the Rb protein and, thereby, promotes the G1 to S transition. Although ß-catenin may bind to TCF and transcriptionally up-regulate the cyclin D1 gene expression (1315), various other signaling pathways may cause the cyclin D1 over-expression. For example, ras signaling can up-regulate transcription from the cyclin D1 gene (1621) and cyclin D1-cdk 4 assembly (22), via a pathway that depends upon the sequential activation of ras, raf-1, ERK kinase and mitogen-activated protein kinase/extracellular signal regulated protein kinase (MAPK/ERK). On the other hand, recent evidence suggested that both ß-catenin and cyclin D1 might be simultaneously up-regulated by various signaling pathways. For example, activation of growth factor receptors, erbB2 (23), met (24) and insulin like growth factor (Igf) 1 receptor (25), and the activated ras (26,27) have been shown to lead to ß-catenin tyrosine phosphorylation which results in disruption of adherens complexes and liberation of ß-catenin from cell membrane to cytoplasm and nuclei. Furthermore, the growth factor receptors and ras activate the phosphatidyl inositoside 3-phosphate kinase (PI3K)–Akt pathway, which results in inactivation of glycogen synthetase kinase 3ß (GSK3ß) by phosphorylation at serine 9, resulting in accumulation of ß-catenin (28,29) and cyclin D1 (3033).

Both activating mutations of H-ras, usually at codon 61 (34,35), and ß-catenin mutations, usually in its exon 2 encoding possible N-terminal GSK3ß phosphorylation sites (2,4,5,8,9), were detected in liver tumors in mice. Moreover, mouse hepatic tumors frequently over-express cyclin D1 (36). The present study was conducted to address whether ß-catenin or H-ras mutations may be linked to the cyclin D1 expression in hepatic tumors, whether phosphorylation of Akt, GSK3ß and ERK1/2 may be correlated with the ß-catenin/cyclin D1 activation, and at what stage such changes may occur during multi-step hepatic carcinogenesis.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Treatment of animals
Male B6C3F1 mice (Clea Japan, Tokyo, Japan) were intraperitoneally administered diethylnitrosamine (DEN) at a dose of 5 µg/g body weight at the age of 2 weeks. The mice were then maintained on normal diet (CE-2, Clea Japan) and water ad libitum and killed after 8–14 months.

Immunohistochemistry
The liver of the DEN-treated mice were fixed by perfusion through the portal vein or vena cava with 4% formaldehyde solution, processed and embedded in paraffin for sectioning. Sections through lesions were histologically examined after hematoxylin and eosin staining, and hepatic tumors were classified according to the published criteria (37). For immunohistochemistry, deparaffinized sections were microwaved in 0.01 M citrate buffer, pH 6.0, for 10 min in an electric oven, treated with 3% hydrogen peroxide dissolved in ethanol for 60 min and incubated with the different antibodies for overnight at 4°C. The antibody dilution was 1:100 for anti-ß-catenin (Transduction Laboratories, Lexington, KY) and 1:200 for anti-cyclin D1 (Upstate Biotechnology, Lake Plasid, NY). Antibody binding was visualized by a Histofine kit (Nichirei, Tokyo, Japan). Whole views of immunohistochemically stained sections were scanned using a Nikon 35 mm film scanner and saved in the Macintosh computer for comparison of the intensity and pattern of staining in individual lesions.

