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TITLE PAGE
The ß-catenin-target Fascin-1, altering hepatocyte differentiation, is a new
marker of immature cells in hepatoblastomas
Short title: Fascin-1 alters hepatocyte differentiation
Caroline Gest1,#, Sandra Sena1,#, Véronique Neaud1, Robin Loesch2, Nathalie
Dugot-Senant3, Lisa Paysan1, Léo Piquet1, Terezinha Robbe1, Nathalie Allain1,
Doulaye Dembele4, Catherine Guettier5, Paulette Bioulac-Sage1, Brigitte Le Bail1,
Christophe F. Grosset6, Frédéric Saltel1, Valérie Lagrée1, Sabine Colnot2, and
Violaine Moreau1
1 Univ. Bordeaux, INSERM, BaRITOn, U1053, F-33000 Bordeaux, France; 2
INSERM, Sorbonne Université, Université de Paris, Centre de Recherche des
Cordeliers (CRC), Paris, France; 3 Plateforme d'histologie, UMS 005, Bordeaux F-
33076, France; 4 IGBMC, CNRS UMR 7104 - INSERM U 1258 - Université de
Strasbourg, 67400, Illkirch, France; 5 Department of Pathology, Bicêtre University
Hospital, University of Paris-Saclay, Assistance Publique-Hôpitaux de Paris, Le
Kremlin-Bicêtre, F-94275, France; 6 MIRCADE team, Univ. Bordeaux, Inserm,
BMGIC, U1035, Bordeaux, F-33000, France.
# Authors share co-first authorship
Grant support: CG and SS were supported by postdoctoral fellowships from,
respectively, La Ligue Nationale contre le Cancer and the Fondation pour la
Recherche Médicale. L. Piquet and L. Paysan were supported by PhD fellowships
from respectively the SIRIC BRIO and the Région Aquitaine. This work was
supported by grants from La Ligue contre le Cancer (comité régional) (to VL), La
Ligue Nationale contre le Cancer “Equipe labellisée 2016” (to VM and FS), and from
Institut National du Cancer (PLBIO-INCa2014-182) (to VM).
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Abbreviations: BC, bile canaliculus; CFDA, 5-carboxyfluorescein diacetate; HB,
hepatoblastoma; HCC, hepatocellular carcinoma; HNF4a, hepatocyte nuclear factor-
4 alpha; KD, knock-down; STED, stimulated emission depletion; WT, wild-type.
Contact Information: Violaine Moreau, 146 Rue Léo Saignat, F-33076 Bordeaux,
+33 5 57 57 12 72, violaine.moreau@inserm.fr
Disclosures: The authors disclose no conflicts.
Transcript Profiling: The transcriptomic data are archived in the public GEO data
repository under the GEO accession number GSE144107.
Author Contributions: Study design: VM, Generation of experimental data: CG, SS,
VN, LP, LP, TR, NDS, RL, NAC, DD, VL. Analysis and interpretation of data: CG, SS,
DD, PBS, CFG, BLB, VL, SC, VM. Drafting of the manuscript: FS, VL, SB, VM.
Data Transparency Statement: study materials will be made available to other
researchers upon request.
Acknowledgements
We thank Dr D. Vignjevic (Curie Institute, Paris, France) for Fascin-1 DNA
constructs. We thank the plateforme GenomEast of Strasbourg (Strasbourg, France).
Microscopy was done in the Bordeaux Imaging Center, a service unit of the CNRS-
INSERM and Bordeaux University, member of the national infrastructure France Bio
Imaging, with the help of Dr. Philippe Legros for STED analysis. We thank Anne-
Aurélie Raymond (Inserm U1053, Bordeaux, France) for her help in transcriptomic
analysis.
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ABSTRACT
BACKGROUND & AIMS: ß-catenin is a well-known effector of the Wnt pathway and
a key player in cadherin-mediated cell adhesion. Oncogenic mutations of ß-catenin
are highly frequent in pediatric liver primary tumors. Those mutations are mostly
heterozygous allowing the co-expression of wild-type (WT) and mutated ß-catenins in
tumor cells. We investigated the interplay between WT and mutated ß-catenins in
liver tumor cells, and searched for new actors of the ß-catenin pathway. METHODS:
Using an RNAi strategy in ß-catenin-mutated hepatoblastoma (HB) cells, we
dissociated the structural and transcriptional activities of β-catenin, carried mainly by,
respectively, WT and mutated proteins. Their impact was characterized using
transcriptomic and functional analyses. We studied mice that develop liver tumors
upon activation of ß-catenin in hepatocytes (APCKO and ß-catenin∆exon3 mice). We
made use of transcriptomic data from mouse and human HB specimens and
analyzed samples by immunohistochemistry. RESULTS: We highlighted an
antagonist role of WT and mutated ß-catenins on hepatocyte differentiation as
attested by alteration of hepatocyte markers expression and bile canaliculi formation.
We characterized Fascin-1 as a target of ß-catenin involved in hepatocyte
differentiation. Using mouse models that allow the formation of two phenotypically
distinct tumors (differentiated or undifferentiated), we found that Fascin-1 expression
is higher in undifferentiated tumors. Finally, we found that Fascin-1 is a specific
marker of the embryonal component in human HBs. CONCLUSIONS: In mice and
human, Fascin-1 expression is linked to loss of differentiation and polarity of
hepatocytes. Thus, we highlighted Fascin-1 as a new player in the modulation of
hepatocyte differentiation associated to ß-catenin pathway alteration in the liver.
Keywords: hepatoblastoma; differentiation; beta-Catenin; Fascin-1
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INTRODUCTION
ß-catenin is an evolutionary conserved protein that plays a dual role in cells. It is the
key effector of the canonical Wnt pathway, acting as a transcriptional cofactor in
complex with lymphoid enhancer factor/T-cell factor (LEF/TCF)1. In addition, ß-
catenin plays a central role in cadherin-mediated cell adhesion. In epithelial cells, in
the absence of Wnt signaling, ß-catenin is associated with E-cadherin at cell-cell
junctions, and β-catenin is maintained at low cytoplasmic levels through its
destruction complex of Axin, adenomatosis polyposis coli (APC), and glycogen
synthase 3ß kinase (GSK-3ß). GSK-3ß phosphorylates ß-catenin causing its
degradation by the proteasome. Upon Wnt signal, the destruction complex is
disrupted and cytoplasmic stabilized ß-catenin translocates in the nucleus where it
drives the transcription of target genes. Thus, ß-catenin is endowed with two main
functions, a structural function at cell-cell adhesion and a transcriptional function in
the nucleus. Imbalance between signaling properties of ß-catenin may lead to
deregulated cell growth, adhesion and migration resulting in disease such as tumor
development and metastasis. However, given the dual function of ß-catenin, it is
difficult to distinguish which function, structural versus transcriptional, of ß-catenin, is
precisely involved in cellular processes.
