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The role of fibroblast growth factor signaling on postnatal hepatic progenitor cell expansion
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The role of fibroblast growth factor signaling on postnatal hepatic progenitor cell expansion
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i
THE ROLE OF FIBROBLAST GROWTH FACTOR SIGNALING ON POSTNATAL
HEPATIC PROGENITOR CELL EXPANSION
by
Sarah Utley
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(INTEGRATIVE BIOLOGY OF DISEASE)
August 2013
Copyright 2013 Sarah Utley
ii
Acknowledgments
It has certainly been a long and adventurous course to get me to this point and I have so
many people to thank for their support and guidance. First and foremost I want to thank my
mentor, Dr. Kasper Wang, MD, for being the most supportive mentor I could ever ask for. With
Dr. Wang’s guidance and support, I have become a more independent scientist which I know
will invaluable to my future career. I would also like to thank all of my committee members who
have guided me and pushed me to be a better scientist: Dr. Martin Pera, Dr. David Warburton,
Dr. Hide Tsukamoto, Dr. Laurie DeLeve, and Dr. Mark Frey.
I have also had the pleasure of working with many wonderful co-workers throughout my
journey. I could not have found my love for liver biology without the guidance and joy that
Tove Berg, Ph.D. brought to every day we worked together in the lab. I would not have been so
inspired to join the lab if it hadn’t been for her. I also would like to thank Nirmala Mavila, Ph.D.
for all of her support and mentorship she has provided over the past few years. All of the long
hours of work could not have been possible without the amazing lab technicians we’ve had over
the years, including Mike Salisbury, Jenn Phan, and David James. I could not have gotten all of
this work done without their technical help, encouragement, fun we had together in the lab.
I would also like to thank my fellow graduate students who helped to push me to be a
better scientist while being an amazing group of friends: Lauren Geary, Yvonne Leung, Ben
Lam, and Christine Cao. I could not have made it through this journey without their friendship,
support, and scientific inspiration throughout the years. I would especially like to thank Lauren
Geary for being an amazing friend every step of the way. My Ph.D. would not have been the
same without you! I also want to thank Orchid Rogers Ph.D. for being such a wonderful friend
iii
and inspiration to me. I could not have made it through my Ph.D. without our many coffee
breaks and long discussions about experiments and career choices.
Last, but certainly not least, I am so grateful for my amazing family. I would never have
been able to make it this far without all of their support over the past six years. My parents have
pushed me to become a better person and scientist. My brothers and sister-in-law have made this
whole journey wonderful by keeping me laughing about it all. I would also like to acknowledge
the administrative staff at CHLA and in the PIBBS and SBD programs: Joanna Cavallero, Dawn
Burke, Raquel Gallardo, and Marisela Zuniga, for all of their help throughout my Ph.D.
iv
Dedication
I would like to dedicate this body of work to my amazing parents, Debbie and David Utley. I
would have never aspired to pursue a Ph.D. and a career and science without the endless support
and inspiration you have given me my whole life.