PCR-single strand conformation polymorphism (SSCP) analysis
Hepatic tumor tissues were dissected from the sections serial to the immuno-stained sections using a dissecting microscope. DNA was extracted from the dissected tissues with DexpatTM (Takara, Otsu, Japan) according to the manufacturer’s instructions, precipitated in ethanol, dried and dissolved in H2O. DNA and RNA were also extracted from the HCC cell lines that had been established in our laboratory (38). For detection of ß-catenin mutations, 150 bp fragments including the codons encoding the GSK3ß phosphorylation sites in exon 2 were amplified from the isolated DNA using the primer pairs, 5'-GGAGTTGGACATGGCCATGG-3' (forward) and 5'-TCAACATCTTCTTCCTCAGG-3' (reverse). For detection of exon 2 deletions, the total RNA was reverse-transcribed for generation of first strand cDNA using oligo(dT)16 primers, and aliquots of the reaction mixtures were used for the subsequent PCR. The forward PCR primer (5'-GCGTGGACAATGGCTACTCAAG-3') for this purpose targeted the end of exon 1, and the reverse primer (5'-GTCATTGCATACTGCCCGTCAA-3') the beginning of exon 3. The primer sequences for H-ras exon 2 were 5'-GACAGAATACAAGCTTGTGG-3' (forward) and 5'-AGTGGGATCATACTCGTCCAC-3' (reverse), and those for exon 3 were 5'-GTGGTCATTGATGGGGAGAC-3' (forward) and 5'-CTGATGGATGTCCTCGAAGG-3' (reverse). PCR was carried out in a reaction volume of 15 µl for 3 min for first denaturation at 94°C, followed by 35 or 45 cycles of 94°C for 1 min, 56°C for 1 min and 72°C for 1 min and finally 72°C for 7 min. Mutations were detected by the SSCP analysis. Amplified DNA was mixed with an equal volume of formamide loading dye (95% formamide, 0.05% bromphenol blue and 0.05% xylene cyanol), denatured at 95°C for 5 min and rapidly cooled on ice. The samples were then electrophoresed with a GeneGel Excel 12.5/24 Kit (Amersham Biosciences, Uppsala, Sweden) and stained with a PlusOne DNA silver staining kit (Amersham Biosciences).

DNA sequencing
When abnormal bands were detected by the SSCP analysis, they were dissected from the gel, and DNA was extracted by boiling in 10 µl distilled water for 15 min. Extracted DNA was amplified by PCR as described above, purified with the Microcon filters (Millipore, Bedford, MA), and sequenced using dye terminator cycle sequence chemistry with AmpliTaq polymerase FS (Applied Biosystem, Foster City, CA). Sequencing was performed in both directions using the PCR primers described above. The samples were run on a fluorescence automated DNA sequencer (ABI Prism 377 DNA Sequencer, Applied Biosystems).

Western blotting
Fresh hepatic tumors were removed from the livers of DEN-treated mice between 8 and 14 months. The tissues were divided into two pieces, one of which was frozen by liquid nitrogen and the other fixed in the 4% formaldehyde solution for histological examination and immunohistochemical analysis for ß-catenin and cyclin D1. Cells were also isolated from subconfluent cultures of the HCC cell lines. Both tissues and cells were lysed using a solution containing 50 mM Tris (pH 8.0), 120 mM NaCl, 0.5% Nonidet P40 and 100 mM NaF, and aliquots of 50 µg of denatured protein were run on 10% polyacrylamide gels containing 0.1% sodium dodecyl sulfate and then electro-blotted onto nitrocellulose filters. The filters were then blocked with 5% non-fat dry milk in phosphate-buffered saline/0.2% Tween 20 and incubated with 1:100 diluted anti-ß-catenin, 1:200 diluted anti-cyclin D1 or 1:200 diluted {alpha}-tubulin antibodies. For ERK1/2, Akt and GSK3ß, the filters were incubated with the antibodies against 1:100 diluted either total or phosphorylated ERK1/2 (Promega, Madison, WI), Akt (Cell Signaling Technology, Beverly, MA) and GSK3ß (Cell Signaling Technology). After extensive washing, they were then reacted with peroxidase-labeled anti-mouse or anti-rabbit secondary antibodies (Santa Cruz Biotechnology, Santa Cruz, CA) and washed again. Finally, bound antibodies were visualized using the ECLTM system (Amersham Biosciences). The band intensity in the western blot analysis was quantified by the NIH image software and standardized using {alpha}-tubulin.

Gel retardation assay
Gel retardation assays were performed as described previously (39). Nuclear protein extracts were prepared from the HCC cell lines as described (40), carefully avoiding contamination with cytoplasmic proteins. A double strand oligonucleotide probe containing the TCF4 binding sequence (5'-CCCTTTGATCTTACC-3') was synthesized and end-labeled with [{gamma}-32P] using a MegalabelTM DNA 5'-end labeling kit (Takara). A typical binding reaction contained 5 µg nuclear proteins, 32P-labeled probe (0.05 ng) and 50 ng deoxyinosine-deoxycytidine in 25 µl buffer containing, 10 mM Tris–HCl, 50 mM NaCl, 5 mM dithiothreitol, 1 mM EDTA, 0.5 mg/ml bovine serum albumin and 4% glycerol. Samples were incubated for 30 min at room temperature. Unlabeled TCF4 probe in excess was used as a specific competitor, and an unlabeled NF{kappa}B probe as a negative control. For the super-shift reaction, nuclear protein extracts were pre-incubated with the anti-ß-catenin antibody for 1 h on ice. The anti-c-myc antibody was used as a negative control in this case. DNA binding reactions were visualized by autoradiography after resolution on 4% polyacrylamide gels.