In the liver, the Wnt/ß-catenin pathway plays important roles regulating embryonic
and postnatal development, zonation, metabolism and regeneration2. This pathway is
also strongly involved in the hepatocarcinogenesis. Its aberrant activation, due to
mutations in the CTNNB1 gene encoding ß-catenin or in components of the
degradation complex, such as AXIN and APC, are detected in primary liver cancers.
ß-catenin gene alterations are identified in up to 80% of human hepatoblastomas
(HBs), the primary hepatic malignancy in children, and in 30 to 40% of human
hepatocellular carcinomas (HCCs)3, 4. Moreover, whereas
APC
loss-of function
mutations remain rare in HCCs, they are more frequent in HBs, associated to familial
or sporadic cases5. Even if HBs remain poorly studied and complex tumors with
various histologic components (epithelial, mesenchymal, fetal, and embryonal) within
the same tumor6, two subtypes were described based on molecular analyses; the
fetal C1 subtype with favorable outcome shows enhanced membranous staining and
cytoplasmic accumulation of ß-catenin with occasional nuclear localization, whereas
the proliferative poorly differentiated C2 subtype is characterized by an intense
nuclear staining of ß-catenin3. The C2 subtype was recently split into subgroups,
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C2A and C2B, with C2A subgroup containing more proliferative tumors7. The majority
of ß-catenin mutations affect exon3 at sites of phosphorylation by GSK3ß, avoiding
its degradation and so constitutively activating the Wnt pathway. Interstitial deletions
in the third exon of the ß-catenin gene are highly prevalent in HBs, while point
mutations are more common in HCCs8. Interestingly, most of mutations in exon3 are
monoallelic mutations, leaving a wild-type (WT), non-mutated allele in tumor cells.
We thus attempt to address the interplay between the WT and the mutated ß-
catenins in liver tumor cells.
To do so, we made use of the human HB HepG2 cell line. HepG2 cells have an
heterozygous deletion of 348 nucleotides in exon3 of the CTNNB1 gene, resulting in
an abundant truncated form of ß-catenin and a lower amount of WT ß-catenin4. The
large deletion (amino acid residues 25–140) removes the GSK-3ß phosphorylation
sites and the binding site for α-catenin, the ß-catenin partner in the E-cadherin-
mediated cell adhesion. Thus, both WT and mutated (∆aa25-140) forms of ß-catenin
co-exist in these cells and the interplay between both has never been explored. In
addition, HepG2 cells exhibit a good degree of differentiation and show most cellular
features of normal human hepatocytes such as bile canaliculi (BC) formation9. We
thus designed an RNA interference approach to specifically knockdown WT and/or
mutated ß-catenin and address their reciprocal role in hepatocyte differentiation.
Using this model, we dissociated the structural and transcriptional activities of β-
catenin, carried mainly by, respectively, the WT and the mutated proteins. Moreover,
we found that whereas knockdown (KD) of WT ß-catenin induced a loss of BC, KD of
mutated ß-catenin lead to an increase of their number and size, suggesting that both
play antagonistic role in tumor hepatic cell differentiation. Moreover, molecular
characterization revealed that FSCN1 (fascin-1) is a target of ß-catenin involved in
the differentiation state of hepatocytes. We further found that fascin-1 expression is
associated to undifferentiated ß-catenin mutated tumors in mice, which are closed to
human HBs. Using human samples, we found that fascin-1 is specifically expressed
in the embryonal component of HBs. Thus, we described FSCN1 as a ß-catenin
target gene associated with hepatic tumors of poor outcome, such as poorly
differentiated HBs.
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MATERIALS AND METHODS
Cell culture
Human HB cell line, HepG2, and human HCC cell lines, Hep3B, Huh7 and SNU398,
were purchased from American Type Culture Collection. HB cell line Huh6 was
generously provided by C. Perret (Paris, France). All cell lines were cultured as
previously described10, 11.
Transfection
DNA and siRNA transfections were realized as described previously11. pEGFP-
Fascin-1 vector was a generous gift from Dr D. Vignjevic (Paris, France). SiRNA
were purchased from Eurofins genomics. Sißcat WT targets human ß-catenin mRNA
at 5’-GTAGCTGATATTGATGGACAG-3’, sißcat both at 5’-
ACCAGTTGTGGTTAAGCTCTT-3’ and sißcat mut at 5’-
TGTTAGTCACAACTATCAAGA-3’. SiFascin-1#1 targets human Fascin-1 mRNA at
5’-CAGCTGCTACTTTGACATCGA-3’, siFascin-1#2 at 5’-
CAAAGACTCCACAGGCAAA-3’, siFascin-1#3 and siFascin-1#4 were purchased at
Ambion. The AllStars negative-control siRNA from Qiagen was used as control
siRNA. For KD, a reverse transfection was performed on day 1, a second forward
transfection on day 3 and experiments were performed on day 5.
Luciferase reporter assays
Assays were performed as previously described11. pGL4-TOP reporter with TCF
responsive elements to quantify ß-catenin transcriptional activity was a generous gift
from Pr. H. Clevers (Utrecht, The Netherlands). The pmFascin-Luciferase vector
containing luciferase under fascin promoter was a generous gift from Dr D. Vignjevic
(Paris, France).
Western blot
Cells were scraped off on ice and homogenized in RIPA buffer (150 mM NaCl, 0.1%
Triton X‐100, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate, and 50 mM
Tris‐HCl pH 8.0) containing protease and phosphatase inhibitors cocktail (Thermo
Scientific). Cell lysates were cleared of cellular debris and nuclei by a 16,000 × g
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centrifugation step for 10 min. Lysates were analyzed using a western-blot protocol
described previously11. The antibodies used are listed in Table S1.
Quantitative real-time PCR
RNA was collected from cultured cells using the Trizol reagent (Invitrogen), according
to manufacturer’s protocol. Reverse transcription and SYBR® Green-based real-time
PCR were performed as described previously11. Gene expression results were first
normalized to internal control r18S. Relative levels of expression were calculated
using the comparative (2-ΔΔCT) method. All primers used for qRT-PCR experiments
are listed in Table S2.
Immunofluorescence
Glass coverslip-plated cells were prepared for immunofluorescence microscopy and
imaged as previously described12. The antibodies used are listed in Table S1. F-actin
was stained using fluorescent phalloidin (1/250) (Molecular Probes). To detect
functional BC, cells were incubated in PBS with 5-carboxyfluorescein diacetate
(CFDA) (Sigma) at a final concentration of 0.5 µM for 30 min at 37°C to allow its
internalization into the lumen. Cells were washed three times with PBS and CFDA
positive BC were counted as functional under a fluorescence microscope. Stimulated
emission depletion (STED) microscopy was used to image microvilli of BC, by
labelling actin with phalloidin-Atto647N (Thermo Fischer Scientific). Sample was
imaged with an inverted Leica SP8 STED microscope equipped with an oil immersion
objective (Plan Apo 100X NA 1.4), white light laser (WLL2) and internal hybrid
detectors. BC features were quantified using image J on STED images.
