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Short Communication |
From the Département de Pathologie*
and Service de
Chirurgie,
Hôpital Henri Mondor -
AP-HP, Créteil; and the Unité de Recombinaison et
Expression Génétique,
Institut
National de la Santé et de la Recherche Médicale U163,
Institut Pasteur, Paris, France
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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ß-catenin, like its homologue armadillo in Drosophila, is
an important multifunctional protein involved in cell-cell adhesion, by
strengthening the linkage of cadherin and 
-catenin to the actin
cytoskeleton.12
It is also involved in Wingless/Wnt
signaling during embryonic development13
and inappropriate
reactivation of this pathway has been implicated in tumorigenesis. In
the absence of Wnt signaling, ß-catenin is phosphorylated at
N-terminal serine-threonine residues by functional interactions with
glycogen synthase kinase (GSK)-3ß, axin and the adenomatous
polyposis coli protein (APC), and subsequently targeted to degradation
by the ubiquitin-proteasome system.14
Activation of the
Wnt signal inhibits GSK-3ß activity and induces ß-catenin
stabilization. Translocation of ß-catenin to the nucleus and its
association with high mobility group domain factors Tcf/LEF causes
transcriptional activation of target genes.15
Lately, the
c-myc, cyclin D1, and WISP genes have been reported to be
directly or indirectly activated by Wnt/ß-catenin signaling in human
epithelial cells.10,11,16
Recently, mutant ß-catenins that are resistant to down-regulation by GSK-3ß phosphorylation and ubiquitination have been characterized in human colorectal cancers and in a variety of carcinomas.17-19 In colon cancers, immunohistochemical studies have demonstrated increased expression of ß-catenin and its nuclear localization in tumors harboring either APC defects or ß-catenin mutations in the GSK-3ß phosphorylation domain.18,20 Nuclear and cytoplasmic localization of ß-catenin was also frequently seen in ovarian and uterine carcinomas and in melanomas, although genetic alterations of the ß-catenin gene could be detected only in a minority of these tumors.21-23 Whether inactivation of APC or other genetic or epigenetic event contributed to activating the Wnt/ß-catenin pathway in these cancers remains to be determined.
In human HCC, mutations of the ß-catenin gene have been reported in 19 to 26% of primary tumors,8,9 but immunolocalization of ß-catenin in tumoral tissues has not been investigated. The purpose of the present work was to determine the expression level and subcellular localization of ß-catenin in primary HCC specimens and in adjacent livers, to assess the relationship between nuclear accumulation and presence of ß-catenin mutations, and to investigate the effects of ß-catenin activation on cell proliferation. The expression of ß-catenin was also evaluated with respect to tumor size and grade and to patients' clinical follow-up.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Surgical liver resections for HCC from 32 patients (29 men, 3 women, mean age 59 years, range 36–78 years) were retrospectively studied. HCC was associated with cirrhosis or chronic hepatitis due to HCV (11 cases) or HBV infection (5 cases), to alcohol alone (9 cases) or in association with HCV infection (2 cases), or to genetic hemochromatosis (2 cases). Etiology was undetermined in one case. Two HCCs developed on normal liver. Twenty-two patients underwent segmental hepatectomy and 10 patients underwent orthotopic liver transplantation.
Fresh tissue samples from tumoral nodules and nontumoral livers were snap-frozen in liquid nitrogen and stored at -80°C for DNA studies.
For histopathological and immunohistochemical studies, tumor and liver specimens were fixed in 10% formalin, then embedded in paraffin. Four-micron sections were stained with hematoxylin-eosin-safran and with Masson's trichrome or picrosirius red for collagen. Tumor size, number of tumoral nodules, and presence of vascular invasion were determined at gross and/or microscopic examination.