Statistics
The data were statistically evaluated with Statview software (SAS Institute, Cary, NC). Differences were analyzed using the two-tailed {chi}2, and significance was defined as a P value <0.05. Correlation between the expression levels of cyclin D1 and phosphorylated/total Akt, GSK3ß and ERK1/2 was analyzed by the Pearson’s coefficient test, and the significance was defined as R2 value >0.187.


    Results
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 Abstract
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 Materials and methods
 Results
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Immunohistochemical staining of ß-catenin and cyclin D1
In normal hepatocytes surrounding the hepatic tumors, ß-catenin staining was evident at the cell membranes (Figure 1A, b and B, bGo), and the nuclear cyclin D1 labeling was rarely observed (Figure 1A, c and B, cGo). In the foci, ß-catenin was localized to cell membranes as in normal hepatocytes, but the staining was sometimes weaker than in the surrounding normal hepatic tissue (Figure 1A, bGo). The nuclear cyclin D1 labeling index (L.I.) was <5% in most foci (Figure 1A, cGo), but 5–20% in a few cases (Table IGo). In adenomas, ß-catenin was stained in the nuclei/cytoplasm/membranes in one case and the cytoplasm/cell membranes in about half of the cases, but only in the cell membrane in the other (Table IGo, Figure 1B, bGo). The nuclear cyclin D1 L.I. for adenomas was almost the same as for the foci (Table IGo, Figure 1B, cGo). In HCCs, nuclear ß-catenin staining was observed, at least in part, together with cytoplasmic/membrane staining in all but one of the cases (Table IGo, Figure 1B, bGo). The nuclear cyclin D1 L.I. was high with eight of 22 HCCs having values >20%, six of 22, 10–20% and eight of 22, 5–10% (Table IGo, Figure 1B, cGo).



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Fig. 1. Hematoxylin and Eosin (a) and immunohistochemical staining (b, c) for a focus and an adenoma adjacent to a HCC. While closed arrowheads indicate the boundary between the normal liver (N) and the focus (A, b, c) or adenoma (A, d) (B, a), and open arrowheads indicate the boundary between the adenoma and the HCC (B, a). Although ß-catenin is expressed only in the cell membrane in the focus (A, b) and the adenoma (B, b), the cytoplasmic/nuclear expression is evident in the HCC (B, b, inset). Nuclear cyclin D1 staining is rare in the focus (A, c) and the adenoma (B, c), but frequent in the HCC (B, c, inset).

 

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Table I. Correlation between the ß-catenin staining pattern and cyclin D1 expression
 
Mutations of ß-catenin and H-ras
Tissues from 15 foci, 55 adenomas and 22 HCCs, and 25 HCC cell lines were examined for ß-catenin and H-ras mutations. SSCP analysis of ß-catenin exon 2 detected abnormal bands in one of 55 adenoma and seven of 22 HCCs and four of 25 HCC cell lines, but in none of the 15 foci (Table IIGo, Figure 2A, aGo). DNA sequencing revealed missense mutations at codons 32, 33, 34, 41, 44 and 45 that encode serine or threonine residues of putative GSK3ß phosphorylation or neighboring sites in all except for two cell lines (Table IIGo, Figure 2A, bGo). In the latter cases, exon 2 was completely deleted in the cDNA (Figure 2B, a, bGo).