Quantification of BC was assessed in three independent experiments in which at
least 100 cells were counted.
Cell growth assay
For 2D assay, cells were seeded at 5000 cells/well in 96-well plates and grown for 1
to 6 days. Each assay was performed in five replicates. For indicated time points,
cells were fixed with 50% trichloroacetic acid at 4°C for 1 hour, and processed using
the SulfoRhodamine B assay kit (Sigma).
Mouse samples
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We collected tumoral and non tumoral livers from mouse transgenic models with
hepatic β-catenin activation. All animal procedures were approved by the ethical
committee of Université de Paris according to the French government regulation. The
Apcfs-ex15 and β-cateninΔex3 mouse tumors were obtained from compound
Apcflox/flox/TTR-CreTam and β-cateninex3-flox/flox respectively, injected with 0.75 mg
Tamoxifen or with 5 x 108 ip Cre-expressing adenovirus, as previously described13-15.
All the mice were maintained at the animal facility with standard diet and housing.
They were followed by ultrasonography every month until tumor detection, thereafter
ultrasound imaging was continued every 2 weeks. Table S3 described the cohort of
mice used for this study.
Patient samples
Liver tissues were immediately frozen in Isopentane with Snapfrost and stored at
−80°C until used for molecular studies. Samples were obtained from the Centre de
Ressources Biologiques (CRB)-Paris-Sud (BRIF N°BB-0033-00089) with written
informed consent, and the study protocol was approved by the French Government
and the ethics committees of HEPATOBIO (HEPATOBIO project: CPP N°CO-15-
003; CNIL N°915640). Liver samples were clinically, histologically, and genetically
characterized (Table S4). Among the 20 cases, 12 were classified as C1, 3 as C2A
and 5 as C2B in a C. Grosset’s previous study7.
Immunohistochemistry
Specimens were fixed in buffered formaldehyde. The 2.5 µm thick sections were
dewaxed and rehydrated and antigen retrieval was performed in a Tris-EDTA buffer
(pH9 solution for 20 min). All staining procedures were performed in an automated
autostainer (Dako-Agilent Clara, United States) using standard reagents provided by
the manufacturer. Endogenous peroxidase was inhibited with 3% H2O2 in H2O, and
non-specific sites were saturated with 20% goat serum in TBS-Tween. The sections
were incubated with the anti-Fascin-1 monoclonal antibody at 2 µg/mL for 45 min at
room temperature. EnVision Flex/HRP was used for signal amplification. 3,3’-
Diamino-benzidine development was used for detecting primary antibodies. The
slides were counterstained with hematoxylin, dehydrated and mounted.
Statistical tests
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Data were reported as the mean ± SEM of at least three experiments. Statistical
significance (P < 0.05 or less) was determined using a Student’s t-test or analysis of
variance (ANOVA) as appropriate and performed with GraphPad Prism software. P
values are indicated as such: * P < 0.05; ** P < 0.01; *** P < 0.001; **** P <0.0001;
ns, non significant.
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RESULTS
The dual ß-catenin knocked-down HepG2 model
The N-terminus part of ß-catenin contains the GSK-3ß phosphorylation sites that are
crucial for the regulation of its turnover. This region encoded by the exon3 of ß-
catenin is deleted on one allele of the CTNNB1 gene in HepG2 cells4. Thus, a high
amount of a truncated form of 76kDa is co-expressed with a 92kDa full-length ß-
catenin (Fig. 1). We designed small interfering RNAs (siRNAs), named “sißcat-WT”,
“sißcat-mut” and “sißcat-both” to target, respectively and specifically, either the WT,
the mutated form of ß-catenin, or both (Fig. 1A). All three siRNAs efficiently knocked-
down the protein expression of their targeted form of ß-catenin (Fig. 1B and S1A). At
the mRNA level, we found that sißcat-WT and sißcat-mut decreased the amount of
ß-catenin transcripts by two fold and that, as expected, sißcat-both led to almost a
full extinction of ß-catenin in HepG2 cells (Fig. 1C).
We then characterized this model by analyzing the transcriptional activity of ß-catenin
upon silencing of both alleles. Using the TOP-flash reporter system, we found that
whereas silencing the WT allele did not impact TCF/LEF reporter activity, silencing
the expression of the mutated allele or both alleles strongly inhibited it (Fig. 1D). We
further showed that expression of positive targets of ß-catenin, such as GPR49,
Axin2 and cyclinD1 were strongly inhibited upon treatment with sißcat-mut and
sißcat-both (Fig. 1E). Consistently, the reverse result was obtained for negative
targets of ß-catenin such as Arg1 (Fig. 1E). Thus, the data suggest that, in HepG2
cells, the TCF-dependent transcriptional activity of ß-catenin is mainly carried by the
mutated form of ß-catenin.
According to this impact on ß-catenin targets, we found that HepG2 cell growth was
highly dependent on mutated ß-catenin expression, whereas it was insensitive to the
KD of the WT form of ß-catenin (Fig. 1F). This data obtained in 2D culture, were also
confirmed on spheroids (Fig. S1B), showing that the mutated ß-catenin is required for
HepG2 cell growth in a 2D and 3D environment. This alteration of cell growth
correlated with the down-expression of cyclinD1 upon mutated ß-catenin KD (Fig.
1E). Thus, these results confirm that the mutated form of ß-catenin acts as an
oncogene independently of the WT ß-catenin, and consistent with the notion of
oncogene addiction, this allele is strictly required for HepG2 cell growth.
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The large deletion (residues 25–140) present in HepG2 cells also removes α-catenin
binding site of ß-catenin, involved in the formation of E-cadherin-based adherens
junctions. We thus found that the silencing of the mutated allele of ß-catenin did not
impact E-cadherin localization at cell-cell junctions. Interestingly, in an opposite
manner, WT ß-catenin KD strongly affected E-cadherin engagement at adherens
junctions in HepG2 cells (Fig. 2A). In cells lacking the expression of both ß-catenins,
E-cadherin staining appears strongly affected with a more intracellular localization.
Thus, these results suggested that the structural role of ß-catenin is altered only
upon WT ß-catenin KD. Thus, this dual ß-catenin KD HepG2 model allows the
uncoupling of the two functions of ß-catenin in the same cellular background:
membrane/structural activity mediated by the degradable WT ß-catenin and the
nuclear/transcriptional activity mediated by the mutated β-catenin. This conclusion is
further supported by ß-catenin localization (Fig. 2B). In control HepG2 cells, ß-
catenin localized at cell-cell junctions, in the cytoplasm and in nuclei, stainings which
are lost when cells are transfected with sißcat-both. In cells transiently transfected
with sißcat-mut, remaining ß-catenin, i.e. WT ß-catenin, is enriched at cell-cell
junctions, but failed to localize in the cytoplasm and in nuclei. At the opposite, in cells
transfected with sißcat-WT, remaining ß-catenin, i.e. mutated ß-catenin, is still
cytosolic and less at the membrane (Fig. 2B). These results suggest that, in HepG2
cells, WT β-catenin is mainly involved in adherent junctions whereas mutated β-
catenin is preferentially involved in the regulation of gene expression. Based on
these results, we believe that this dual ß-catenin KD HepG2 model is suitable to
address independently the structural and the transcriptional functions of ß-catenin.