v
Table of Contents
Acknowledgements ii
Dedication iv
List of Tables vi
List of Figures vii
Abbreviations ix
Abstract xii
Chapter 1: Introduction
1 Liver Development
2 Hepatic Progenitor Cells in Liver Disease
1
1
19
Chapter 2: Over-expression of Fgf10 Expands the Periportal
Hepatic Progenitor Cell Population in Early Postnatal Liver
2.1 Abstract
2.2 Introduction
2.3 Materials and Methods
2.4 Results
2.5 Discussion
25
25
26
27
29
41
Chapter 3: Fibroblast Growth Factor Signaling Regulates the
Expansion of A6-Expressing Progenitor Cells via AKT-Dependent
β-catenin activation during DDC-induced liver injury
3.1 Abstract
3.2 Introduction
3.3 Materials and Methods
3.4 Results
3.5 Discussion
44
44
45
47
50
63
Chapter 4: Concluding Remarks and Future Perspectives
5.1 Concluding Remarks
5.2 Future Perspectives
67
67
68
References
72
Bibliography 83
vi
List of Tables
Table 2.1 Antibodies for Immunostaining
28
Table 3.1
Antibodies for Immunostaining 48
Table 3.2
qRT-PCR Primers 49
Table 3.3
Effects of dnFGFR and Fgf10 Over-Expression
on DDC Liver Injury
52
Table 3.4
Effects of pAKT Inhibition with Fgf10 Over-
Expression on DDC Liver Injury
59
vii
List of Figures
Figure 1.1 Generic FGF Ligand and Receptor Protein Structure
3
Figure 1.2 FGFR Activated Intracellular Signaling Pathways
5
Figure 1.3 Canonical Wnt Signaling Pathway
8
Figure 1.4 Non-canonical β-catenin Activation
10
Figure 1.5 Notch Signaling Pathway
16
Figure 2.1 Early Postnatal Periportal Cell Population Peaks at Postnatal Day 7
29
Figure 2.2 Fgf10 Over-Expression Retains the P7 Periportal Cell Population
31
Figure 2.3 Dominant Negative FGFR2b Inhibition Does Not Disrupt the
Expansion of the P7 Periportal Cell Population
31
Figure 2.4 Periportal Cells Express FGF Receptors
33
Figure 2.5 Periportal Cells are a Heterogeneous Cell Population of Epithelial and
Hematopoietic Cells
34
Figure 2.6 Periportal Cells Express ALBUMIN but not HNF4 α
35
Figure 2.7 Fgf10 Induced Periportal Cells Express Hepatic Progenitor Cell
Markers
36
Figure 2.8 CD133
+ive
Cell Progeny Form a Subset of the Liver Parenchymal Cells
During Postnatal Development
38
Figure 2.9 CD133 Progeny Form a Subset of Biliary Epithelial Cells and
Hepatocytes in Postnatal Development
39
Figure 2.10 Notch1 Signaling is Activated with Fgf10 Over-Expression
40
Figure 2.11 AKT-mediated β-catenin activation is not regulated in the early
postnatal periportal cell population
41
Figure 3.1 FGF Signaling is Up-Regulated During DDC Injury
51
Figure 3.2 FGF Signaling Regulates A6
+ive
HNF4 α
+ive
Cell Expansion
55
Figure 3.3 Fgf10 over-expression in non-injured adult livers induces A6
expression in hepatocytes.
56
viii
Figure 3.4 FGF Signaling Regulates Heterogeneous Periportal Cell Population
Proliferation and AKT-Dependent β-catenin Phosphorylation
58
Figure 3.5 Wortmannin Inhibition of Fgf10 Induced Livers Disrupts the
A6
+ive
HNF4 α
+ive
Cell Differentiation and Migration
60
Figure 3.6 Wortmannin Inhibition Decreases Periportal Cell Proliferation
61
Figure 3.7 CD133
+ive
cells form a subset of biliary epithelial cells and hepatocytes
in non-injured livers and DDC injured livers
62
ix
Abbreviations
αSMA alpha smooth muscle actin
AFP α-fetoprotein
AGS Alagille syndrome
AKT Protein Kinase B
Alb Albumin
APC Adenomatous Polyposis Coli
BA biliary atresia
BDL bile duct ligation
BECs biliary epithelial cells
BMP Bone Morphogenetic Protein
CD13 Aminopeptidase N
CD49f Integrin α6
CD133 Prominin 1
CITED-1 CBP/P-300 interacting trans-activator 1
CK19 Cytokeratin 19
CYP7A1 cholesterol 7alpha hydroxylase
DDC 3,5-diethoxycarbonyl-1,4-dihydrocollidine
Dlk1 delta-like 1 homolog
DLL delta-like ligand
Dvl Dishevelled
dnFGFR dominante negative fibroblast growth factor receptor
E8.5 embryonic day 8.5
E-Cadherin epithelial cadherin
EHBD extra-hepatic bile duct