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Table II. Summary of muation analysis
 


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Fig. 2. Representative results of mutation analysis. (A) Screening for ß-catenin mutations by SSCP (A, a). Arrowheads point to abnormal bands. Sequencing analysis results are shown (A, b). Left: HCC on lane 1 (A, a) showing a C to G substitution at the second base of codon 33. Right: HCC on lane 2 showing a C to T substitution at the second base of codon 45. (B) Results of RT–PCR analysis for deletion of ß-catenin exon 2 ({Delta}exon 2) in HCC cell lines (B, a). In addition to the normal exons 1–3 bands, a lower band is seen in two cell lines. DNA sequencing reveals deletion of ß-catenin exon 2 in the lower band (B, b). (C) Screening for H-ras mutations by SSCP (C, a). Arrowheads: abnormal bands. Results of DNA sequencing of abnormal bands (C, b). Left: adenoma on lane 6 (C, a) showing a G to A substitution at the first base of codon 12. Right: HCC cell line on lane 11 (C, a) with an A to T change at the second base of codon 61.

 
Abnormal SSCP bands were detected in H-ras exons 1 or 2 in two of 15 foci and five of 55 adenomas, and in H-ras exon 2 in three of 25 cell lines (Table IIGo, Figure 2C, aGo), but in none of the HCCs. DNA sequencing revealed the H-ras mutations to be G to A substitution at the first base of codon 12 in two foci and three adenomas, A to G change at the second base of codon 61 in two adenomas, and A to T change at the second base of codon 61 in three cell lines (Table IIGo, Figure 2C, bGo).

Correlations of ß-catenin mutations, nuclear/cytoplasmic ß-catenin staining and cyclin D1 expression
All of the seven HCCs with ß-catenin mutations exhibited nuclear/cytoplasmic ß-catenin staining, and one adenoma with the ß-catenin mutation showed cytoplasmic/membrane localization. The nuclear cyclin D1 L.I. was 10–20% or >20% in most of the cases (Figure 3Go). On the other hand, two foci and three adenomas with H-ras mutations showed membrane staining of ß-catenin, and the other two adenomas with H-ras mutations showed cytoplasmic/membrane staining. The cyclin D1 L.I. was <5% in all the lesions with H-ras mutations. The cyclin D1 L.I. was also <5% in the lesions with membranous ß-catenin staining except for a few cases.



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Fig. 3. Correlation of ß-catenin staining patterns and ß-catenin/H-ras mutations with cyclin D1 expression in foci (open columns), adenomas (gray columns) and HCCs (closed columns). The intensity of cyclin D1 staining is clearly correlated with the nuclear/cytoplasmic ß-catenin staining (P < 0.0001) as well as ß-catenin mutations (triangle), but not with H-ras mutations (circle). M, the membrane staining of ß-catenin; C/M, cytoplasmic/membrane staining; N/C/M, nuclear/cytoplasmic/membrane staining.

 
ß-Catenin/TCF4 complexes in HCC cell lines
It was further investigated whether ß-catenin formed complexes with TCF4 in hepatic tumor nuclei using six HCC cell lines. These cell lines showed high ß-catenin and cyclin D1 protein expression regardless of the presence or absence of ß-catenin or H-ras mutations (Figure 4AGo). One cell line showed a truncated ß-catenin protein probably derived from ß-catenin mRNA lacking exon 2 (Figure 4AGo, lane 5). Nuclear proteins extracted from one of the ß-catenin mutated HCC cell lines (cell line 1 in Figure 4AGo) showed a ß-catenin/TCF4 complex shift band (Figure 4BGo). The band was further super-shifted in the presence of the anti-ß-catenin antibody, but not by the anti-c-myc antibody. It disappeared on addition of excess unlabeled TCF4 probe, but not unlabeled NF{kappa}B probe. Shift and super-shift bands were also detected in all the other HCC cell lines examined (Figure 4CGo).



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Fig. 4. ß-Catenin expression and nuclear ß-catenin/TCF4 complexes in HCC cell lines. (A) Western blot analysis of ß-catenin and cyclin D1 in HCC cell lines, with or without ß-catenin and/or H-ras mutations. All cell lines express various amounts of ß-catenin and high cyclin D1. Lane 5 shows a truncated band as well as the normal band. Note {alpha}-tubulin levels as an internal standard. (B) Representative results of the gel retardation assay. Incubation with the 32P-labeled TCF4 probe and nuclear extract of a HCC cell line with a ß-catenin mutation (HCC cell line No. 1) results in a band shift (lane 2) which is further super-shifted by addition of anti-ß-catenin antibody to the reaction mixture (lane 3). Such a super-shift band is not apparent with anti-c-myc antibody (lane 4). The band shift completely disappears on addition of the excess unlabeled TCF4 probe (lane 5), but not unlabeled NF{kappa}B probe (lane 6). (C) Gel retardation assay results for five HCC cell lines. The super-shift band is evident in not only the cell lines with ß-catenin and/or H-ras mutations, but also a cell line without the mutations.