WT and mutated ß-catenin have distinct gene expression patterns
To identify the impact of the silencing of each ß-catenin allele on gene expression,
we performed a transcriptional analysis of HepG2 KD cells (Tables S5-7). To validate
the data, we first checked the expression of ß-catenin and known-ß-catenin targets
(Fig. S2A); they were altered in a similar way than previously observed by qRT-PCR
(Fig. 1C and 1E). Global analysis demonstrated that WT ß-catenin KD cells and
mutated ß-catenin KD cells have distinct gene expression patterns, and silencing of
both alleles led to a gene expression pattern closer to the mutated than the WT ß-
catenin KD, suggesting a dominant impact of the oncogenic ß-catenin on gene
expression in HepG2 cells (Fig. S2B). Venn diagram revealed that only a small
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proportion of genes are altered in common upon silencing of WT or mutated ß-
Catenin (Fig. S2C). We further used FuncAssociate 3.0 to analyze alterations in
pathway and biological functions (Table S8). We found that removal of the WT ß-
catenin led to a decrease in expression of genes involved in metabolic processes. It
was striking to observe an opposite behavior upon removal of the oncogenic ß-
catenin. We also found a cell cycle and a TGF-ß signature for genes that were down-
regulated upon silencing of both or of the mutated allele. As metabolic functions are
key features of differentiated hepatocytes, these alterations led us to explore the
impact of both ß-catenin functions on cell differentiation.
Alteration of the hepatocyte differentiation and polarity upon ß-catenin knock-
downs
Differentiated hepatocytes are characterized by the expression of specific markers,
including xenobiotic-metabolizing enzymes, transporters, transcription factors and
bile canaliculi molecules. Interestingly, we observed an alteration in mirror upon
silencing of either the WT or the mutated allele of ß-catenin. Hepatocyte markers
were found up-regulated upon mutated ß-catenin silencing and down-regulated upon
WT ß-catenin silencing (Fig. 3A). As HNF4a (hepatocyte nuclear factor-4 alpha) is a
transcription factor involved in hepatocyte differentiation, we further explored the
impact of ß-catenin KD on HNF4a signaling. We first analyzed the expression of
HNF4a by qRT-PCR and found a slight decrease of its expression upon WT ß-
catenin KD, whereas it is not significantly altered upon silencing of the oncogenic ß-
catenin (Fig. 3B). We found a significant increase of HNF4a transcriptional activity
upon silencing of the oncogenic ß-catenin using ApoC3 promoter reporter assay (Fig.
3C). Accordingly, in our transcriptional analysis, expressions of transcriptional
positive targets of HNF4a were found upregulated upon silencing of the mutated ß-
catenin (Fig. S3 A-C) confirming the results obtained for ApoC3 and ApoM mRNA
expressions by qRT-PCR (Fig. 3D). In both assays, the removal of WT ß-catenin
slightly decreased the expression of HNF4a target genes. As described earlier16, this
result confirms that transcriptional activity of ß-catenin may repress the hepatocyte
differentiation program of HNF4a. Altogether, these results suggest that the structural
function of ß-catenin, mainly supported by the WT ß-catenin, is necessary to
maintain a differentiated state of hepatocytes, and that the inhibition of the
transcriptional activity of oncogenic ß-catenin reverses the dedifferentiation program
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of HepG2 cells.
As differentiated hepatocytes are polarized epithelial cells endowed with the capacity
to produce and excrete bile into a specialized structure called bile canaliculus (BC),
we also analyzed the impact of ß-catenin KD on BC markers. As described above for
hepatocyte markers, we observed an alteration in mirror in the expression of junction-
and polarity-associated genes (Fig. S3D-E), such as JAM-A (F11R gene), connexin-
32 (GJB1 gene) and claudin-1 (CLDN1 gene) (Fig. 3E, S3D-E). As HepG2 cells
retained the ability to form BC in culture, we addressed the impact of ß-catenin on
their maintenance using confocal microscopy by staining F-actin and radixin (Fig.
4A). Interestingly, we found that depletion of WT β-catenin induced a decrease,
whereas depletion of mutated β-catenin an increase, in the number of cells exhibiting
BC (Fig. 4A-B). We also performed super-resolution STED microscopy on HepG2
cells stained with phalloidin in order to visualize BC with an improved resolution than
confocal (Fig. 4C and Fig. S4A). Quantification of BC features demonstrated that
mutated ß-catenin KD increases their size, as attested by the increase of their area,
perimeter and diameter, whereas their circularity remains unchanged (Fig. 4D), and
the number of cells engaged in the formation of each canaliculus was found higher
(Fig. S4B). Finally, to check the functionality of BC, we examined their ability to
translocate CFDA, a fluorescent substrate of apical plasma membrane ABC
transporters, into the apical lumen. As shown between adjacent HepG2 cells on
Figure 4E, removing the mutated form of ß-catenin increased the percentage of
CFDA positive BC, demonstrating they are fully functional (Fig. 4E). Our results
demonstrated that in tumor hepatocytes, WT and mutated ß-catenin act
antagonistically on hepatocyte BC. These results suggest that WT ß-catenin still acts
as a gatekeeper of hepatocyte differentiation and polarity, even in tumor hepatocytes.
Fascin-1, as a target of ß-catenin, involved in hepatocyte dedifferentiation
We searched, in our transcriptomic data, for genes antagonistically expressed upon
silencing of WT and mutated ß-catenin in HepG2 cells, potentially impacting both
their differentiation and polarity. We focused on FSCN-1 found up-regulated upon
WT ß-catenin KD and down-regulated upon mutated ß-catenin KD (Fig. S5A). FSCN-
1 encodes fascin-1, an actin-bundling protein, which is normally not expressed in
epithelial cells, i.e. absent in mature hepatocytes. Instead, villin, encoded by the VIL1
gene, is the actin-bundling protein associated to BC microvilli in differentiated cells.
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14
As villin and fascin-1 that share similar function, were found altered in an opposite
manner upon ß-catenin KD (Fig. S3D-E and S5A), we further studied the role of
fascin-1 in hepatocyte dedifferentiation. Moreover, fascin-1 is expressed in tumors
including HCC17 and described as a transcriptional target of β-catenin in breast,
gastric and colon cancer cells18-20. We first analyzed fascin-1 protein expression in
HB and HCC cell lines, with different ß-catenin status (Fig. S5B). We found that
fascin-1 was more expressed in cell lines bearing-CTNNB1 deletion (HepG2) or
mutations (SNU398 and Huh6), compared to non-mutated (Huh7, Hep3B) cell lines.