DNA Analysis
DNA was extracted and purified by standard techniques from frozen tumoral and nontumoral liver samples.3 Genomic DNAs were amplified by a step-down polymerase chain reaction (PCR) protocol using 100 ng of template DNA, the forward primer 5'-GCGTGGACAATGGCTACTCAAG-3' (3' end of exon 2), and the reverse primer 5'-CTGGTCCTCGTCATTTAGC-3' (3' end of exon 4). PCR was performed in 50 µl of PCR buffer (Eurobio, Les Ulis, France) with 1.5 mmol/L MgCl2, 200 µmol/L dNTP, 1 µmol/L each primer, and 1 unit of Taq polymerase (Eurobio). The PCR conditions consisted of 1 cycle at 94°C for 4 minutes; 3 cycles of denaturation (94°C, 45 seconds), annealing (68°C, 45 seconds), and extension (72°C, 45 seconds); 3 cycles with annealing steps at 65, 62, 59, and 56°C; 20 cycles with annealing step at 52°C, and a final extension cycle at 72°C for 5 minutes in a Cyclogene thermal cycler (Techne, Cambridge, UK). Reaction products were resolved in 1% agarose and visualized with ethidium bromide. In two samples, shorter PCR products corresponding to deleted ß-catenin were excised from the gel and eluted. PCR products were sequenced using the T7 Sequenase PCR product sequencing kit (Amersham France, Les Ulis, France) according to the manufacturer's instructions. The primers for sequencing were the forward primer 5'-TGATGGAGTTGGACATGGCCATG-3' and the reverse primer 5'-CCCACTCATACAGGACTTGGGAGG-3' in exon 3.
Immunohistochemical Study
Analysis of ß-catenin and MIB-1 (Ki-67) expression was performed on serial sections for each tumor. For ß-catenin immunostaining, an immunoenzymatic method using alkaline phosphatase-anti-alkaline phosphatase (APAAP) complexes (Dakopatts, Trappes, France) and a ß-catenin monoclonal antibody (Transduction Laboratories, Lexington, KY) was used at a dilution of 1:500. For increased sensitivity and better subcellular localization of ß-catenin, an antigen retrieval method using microwave oven heating (2 x 5 minutes) in 1 mmol/L EDTA, pH 8.0,24 was applied on the deparaffinized and rehydrated sections. Slides were immersed in 20% AB human serum in 0.05 mol/L Tris-buffered saline, pH 7.6, and then incubated overnight at 4°C with the primary antibody. After washing, the slides were incubated for 30 minutes with a second rabbit anti-mouse antibody (Dakopatts) diluted 1:20, washed again, and incubated with the APAAP complexes at a 1:50 dilution. A single amplification was performed by adding sequentially the second antibody for 10 minutes and the APAAP complexes for 10 minutes. Sections were incubated with Fast Red salt-TR - Naphtol AX-TR phosphate (Sigma, Saint-Quentin Fallavier, France) in the presence of levamisole to block endogenous activity, then counterstained with aqueous hematoxylin and mounted in Immumount (Shandon, Cergy-Pontoise, France).
For Ki-67 immunostaining, deparaffinized sections were pretreated in 0.01 mol/L citrate buffer, pH 2.5. Immunolabeling, with the monoclonal antibody MIB-1 (Immunotech, Marseille, France) diluted 1:50, was performed using the immunohistochemistry automat Ventana NexES (Ventana Medical System, Strasbourg, France). This automat used the avidin-biotin peroxidase complex (ABC) method with 3,3-diaminobenzidine as chromogen and hematoxylin for counterstaining. As a control, no labeling was observed by omitting the primary antibody.
Evaluation of Immunostaining
The immunostaining of tumor samples was evaluated on coded slides on two different occasions. Membranous and intracytoplasmic ß-catenin signals were scored independently and graded in comparison with the adjacent nontumoral liver. In tumors showing nuclear expression, the percentage of strongly positive nuclei was determined by counting at least 1000 tumoral cell nuclei in homogeneous areas. For determination of the proliferative index, stromal cells were excluded and MIB-1-positive tumoral cell nuclei were counted on 10 consecutive high power (x400) fields.25
Statistical Analysis
Results of quantitative data in immunohistochemical analyses were expressed as mean ± one SE. Statistical differences between groups were tested with the non-parametric Mann-Whitney U test and the Fisher exact probability test. Statistical correlations between immunostaining counts were assessed by the Spearman rank correlation test. The significant level was defined as a P value <0.05.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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All tumors were HCCs grade I to III according to the World Health Organization international histological classification.26 Most tumors were single, although five were multinodular, and tumor sizes ranged from 1.6 to 19 cm (mean, 7.8 cm). Vascular invasion in portal vein and/or hepatic vein branches was observed in nine cases.
Nontumoral liver was normal in three, fibrotic in six, and cirrhotic in 23 cases.
Identification of ß-Catenin Mutations
Thirty-five HCC specimens from 32 patients were screened for
mutations in the ß-catenin gene by PCR amplification of genomic DNA
over intron 2 to exon 4 sequences. The wild-type 1.1-kbp PCR product
was present in all tumors, and additional, faster migrating bands were
detected in two cases (Figure 1A)
.