 
Akt, GSK3ß and ERK1/2 phosphorylation and ß-catenin/cyclin D activation
After ß-catenin mutations were examined by the PCR–SSCP analysis using the paraffin-embedded tissues (data not shown), the corresponding frozen samples without the mutations were used for the western blot analysis of Akt, GSK3ß and ERK1/2. Histological examination demonstrated that five of these tumor samples contained both adenoma and HCC components with various ratio, whereas 13 and three samples were composed of adenoma and HCC, respectively. Immunohisotochemical studies revealed that the nuclear/cytoplasmic ß-catenin staining was correlated to the nuclear cyclin D1 index (Figure 5AGo) as described above. By western blot analysis, both total and phosphorylated Akt were detected in all hepatic tumors to the comparable levels in the normal livers (Figure 5BGo). On the other hand, total GSK3ß levels were variably elevated in 14 of 21 (66.6 %) tumors, and the phosphorylated form was increased in parallel with the total levels (Figure 5B and C, aGo). The expression levels of total ERK2 protein did not differ between the normal livers and hepatic tumors, but ERK1 levels were markedly elevated in the tumors than the normal livers. Although the phosphorylated ERK1/2 were detected in some tumors to the comparable levels with the normal liver, they were negative in other tumors. There was no significant correlation between the cyclin D1 expression levels and phosphorylation of Akt, GSK3ß and ERK1/2 (Figure 5C, b–eGo), although there was correlation between phosphorylation of ERK2 and the total or phosphorylated GSK3ß levels (Figure 5C, fGo).



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Fig. 5. (A) Results of immunohistochemical staining for ß-catenin and cyclin D1. N, normal livers; A, adenomas; A, H, adenomas and HCCs; H, HCCs. ß-Catenin staining pattern: M, membrane; C, cytoplasmic; N, nuclear. Cyclin D1 L.I.; the values above the bars represent the nuclear cyclin D1 L.I. in adenomas, and those under the bars represent the L.I. in HCCs. (B) Western blot analysis of Akt, GSK-3ß, ERK1/2 and cyclin D1. The total and phosphorylated Akt are almost equally expressed in the normal livers and hepatic tumors. GSK3ß levels are low in normal livers, but both total and phosphorylated forms are elevated in the tumors to various degrees. Although total ERK2 (42 kDa) levels are not different between the normal livers and hepatic tumors, ERK1 (44 kDa) levels are higher in the tumors. Phosphorylated ERK1/2 are detectable in the normal liver, but their levels are variable in the tumors. (C) There is significant correlation between the expression levels of phosphorylated and total GSK3ß (a). On the other hand, there is no correlation between cyclin D1 expression levels and phosphorylation of Akt (pAkt/total Akt) (b), GSK3ß (pGSK3ß/total GSK3ß) (c), ERK1 (pERK1/total ERK1) (d) and ERK2 (pERK2/total ERK2) (e) (R2 < 0.187), although there is correlation between phosphorylation of ERK2 (pERK2/total ERK2) and amounts of phosphorylated GSK3ß (f).

 

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The immunohistochemical study of ß-catenin/cyclin D1 and the ß-catenin/H-ras mutation analysis in 92 focal lesions demonstrated: (i) a significant correlation between the nuclear/cytoplasmic ß-catenin staining and ß-catenin mutations; (ii) a correlation between nuclear/cytoplasmic ß-catenin staining and high nuclear cyclin D1 L.I.; (iii) no correlation between H-ras and ß-catenin mutations; and (iv) no correlation between H-ras mutations and cyclin D1 expression. These results indicate that ß-catenin activation is associated with cyclin D1 over-expression, which may play an important role for progression of hepatic carcinogenesis, whereas H-ras mutations may be less responsible for this process. However, the fact that a few foci and adenomas showed relatively high cyclin D1 L.I. (10–20%) suggests that ß-catenin activation may not be a sole reason for the cyclin D1 activation.