The level of Fascin-1 protein was positively correlated to ß-catenin protein level,
which accumulates upon mutations (Fig. S5C). As a ß-catenin transcriptional target,
fascin-1 mRNA expression was strongly inhibited upon treatment of HepG2 cells with
siRNAs sißcat-mut and sißcat-both (Fig. 5A). As previously observed for Axin2 and
CyclinD1 (Fig. 1E), the expression of fascin-1 is slightly but significantly upregulated
upon removal of WT ß-catenin. This alteration was also detected at the protein level
(Fig. S5D). Using a luciferase assay with fascin-1 promoter, we show that this
regulation occurs at the transcriptional level, with an antagonistically regulation of
promoter activity upon silencing of either WT or mutated ß-catenin (Fig. 5B). Thus, as
previously shown in other cancer cells, our results confirm that FSCN1 is a target
gene of β-catenin in tumor hepatocytes. We next wonder whether the variation of
fascin-1 expression may play a role in the alteration of hepatocyte differentiation in a
tumor context. Interestingly we found that fascin-1 silencing led to an increase of
ApoC3, E-cadherin and claudin1 mRNA expressions in HepG2 cells (Fig. 5C). The
same tendency was observed in HCC cell lines (Fig. S6A-B). We also found that
fascin-1 KD (Fig. 5D) led to an increase of two to three folds in the number of BC in
HepG2 cells (Fig. 5E). Moreover, we show that the depletion of fascin-1 upon
inhibition of WT β-catenin expression restores the formation of BC (Fig. 5F),
demonstrating that fascin-1 is one of the effectors responsible of the impact of ß-
catenin on BC formation. Finally, overexpression of fascin-1 induced a decrease in
BC formation in comparison to control (Fig. 5G). Altogether these results
demonstrated that the level of fascin-1 regulates hepatocyte differentiation status.
Moreover, the impact of WT and mutated ß-catenin silencing on hepatocyte
differentiation is in part due to fascin-1 expression.
Fascin expression is high in undifferentiated tumors in mice
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15
We next aimed at validating fascin-1 as a ß-catenin target in liver tumors in mice. We
used mouse models that mimic ß-catenin dependent tumorigenesis, such as the APC
loss-of-function and the ∆exon3 ß-catenin models. These models were shown to lead
to ß-catenin pathway activation and the development of phenotypically
undistinguishable liver tumors within about 10 months15, 21. Interestingly, two
phenotypically distinct tumors defined as differentiated and undifferentiated tumors
are generated in these mice15, 21. Well-differentiated tumors are characterized by
hepatocyte-like tumor cells that maintain a β-catenin-induced expression of glutamine
synthetase (GS) and present nuclear β-catenin. Undifferentiated tumors are
composed of small cells with basophilic nuclei, that they strongly express nuclear β-
catenin15. A RNA-Seq analysis showed that these tumors lose the expression of GS,
which is reminiscent of a loss of differentiation (Fig. S7A). We thus made use of this
mouse cohort to analyze fascin-1 expression (Table S3). RNA sequencing data from
these murine tumors demonstrated that fascin-1 expression is increased in both,
well-differentiated and undifferentiated types of tumors in comparison to normal liver
(Fig. 6A and S7B). We further highlighted that fascin-1 expression is higher in
undifferentiated samples than in the differentiated ones (Fig. 6B). Moreover, fascin
mRNA expression was found to correlate positively with various markers of
undifferentiated tumors such as MMP2, VIM, HIF1A and YAP1, and negatively with
markers of differentiated tumors such as HNF4a, APOC3 and GJB1 (Fig. S7C).
Finally, as a bona fide ß-catenin target, fascin-1 expression correlated positively with
level of CTNNB1, LEF1 and TCF4 (Fig. S7C). These new findings were confirmed by
immunohistochemistry performed on both types of murine tumors obtained from a
new cohort of APC KO mice. As described previously15, activation of the β-catenin
pathway in the tumor cells was characterized by nuclear localization of the β-catenin
along with high expression of GS in well-differentiated tumors (Fig. 6C) and a lower
to almost absent expression in undifferentiated ones (Fig. 6D). Fascin-1 staining,
which is absent of the non-tumoral hepatocytes, was found increased in both types of
tumors. Moreover, whereas its expression is low in differentiated tumors, fascin
expression appears high in undifferentiated tumor cells, with a cytoplasmic and
membranous staining. Thus, in mice, fascin-1 expression is a marker of ß-catenin-
induced undifferentiated tumors. As those murine tumors were found to be
transcriptionally close to human mesenchymal HBs15, this prompted us to explore
Fascin-1 expression in human HBs.
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16
Fascin-1 is a marker of embryonal contingent in human HB
In order to analyze fascin-1 expression in human HBs we first made use of public
datasets3,7. We found that fascin-1 mRNA is specifically expressed in the C2-subtype
of HBs that corresponds to poor-prognosis tumors, with no apparent difference
between C2A and C2B subtypes (Fig. 7A). As observed in mice, we found that
FSCN1 mRNA expression correlates negatively with markers of differentiated
hepatocytes such as HNF4a, APOC3, GJB1 and CLDN1 (Fig. S8A). We then used
immunohistochemistry to confirm expression of the protein product in HB samples.
Whereas fascin1 is not expressed in human normal hepatocytes and restricted to
sinusoidal cells in normal and peri-tumoral tissues (Fig. S8B), we found that fascin-1
is expressed in a specific contingent of tumor cells in HBs (Fig. 7B). Fascin-1 staining
is highly consistent with the results obtained in mice, showing that fascin-1 is
expressed in GS-negative and ß-catenin highly positive cells (Fig. 7B and S7C).
These cells correspond to the embryonal contingent of undifferentiated small cells
with basophilic nuclei. Thus, in human, as found in mice, fascin-1 expression is a
marker specific of ß-catenin-induced undifferentiated tumors.
DISCUSSION
The two functions of ß-catenin, structural at cell-cell junctions and transcriptional in
the nucleus are difficult to dissociate and to study individually. Indeed as carried by
the same protein, both functions are intrinsically linked. Here, we developed the dual
ß-catenin KD HepG2 model, which allow us to address the interplay between the WT
and the mutated ß-catenin in liver tumor cells. We found that the mutated form of ß-
catenin is dedicated to the transcriptional function, whereas the WT ß-catenin,
without Wnt stimulation, is more endowed with its adhesive function in HepG2 cells.
Thus, this model is suitable to address independently the structural and the
transcriptional functions of ß-catenin. Interestingly, the transcriptomic analysis of the
dual ß-catenin KD HepG2 model revealed that disrupting each of the function alter
gene expression. Consistent with the transcriptional activity of ß-catenin, genes that
were dysregulated upon removal of mutated ß-catenin were coherent with a Wnt/ß-
catenin signature. In contrast, genes altered upon silencing of the WT allele were
endowed with different signaling pathways that remain to be explored. We found that
transcriptional and adhesive activities of β-catenin play antagonistic role in tumor
hepatocytes. Whereas the structural function of ß-catenin is necessary to maintain a
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17
differentiated state of hepatocytes, the transcriptional activity of ß-catenin induces the
dedifferentiation program of HepG2 cells. KD of each allele specifically reverts those
programs. As published earlier16, we found that this antagonism is in part due to the
regulation of the transcriptional activity of HNF4a, required to maintain a hepatocyte
differentiation program. Consistent with this antagonism between the transcriptional
and the structural activities of ß-catenin we found that WT and mutated ß-catenins
act in an opposite manner on hepatocyte polarity.