Subsequent DNA sequencing of these products revealed interstitial
deletions of 553 bp in Patient 11 and 595 bp in Patient 12,
encompassing in both cases most of exon 3 and 4 sequences of the
ß-catenin gene (Figure 1B
and Table 1
).
In the remaining tumors, missense mutations affecting ß-catenin
residues 32, 33, 34, 35, 36, 40, 41, 45, or 47 were detected in 10
patients (Figure 1D
and Table 1
). It is noteworthy that these changes
affected not only all four GSK-3ß phosphorylation sites and flanking
residues potentially implicated in ß-catenin ubiquitination, but also
several residues (H36, T40, and S47) that were not found to have
mutated in previous studies of various neoplasms.19
In
Patient 2, among four HCC nodules in the explant liver, only one
displayed a mutation changing amino acid 36 from a histidine to a
proline. Tumors 7 and 9 are also interesting in that they carried more
than one point mutation in exon 3. For Patient 7, in addition to a
silent mutation at codon 34, mutations changing isoleucine 35 to an
asparagine and threonine 40 to an alanine were demonstrated (Figure 1C)
, and Patient 9 harbored the classical threonine 41-to-alanine
mutation together with a change of serine 47 to an arginine.
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) might be coincidental, and larger studies are needed to
answer this question. ß-Catenin Immunostaining
In nontumoral livers, a thin membranous ß-catenin signal
delineated the hepatocytes. Strong membranous and pale cytoplasmic
staining of bile ductules, as well as milder staining of nerves and
endothelial cells, were observed (Figure 2A)
. Normal hepatocytes or dysplastic
cells from cirrhotic nodules or from livers with chronic hepatitis did
not display nuclear expression of ß-catenin. In 20 of the 35 tumors
analyzed (57%), membranous ß-catenin immunostaining was markedly
increased by comparison with adjacent livers and was frequently
associated with intracytoplasmic staining (data not shown). Nuclear
staining could be observed in 23 of the 35 HCC nodules (66%). In eight
of these nodules, only very scarce (<1%) tumoral cells displayed
ß-catenin positivity in the nucleus. In the other 15 cases, a
strongly positive nuclear signal could be observed in 8 to 90% of
tumoral nuclei (Figure 2B)
. ß-catenin gene mutations were detected in
nine of these 15 cases (Table 1)
. The mean rate of ß-catenin-positive
nuclei was significantly higher in HCCs carrying a mutated form of
ß-catenin than in HCCs without mutation (49.6 ± 11.4%
versus 15.5 ± 6%, P < 0.005,
Mann-Whitney test). Conversely, the number of HCCs with ß-catenin
mutation was significantly higher in the group of tumors harboring
nuclear immunostaining for ß-catenin in 8 to 90% of cells than in
the group of tumors without ß-catenin nuclear positivity (Table 2)
.
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) showed only membranous ß-catenin staining, with
lower or equal intensity compared to corresponding nontumoral liver. On
the other hand, four tumors in which ß-catenin gene alteration could
not be detected showed 30 to 80% of nuclei strongly positive for
ß-catenin staining. Further studies are required to determine whether
these results reflect tumor heterogeneity, because different specimens
were used for DNA analysis and for immunostaining. Finally, ß-catenin
expression was markedly elevated at the invasive front in three tumors
showing otherwise moderate staining and in the tumoral intravascular
compartment, compared with other tumor areas in 4 of the 9 cases of
vascular invasion (Figure 2C)
. Tumor Cell Proliferation
Proliferative index, determined by the count of tumoral nuclei
that scored positive after immunostaining with a Ki-67 specific
antibody, was twofold to more than 60-fold higher in tumors than in the
nontumoral liver counterpart; the mean proliferative index was 498.5
(range, 12–1,854) in tumors versus 18.1 (range, 0–100) in
nontumoral liver. A higher number of positive cells was observed at the
invasive front as well as in tumoral vascular invasion (Figure 2D)
. In
some heterogeneous tumors, areas showing markedly elevated nuclear
expression of ß-catenin also exhibited increased number of Ki-67
positive cells (Figure 2, E and F)
. However, in four cases, the
proliferative index was high (from 587 to 1,500 for 10 high power
fields) in the absence of nuclear staining for ß-catenin (data not
shown).
Statistical analysis demonstrated a strong correlation between the rate
at which cells expressed ß-catenin in the nucleus and the number of
Ki-67-positive cells (P < 0.001, Spearman
test). When tumors were divided in two groups according to nuclear
ß-catenin staining, ie, from 8 to 90% of positive cells
versus no nuclear positivity, a significant relationship
between the two factors persisted (P < 0.01,
Mann-Whitney test; Table 2
). In contrast, no significant relationship
could be established between the ß-catenin gene status and the
proliferative index.