An importance of ß-catenin activation for progression of hepatic carcinogenesis was further supported by the fact that all the HCC cell lines examined showed the ß-catenin/TCF4 complex and over-expression of ß-catenin and cyclin D1. These observations are in line with the published results that activated ß-catenin may bind to TCF4 and transcriptionally activate various genes including cyclin D1 in tumor cells (1315). In the absence of ß-catenin, TCF4 is bound to the transcription co-repressor Groucho and CtBP, and this complex acts as a transcription repressor of the TCF4 target genes (41). ß-Catenin competes with the co-repressors for TCF4 binding in a dose-dependent manner, and higher amounts create more active complexes, which could lead to more efficient de-repression. Thus, high nuclear ß-catenin is an important parameter determining the activity of ß-catenin/TCF4 complexes.

In the present study, although almost all HCC tissues showed nuclear/cytoplasmic ß-catenin staining, only 31.8% demonstrated ß-catenin mutations, indicating that ß-catenin may be activated by the mechanisms other than through mutations. One of the possible mechanisms for ß-catenin activation may be activation of the PI3K/Akt pathway, because GSK3ß is a target of the PI3K/Akt pathway, and its inactivation by phosphorylation by Akt has been reported to be associated with the increased accumulation and trascriptional activity of ß-catenin in HepG2 cells (28) and alveolar macrophages (29). Also, GSK3ß can phosphorylate cyclin D1 at threonine 286, which is important for degradation of cyclin D1 via the ubiquitin proteasome pathway (30). Therefore, GSK3ß inactivation by Akt may be causative of activation of both ß-catenin and cyclin D1. However, total and phosphorylated forms of Akt were not different between the normal livers and hepatic tumors, suggesting that the PI3K/Akt pathway may not be up-regulated in these tumors.

On the other hand, the GSK3ß levels were markedly elevated in ~50% of the tumors, and the phosphorylated form levels were almost comparable with the total levels, indicating that most GSK3ß was inactivated. Such an increase of GSK3ß in hepatic tumors, to our knowledge, was not reported previously. As GSK3ß inactivates various proteins that are involved in cell proliferation and survival, such as ß-catenin, cyclin D1, c-jun, c-myc, C/EBP and CREB (42), the GSK3ß inactivation results in activation of these proteins. Over-expression of catalytically active GSK3ß was reported to induce apoptosis of Rat-1 and PC12 cells, whereas dominant-negative GSK3ß prevented apoptosis (43). On the other hand, mice homologous for the deleted GSK3ß allele are embryonic lethal due to extensive hepatic cell death that was caused by the defect in TNF{alpha}-induced activation of NF{kappa}B (44). The abundant expression of GSK3ß then may provide the anti-apoptotic nature to hepatic tumor cells by modulating the NF{kappa}B pathway. However, the observation that there was no correlation between GSK3ß phosphorylation and cyclin D1 levels or nuclear/cytoplasmic ß-catenin staining indicated that, although the GSK3ß over-expression may have a role in the progression of hepatic tumors, it may be independent of the ß-catenin/cyclin D1 activation.

Although the mechanism(s) for ß-catenin activation was not clear in the majority of the present series of hepatic tumors, ß-catenin has recently been shown to be activated by various causes. First, the movement of ß-catenin from the cell membrane to the cytoplasm may be promoted by activation of growth factor receptors (2325) and ras gene products (26,27) by disrupting the adherens complexes by tyrosine phosphorylation of ß-catenin. Secondly, the fate of ß-catenin within the cytoplasm may be dependent on the formation of the Axin/APC/GSK3ß or APC/Siah-1 complexes that promote the degradation (10,11). Especially, dissociation of GSK3ß from the Axin/APC/GSK3ß complex through mutations of APC or Axin (10,11) or by activation of the Wnt pathway (45) may result in activation of ß-catenin. Actually, mutations of Axin were described in human HCC, which produce truncated Axin proteins with reduced binding to ß-catenin (38). Thirdly, the ß-catenin/TCF activity within nuclei may be regulated by various factors. TCF1 and lymphoid enhancer factor 1 (LEF-1) exist as both full length and short truncated forms, and, although the full length TCF1/LEF-1 can activate the gene transcription, the truncated forms that lack ß-catenin binding sites repress the ß-catenin/TCF activity (46,47). Moreover, the TGFß signaling may activate TAK-1, a member of the MAP-kinase family, that promotes the second kinase, NLK which phosphorylates TCF4, resulting in reduction of the binding of ß-catenin/TCF4 complexes to DNA (48). Finally, nuclear ß-catenin may be translocated from the nucleus back to the cell membrane, which may be mediated by E-cadherin (49,50). It is then possible that alterations of any of the factors that are involved in control of the ß-catenin activity may cause the ß-catenin activation. In this context, Igf1 and 2 are of particular interest, because Igf1 treatment has been shown to activate for ß-catenin to be translocated from cell membrane to cytoplasm (25) and to activate transcriptional activity of the ß-catenin/LEF complex through activation of the PI3K/Akt/GSK3ß and ras/MAPK pathways (25,28). Because Igf1 and 2 and Igf1 receptor may create the autocrine loop in mouse hepatic tumors (5153), its possible role in ß-catenin activation will be worth investigating.