Hepatocytes polarization involves the formation of functionally distinct sinusoidal
(basolateral) and bile canalicular (apical) plasma membrane domains that are
separated by tight junctions. A normal membrane polarity is essential for hepatocyte
function and its loss may lead to many diseases including cholestasis-associated
diseases. A link between β-catenin and BC abnormalities is known since the middle
of 2000’s but it remains puzzling. Indeed, on one hand, cholestasis is a key feature of
ß-catenin-mutated HCCs22, but on the other hand the liver-specific β-catenin KO
mice were shown to also develop intrahepatic cholestasis associated with BC
abnormalities and bile secretory defect23. These data suggest that loss of ß-catenin
as well as excessive activation of ß-catenin may lead to cholestasis in the liver.
Whether this phenotype is due to structural or transcriptional activity of ß-catenin is
largely unknown. Several studies reported the involvement of cell adhesion
molecules in the maintenance of BC. In cultured HepG2 cells or embryonic chicken
hepatocytes, E-cadherin was shown to be important for BC lumen extension24, 25.
However, its liver specific knock-out in mice did not alter hepatocyte polarity and BC
formation26. More recently, hepatocyte specific KD of α-catenin was shown to alter
BC, resulting of tight junction disruption and enlarged lumens27. Our approach
permits to uncouple the different functions of WT or mutated β-catenin revealing their
involvement in BC formation. We found that the two forms of ß-catenin have an
antagonist role, mutated ß-catenin playing a repressor role by decreasing the
number, the size, and the functionality of BC, and WT ß-catenin being important for
their maintenance. As demonstrated for the development of bile ducts28, ß-catenin
has to be kept at the right level as loss of ß-catenin or ß-catenin overactivation is
detrimental for BC, formation and/or stabilization.
Our work further highlights the involvement of fascin-1 downstream of ß-catenin in
the regulation of hepatocyte differentiation. Fascin-1 is a protein that links actin
filaments to form bundles of actin present in filopodia or invadopodia29. However,
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18
fascin-1 is normally not present in epithelial cell microvilli. In our experiments, fascin-
1 behaves as a transcriptional target of ß-catenin. Links between fascin-1 and ß-
catenin were reported earlier in the literature. Indeed fascin-1 has been reported to
be a transcriptional target of β-catenin/TCF signaling in colon cancer cells showed by
the inhibition of fascin promoter activation using dominant-negative TCF4 in vitro18. In
addition, ß-catenin was shown to bind constitutively to fascin promoter in carcinoma
cells30. However, this regulation of fascin-1 by ß-catenin seems to be cell type-
specific as no regulation was observed in breast cancer cells20. On the other way
around, fascin-1 was found to induce epithelial-mesenchymal transition of
cholangiocarcinoma cells and promote breast cancer stem cell function by regulating
Wnt/ß-catenin signaling31, 32.
One finding of our study is that fascin-1 expression alters hepatocyte differentiation
status. We indeed found that fascin-1 is highly expressed in ß-catenin-mutated
undifferentiated tumors both in mice and in human. In both, fascin-1 expression
correlates negatively with differentiated hepatocyte markers and positively with
mesenchymal markers. In addition, in vitro, we demonstrated that fascin-1 levels
directly act on the hepatocyte polarity and differentiation status. We found that
silencing of fascin-1 allows an upregulation of epithelial markers and an increase of
BC formation. How fascin-1, an actin-binding protein, may regulate epithelial and
mesenchymal gene expression, remains to be explored. Interestingly, Fascin-1 is
present in a large list of proteins found to bind mRNA, suggesting a potential role of
fascin in post-translational regulation33. But it is also now widely accepted that the
modulation of the actin cytoskeleton acts on gene expression through
mechanotransduction pathways.
Thus, our data demonstrate that fascin-1 is a new target of ß-catenin in the liver that
plays a key role in the modulation of hepatocyte differentiation. We found that Fascin-
1 is a new marker of the embryonal contingent in HBs. Even if fascin-1 is enriched in
the C2 subclass of HBs, we also found fascin-1 staining in C1 samples, reporting the
complexity of these tumors. As Fascin-1 is associated to HBs with bad prognosis,
fascin-1 immunostaining may offer new diagnostic/pronostic opportunities. Moreover,
fascin-1 may be suitable to consider as a new actionable target in these liver
pediatric tumors.
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19
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Author names in bold designate shared co-first authorship
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21
FIGURE LEGENDS
Figure 1: The HepG2 dual ß-catenin KD model. (A) Schema showing the different
siRNAs used to KD wild-type (WT), mutated or both ß-catenins on the two alleles of
CTNNB1 gene in HepG2 cells. (B-E) HepG2 cells were transfected with indicated
siRNAs. (B) Protein extracts were analyzed using anti-ß-catenin and GAPDH
antibodies. Note that due to the W25-I140 deletion, mutated ß-catenin (about 76
kDa) migrates faster and is more abundant (because, more stable) than the WT ß-
catenin (about 92 kDa). (C) ß-catenin mRNA expression was analyzed by qRT-PCR.
(D) Promoter activity was evaluated by luciferase reporter assays in HepG2 cells
transfected with TCF responsive reporter. Shown is the mean relative luciferase
activity, normalized to Renilla luciferase and compared to siRNA control transfected
cells. (E) mRNA levels of indicated genes were analyzed by qRT-PCR. LGR5
(GPR49), Axin2 and CCND1 (cyclinD1) are positive transcriptional targets of ß-
catenin. ARG1 encoding arginase is a negative transcriptional target of ß-catenin.
Shown is the relative mRNA level compared to control transfected cells. (C-E) Each
graph shows the quantification of three independent experiments. Error bars indicate
s.e.m (n=3), P values from one way ANOVA. (F) HepG2 cells transfected with
indicated siRNAs were seeded in 96-well plates and cells were fixed and total
biomass, reflecting the number of cells, was assayed every day. Each time point was
performed in triplicates. Error bars indicate s.e.m (n=3). P value from one way
ANOVA.
Figure 2: HepG2 cells transfected with indicated siRNAs were fixed and stained with
(A) anti-E-Cadherin antibodies (green) and Hoescht (blue), or (B) phalloidin (red),
anti-ß-catenin antibodies (green) and Hoescht (blue). Scale bar: 10 µm.