Correlation with Clinical Course and Disease Status
Clinicopathological parameters were collected for all 32 patients
with a median follow-up of 20 months (range, 8–72 months). Recurrence
of HCC occurred in 11 patients (within one year after surgery in 9 of
them) and was responsible for death in 6 of these 9 cases. In two
cases, recurrence was detected at the 24th month. Two patients for whom
follow-up was shorter than 12 months and seven patients who died in the
immediate postoperative period were excluded from statistical analysis.
Although neither survival nor recurrence rates could be definitely
established because of the short mean follow-up time, statistical
analysis suggested a strong correlation between high cell proliferation
index in the primary tumor and poor survival. Indeed, patients who died
by recurrence (n = 6) had a much higher
proliferative index in resected primary HCC than patients still alive
(n = 15; mean value 1,135.67 ± 304.32
versus 288.07 ± 77.78, P < 0.02).
High proliferation index in the primary tumor also correlated with
tumor recurrence (Table 3)
. In addition,
recurrence was related to a high rate of cells expressing ß-catenin
in their nuclei; tumor recurrence was observed in 10 cases in which the
mean rate of positive nuclei for ß-catenin was high, whereas 11
patients harboring primary HCCs with lower rate of ß-catenin-positive
nuclei remained free of disease (Table 3)
. These results need to be
confirmed by a longer follow-up.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our results, showing a strong correlation between nuclear ß-catenin staining and somatic mutations of the ß-catenin gene in HCC, indicate that activation of the Wnt/ß-catenin pathway in HCC occurs predominantly through activating mutations in the ß-catenin gene itself. It differs from colorectal cancers, in which APC mutations are responsible for ß-catenin stabilization in 80% of the cases,29 and from melanoma or ovarian and uterine carcinomas, in which very occasional mutations can be found despite frequent ß-catenin overexpression.21-23 In our study, 34% of tumor samples had mutations or deletions in the third exon of the ß-catenin gene. Similar mutation rates were reported in previous studies of European and Asian patients with HCC, albeit with some differences in the distribution of mutated amino acids.8,9 Further studies of the functional role of these different mutations on ß-catenin stability might be important to better define the mechanisms involved in the posttranscriptional control of ß-catenin expression. In addition, intense nuclear ß-catenin expression in HCCs carrying apparently normal ß-catenin alleles (11% of the cases) might be associated with alterations of the ß-catenin gene in a region other than exon 3 or with defects in the APC gene, although the very low prevalence of allelic losses at the APC locus on chromosome 5q21 in HCC does not favor the latter hypothesis.3,30 Another possible mechanism is the involvement of other factors mediating Wnt signaling, like the human frizzled (Fz) genes. Indeed, a novel Fz gene (FzE3) has recently been shown to enhance ß-catenin signals in human esophageal carcinomas.31
Nuclear translocation of ß-catenin and its association with Tcf/Lef-1 factors can activate gene expression and cell proliferation in experimental systems.14 The present study revealed a significant relationship between the number of tumoral hepatocytes with nuclear ß-catenin and the number of Ki-67-positive cells. To the best of our knowledge, this is the first in vivo study demonstrating a link between the stabilization and nuclear accumulation of ß-catenin and the proliferation of tumor cells. Whether c-myc or other target genes of Wnt/ß-catenin signaling play a direct role in this process is an important issue. However, our data showing a very high proliferative index in four HCC cases in the absence of ß-catenin mutation and nuclear expression, provide evidence implicating other mechanisms in active tumor cell proliferation. It is of interest that high cell proliferation in the primary tumor has been closely related to poor survival and tumor recurrence in HCC patients in previous reports,32,33 as also suggested in the present study. In addition, our preliminary data seem to indicate that nuclear accumulation of mutated ß-catenin in HCC might be associated with an increased risk of tumor recurrence and poor survival. However, the mean follow-up was short in the present work; larger studies with longer follow-up are clearly needed to confirm these data and to determine whether oncogenic activation of ß-catenin and active cell proliferation represent independent prognostic factors in HCC.
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Acknowledgements |
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Footnotes |
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Accepted for publication May 17, 1999.
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References |
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- and ß-catenins in nontumoral livers and hepatocellular carcinoma. Hepatology 1996, 23:1441-1447[Medline]
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