H-ras mutations were detected only in foci and adenomas at low frequency, and there was no correlation with cyclin D1 expression or ß-catenin activation. This is clearly in contrast to the case of rat colonic tumors in which mutational K-ras activation was correlated to cyclin D1 over-expression (54), suggesting that the mechanism of cyclin D1 activation may vary depending on cell types. It is known that the ras activation may lead to sequential activation of raf-1, MEK and MAPK/ERK in many cellular systems, but in the present series of hepatic tumors, although the total ERK1 protein levels were higher in the tumors than the normal liver, phosphorylated ERK1/2 were not changed or rather lower in the tumors, and there was no correlation between ERK1/2 phosphorylation and cyclin D1 expression levels.

The presence of H-ras mutations in foci and adenomas but not HCCs argues that they were unnecessary for malignant transformation under the condition, which we used in this study. Although this observation is in contrast to the previous ones, it has been documented that the frequency of H-ras mutations is quite variable depending on kinds of carcinogens and on mouse strains (5,34,35). The facts that ras mutations were detected in various normal tissues such as normal human pancreas (55,56) and that transgenic mice with somatic activation of K-ras gene only showed higher incidence of lung tumors without increase of other tumors (57), indicate that additional complementing events may circumvent the oncogenic effects of ras activation. The discrepancy of the presence of H-ras mutations in the early rather than advanced lesions implies that other unknown genetic alteration(s) may be more important and therefore preferentially selected for progression of hepatic carcinogenesis, or H-ras activation may be incompatible with the malignant transformation like the case where the expression of activated H-ras gene resulted in cellular senescence rather than malignant transformation in mouse embryonic fibroblasts (58).

Although ß-catenin mutations were mostly detected in HCCs in the present study, other investigators have reported that they were recognized not only in late tumors but also in early lesions (5,7,9). Although much more early lesions are needed to be examined for the ß-catenin mutations, it is possible that there are multiple genetic pathways for hepatic carcinogenesis in which timing of ß-catenin mutations may vary. In this context, we should note that the ß-catenin mutation frequency has been reported to considerably differ depending on tumor types (8), inducing carcinogens (5) and transduced oncogenes in transgenic mouse models (2,9,59).

In conclusion, the ß-catenin/cyclin D activation was indicated to be important for progression for mouse hepatic carcinogenesis, but the Akt/GSK3ß and ras/ERK1/2 pathways seem to be of little significance in this process.


    Notes
 
2 To whom correspondence should be addressed Email: ogawak{at}asahikawa-med.ac.jp Back


    Acknowledgments
 
We thank Drs Susumu Tamakawa and Yuji Yaginuma, Department of Pathology, and Shinichi Chiba, Central Laboratory for Research and Education, Asahikawa Medical College for valuable discussion and technical support, and Yoshihisa Yamada and Miki Aihara, Tokushima Institute of New Drug Research, Otsuka Pharmaceutical Co. Ltd, Japan, for helpful advice regarding the gel retardation assay. We are also grateful to Dr Mari Iida, Biosafety Research Center, Foods, Drugs, and Pesticides for a gift of hepatic tumor specimens and valuable discussion.

This work was supported by a Grand-in Aid for Cancer Research from the Japanese Ministry of Health Welfare and Labour.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received December 6, 2002; revised December 15, 2002; accepted December 15, 2002.