Figure 3: (A) Alteration of the expression of differentiated hepatocyte markers upon
silencing of both alleles of ß-catenin in HepG2 cells. The graph shows the relative
expression of the indicated genes extracted from the transcriptomic analysis. (B-D)
HNF4a activity is up-regulated upon silencing of mutated ß-catenin. (B) Indicated
siRNA were transfected in HepG2 cells, and HNF4A relative mRNA level was
analyzed. Shown is the relative mRNA level compared to control transfected cells.
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22
(C) HepG2 cells transfected with indicated siRNAs were transfected with HNF4a
responsive luciferase reporter. Shown is the mean relative luciferase activity,
normalized to Renilla luciferase and compared to siRNA control transfected cells. (D)
Relative mRNA levels of APOC3 and APOM, both positive transcriptional targets of
HNF4a were analyzed by qRT-PCR. Shown is the relative mRNA level compared to
control transfected cells. (E) Alteration of the expression of polarity markers upon
silencing of both alleles of ß-catenin in HepG2 cells. The graph shows the relative
expression of the indicated genes extracted from the transcriptomic analysis. (B-E)
Each graph shows the quantification of three independent experiments. Error bars
indicate s.e.m (n=3), P values from one way ANOVA.
Figure 4: Alteration of bile canaliculi formation upon silencing of ß-catenin in HepG2
cells. (A) HepG2 cells transfected with indicated siRNAs were fixed and stained with
phalloidin (red), anti-radixin antibodies (green) and Hoescht (blue). Scale bar: 15 µm.
(B) Quantification of the percentage of cells forming BC in the conditions described in
(A). Graph shows the quantification of four independent experiments, where at least
100 cells were observed per experiment. (C) siRNA transfected HepG2 cells were
fixed, stained with phalloidin-ATTO and observed by STED microscopy. Scale bar: 5
µm. (D) BC features (area, perimeter, circularity, min Feret (diameter)) were
quantified by imageJ on STED images performed as described in (C). Each dot
corresponds to one BC. (E) Quantification of BC functionality by using CFDA
incorporation. Life-Act (red) expressing HepG2 cells were treated with CFDA (green)
and observed with fluorescence microscope. Scale bar: 25 µm. Graph shows the
quantification of three independent experiments. (B, D-E) Error bars indicate s.e.m
(n=4 for B, n=3 for D-E), P value from one way ANOVA.
Figure 5: Fascin-1, a target of β-catenin, alters hepatocyte differentiation status. (A)
Fascin-1 mRNA expression upon depletion of WT and/or mutated β-catenin analyzed
by RT-qPCR in HepG2 cells. (B) Activity of Fascin-1 promoter studied by reporter
luciferase assay. Shown is the mean relative luciferase activity, normalized to Renilla
luciferase and compared to control siRNA transfected cells. (C) mRNA levels of
FSCN1, APOC3, CLDN1 and CDH1 upon treatment of HepG2 cells with siRNA
targeting Fascin-1 (siFascin1#1, #2, #3 and #4). (D) HepG2 were transfected with
indicated siRNAs, and protein extracts were analyzed using anti-Fascin-1, ß-actin
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23
and GAPDH antibodies. The graph shows the quantification of three independent
experiments. (E) siRNA transfected HepG2 cells were fixed and stained with
phalloidin (red), anti-radixin antibodies (green) and Hoescht (blue). Scale bar: 15 µm.
The graph shows the quantification of three independent experiments where the
number of BC formed for 100 cells are indicated. (F-G) Experiments were performed
as described in (E) with either co-transfection of indicated siRNAs (F) or transfection
of pEGFP-C1 (GFP) or pEGFP-C1-Fascin-1 (GFP-Fascin-1) (F). (A-G) Graphs show
the quantification of at least three independent experiments. Error bars indicate s.e.m
(n=4 for A, n=5 for B and n=3 for C-F), P values from one way ANOVA.
Figure 6: Fascin-1 expression in murine ß-catenin-mediated tumors. (A-B) Data were
extracted from RNAseq performed on mouse hepatic tumors induced either from
APC KO or ß-catenin ∆exon3 expression in livers. FSCN-1 expression is shown in
non-tumoral samples (n=12) and in differentiated (n=13) and undifferentiated (n=6)
tumors. P values from one way ANOVA. (C) Representative images of
immunohistochemistry of ß-catenin, Fascin-1, and glutamine synthetase (GS) in
differentiated (HCC-type) (C), and undifferentiated (HB-type) (D) murine tumors.
Boxed regions are enlarged in the zoom images.
Figure 7: Fascin-1 expression in human HBs. (A) Data were extracted from RNAseq
performed on human HBs by Hooks et al.7. FSCN-1 expression is shown in non-
tumoral samples (n=30) and in C1 (n=20) and C2A or C2B (n=9) tumors. P values
from one way ANOVA. (B) Representative images of immunohistochemistry of ß-
catenin, Fascin-1 and GS and HES staining in a C2B HB pediatric case. Boxed
regions are enlarged in the zoom images.
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SUPPLEMENTARY MATERIALS & METHODS
3D Cell growth assay
Three-dimensional multicellular spheroids were prepared by seeding cells on a non-
adherent surface. Briefly, 2000 cells were added per well, in Ultra Low Attachment
96-well tissue culture microplates (COSTAR 7007) and incubated at 37°C in
complete culture medium. Spheroid formation was assessed 24 h later and spheroid
growth was followed using the Incucyte system (Essen BioSciences). Spheroid area
was measured using Image J software.
Transcriptomic analysis
Transcriptomic analysis was performed at the plateforme GenomEast of Strasbourg
(Strasbourg, France). Biotinylated single strand cDNA targets were prepared, starting
from 150 ng of total RNA, using the Ambion WT Expression Kit (Cat # 4411974) and
the Affymetrix GeneChip® WT Terminal Labeling Kit (Cat # 900671) according to
Affymetrix recommendations. Following fragmentation and end-labeling, 3 µg of
cDNAs were hybridized for 16 hours at 45°C on GeneChip® Human Gene 2.0 ST
arrays (Affymetrix) interrogating 24 838 genes and 11 086 LincRNAs represented by
approximately 21 probes spread across the full length of the RNA. The chips were
washed and stained in the GeneChip® Fluidics Station 450 (Affymetrix) and scanned
with the GeneChip® Scanner 3000 7G (Affymetrix) at a resolution of 0.7 µm. Raw
data (.CEL Intensity files) were extracted from the scanned images using the
Affymetrix GeneChip® Command Console (AGCC) version 3.2. CEL files were
further processed with Affymetrix Expression Console software version 1.1 to
calculate probe set signal intensities using Robust Multi-array Average (RMA)
algorithms with default settings. The raw data were archived in the public GEO data
repository with the GEO accession GSE144107. Differentially expressed (DE) genes
were identified using the fold-change rank ordering statistics (FCROS) method13,
which associates an f-value with genes. Small f-values (near zero) are associated
with down-regulated genes, while higher values (near 1) are associated with up-
regulated genes.
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SUPPLEMENTARY FIGURE LEGENDS
Figure S1: (A) HepG2 were transfected with indicated siRNAs and protein extracts
analyzed by immunoblotting using anti-ß-catenin and GAPDH antibodies. Graphs
show the quantification of WT (left-hand graph) and mutated (right-hand graph) ß-
catenin protein levels normalized to GAPDH. Shown is the relative protein level
compared to control transfected cells. Error bars indicate s.e.m (n=3) *** P < 0.001
by one way ANOVA. (B) HepG2 cells transfected with indicated siRNAs were seeded
in non-adherent 96-well plates and growth of spheroids was followed using the
Incucyte system. The graph shows the mean area of 6 spheroids upon time. This
experiment is representative of the three performed.
Figure S2: Transcriptomic analysis of the dual ß-catenin KD HepG2 model. (A) Data
extracted from the microarray, showing the expression alteration of CTNNB1, LGR5,
CCND1 and AXIN2 genes, encoding respectively ß-catenin, GPR49, cyclinD1 and
Axin2. Each graph shows the quantification of the three replicates. Error bars indicate
s.e.m *** P < 0.001; **** P < 0.0001 by one way ANOVA. (B) Pearson’s correlations
between each dataset obtained by Affimetrix. Datasets 1, 5, 9 correspond to HepG2
transfected with control siRNA. Datasets 2, 6, 10 correspond to HepG2 transfected
with sißCat WT. Datasets 3, 7, 11 correspond to HepG2 transfected with sißCat both.
Datasets 4, 8, 12 correspond to HepG2 transfected with sißCat mut. The color code
indicates Pearson’s correlation. (C) Venn diagram of altered genes in sißCat WT,
sißCat mut and sißCat both conditions.
Figure S3: Analysis of the expression of 16 HNF4a positive transcriptional targets
extracted from the transcriptomic data, upon silencing of WT ß-catenin (A) or
mutated ß-catenin (B) in HepG2 cells. The graphs show the relative expression of the
indicated genes extracted from the transcriptomic analysis. Each graph shows the
quantification of three independent experiments. Error bars indicate s.e.m (n=3) * P <
0.05; ** P < 0.01; *** P < 0.001 by t-test compared to control. (C) Pooled data of the
16 HNF4a targets. **** P < 0.0001. (D-E) Alteration of the expression of polarity-
associated genes upon silencing of both alleles of ß-catenin in HepG2 cells. Graphs
show the relative expression of the indicated genes extracted from the transcriptomic
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analysis. Each graph shows the quantification of three independent experiments.
Error bars indicate s.e.m (n=3) * P < 0.05; ** P < 0.01; **** P < 0.0001 by t-test
compared to control.
Figure S4: (A) HepG2 cells transfected with siRNA targeting mutated ß-catenin were
fixed, stained with phalloidin-ATTO and observed by confocal (left-hand) and STED
(right-hand) microscopy. Scale bar: 2.5 µm. (B) HepG2 cells transfected with
indicated siRNAs were fixed and stained with phalloidin (red), anti-radixin antibodies
(green) and Hoescht (blue). The number of cells engaged in the formation of bile
canaliculi was evaluated under the microscope. Graph shows the quantification of
three independent experiments. Error bars indicate s.e.m.
Figure S5: Regulation of Fascin-1 expression by β-catenin. (A) Data extracted from
the transcriptomic data, upon silencing of WT ß-catenin and/or mutated ß-catenin in
HepG2 cells, showing the expression of FSCN1 gene, encoding Fascin-1. The graph
shows the quantification of the three replicates. Error bars indicate s.e.m. * P < 0.05;
** P < 0.01 by one way ANOVA. (B) Protein extracts from indicated cell lines were
analyzed by immunoblotting using anti-ß-catenin, Fascin-1 and ß-actin antibodies.
Arrows indicated mutated (about 76 kDa) and WT (about 92 kDa) ß-catenins in
HepG2 cells. Status of CTNNB1 gene in cells is indicated below the immunoblot. (C)
Correlation of the level of Fascin-1 and β-catenin in various cell lines, as quantified
by immunoblot. (D) Fascin-1 protein expression upon depletion of WT and/or
mutated β-catenin observed by western blot. Actin and GAPDH serve as loading
controls.
Figure S6: (A-B) mRNA levels of FSCN1, APOC3, CLDN1 and CDH1 upon treatment
of Hep3B (A) and Huh7 (B) cells with siRNA targeting Fascin-1 (siFascin1#1, #2, #3
and #4). Graphs show the quantification of three independent experiments. Error
bars indicate s.e.m. * P < 0.05; ** P < 0.01; *** P < 0.001; **** P< 0.0001 by one way
ANOVA.
Figure S7: Fascin-1 expression in murine activated ß-catenin-mediated tumors. (A-C)
Data were extracted from analyses performed on mouse ß-catenin-mediated tumors
by Loesch et al. (Loesch et al., submitted). GLUL encoding glutamine synthetase (A)
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and FSCN-1, encoding Fascin-1 (B) are shown in non-tumoral samples (n=12) and in
differentiated (n=13) and undifferentiated (n=6) tumors. (C) FSCN1 expression
correlates negatively with differentiation markers (highlighted in blue) and positively
with undifferentiation markers (highlighted in red) in mouse liver tumors.
Figure S8: Fascin-1 expression in human HBs. (A) Data were extracted from
published analyses from performed on human HBs by Cairo et al., 2008. FSCN1
expression correlates negatively with HNF4A, APOC3, CLDN1 and GJB1 in human
HBs. (B) Representative image of immunohistochemistry of Fascin-1 in a normal
human liver. (C) Representative images of immunohistochemistry of ß-catenin,
Fascin-1 and glutamine synthetase (GS), and HES staining in a C1 HB pediatric
case. Dashed rectangles show part of the images showed in the zoom images.
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Supplementary Tables
Supplementary Table 1: List of the antibodies used in the study.
Supplementary Table 2: List of the qRT-PCR primers used in the study. Forward and
reverse primers are indicated for each gene.
Supplementary Table 3: Mouse cohorts used in this study.
Supplementary Table 4: Clinical data of patient samples used for
immunohistochemistry analyses.
Supplementary Tables 5-7: List of the deregulated genes upon WT (Table S2),
mutated (Table S3) or both (Table S4) ß-Catenin KD in HepG2 cells. Differentially
expressed genes were identified using the fold-change rank ordering statistics
(FCROS) method (Dembélé D, Kastner P. BMC Bioinformatics 2014;15:14) and
which associates an f-value with genes. Small f-values (near zero) are associated
with down-regulated genes (highlighted in green), while higher values (near 1) are
associated with up-regulated genes (highlighted in pink). “ttest" et "sam" correspond
to statistical analyses using, respectively, the Student t-test and the Significant
Analysis of Microarrays (http://statweb.stanford.edu/~tibs/SAM/) methods.
Supplementary Table 8: Analysis with FuncAssociate 3.0 using the genes listed on
tables S5-7.
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