x
EMT epithelial-to-mesenchymal transition
Ep-CAM epithelial cell adhesion molecule
ERK extracellular signal-regulated kinase
ESC embryonic stem cell
Foxa1 forkhead box protein A1
FGF fibroblast growth factor
FGFR fibroblast growth factor receptor
FOXL1 forkhead box L1
FRS2 fibroblast growth factor receptor substrate 2
Fz frizzled
GAB1 GRB2-associated binder 1
GSK3 β glycogen synthase kinase-3 beta
GRB2 growth factor receptor bound protein 2
H&E hematoxylin and eosin
Hex homeobox gene
HGF Hepatocyte Growth Factor
HNF4 α hepatocyte nuclear factor 4 alpha
HNF1 β hepatocyte nuclear factor 1 beta
HPC hepatic progenitor cell
HSC hematopoietic stem cell
HSPG heparin sulfate proteoglycan
IF immunofluorescence
IFN γ interferon-gamma
IHBD intrahepatic bile duct
JAG Jagged
xi
MAPK Mitogen-Activated Protein Kinase
MET mesenchymal-to-epithelial transition
NICD Notch intracellular domain
NHF non-hepatocyte fraction
P7 postnatal day 7
PBC primary biliary cirrhosis
PBS phosphate buffered saline
PCA periportal cell area
PFA paraformaldehyde
PI3K Phosphatidylinositide 3-kinases
PSC primary sclerosing cholangitis
PVA portal vein area
PVM portal vein mesenchyme
RAS rat sarcoma
RBP-J recombination signal binding protein immunoglobulin kappa J
RRV rhesus rotavirus
RTK receptor tyrosine kinase
SHH sonic hedgehog
SOS son of sevenless
SOX9 SRY-related HMG box transcription factor 9
Tcf/Lef T-cell transcription factor/Lymphocyte enhancer factor
TGF- β Transforming Growth Factor
WNT Wingless
WT wild type
xii
Abstract
Fibroblast Growth Factor (FGF) signaling is an established regulator of endoderm specification
to the hepatic fate. We have previously determined that Fibroblast Growth Factors (FGFs)
promote the proliferation and survival of embryonic hepatic progenitor cells (HPCs), termed
hepatoblasts, and a transformed murine HPC line via AKT-dependent β-catenin activation.
Recent studies have shown that postnatal expansion of A6-expressing HPCs during 3,5-
diethoxycarbonyl-1,4-dihydrocollidine (DDC) induced liver injury is partly mediated by β-
catenin activation and FGF signaling. Herein, we examine the role of FGF signaling in early
postnatal HPCs and acute DDC-induced HPC expansion. Methods: The effects of FGF10 on
early postnatal HPCs was studied by inducible transgenic pups over-expressing Fgf10 (P21
Fgf10 Induced) from postnatal day 7 (P7) to P21. For postnatal liver injury, inducible transgenic
mice were fed 0.1% DDC chow for 14 days concurrent with either over-activation of FGF
signaling by Fgf10 over-expression or inhibition of FGF signaling via dominant-negative
expression of soluble FGF Receptor (R)-2IIIb. Results: During early postnatal development, a
resident population of cells with a high nuclei-to cytoplasmic ratio increases from postnatal day 0
(P0) through P7 before it disappears histologically. Over-expression of Fgf10 from P7 to P21
retains and expands the periportal cell population which normally declines after P7.
Characterization of this cell population revealed they express HPC markers CD49f, and CD133,
but lack expression of markers for mesenchymal cells, differentiated hepatocytes and biliary
epithelial cells. This heterogeneous cell population expressed epithelial cell marker E-Cadherin,
and a subset of the cells was CD45 positive, indicating infiltration of hematopoietic cells as well.
During acute postnatal DDC injury, we observed expansion of cells expressing FGFR1 and
FGFR2 in the periportal ductular reaction. Quantitative PCR analysis of the non-hepatocyte cell
xiii
fraction demonstrated significant up-regulation of genes encoding FGFR2IIIb ligands: Fgf7,
Fgf10, and Fgf22. Over-expression of Fgf10 increased the number of cells co-positive for HPC
marker A6 and hepatocyte marker HEPATOCYTE NUCLEAR FACTOR-4 α (HNF4 α) while
reducing serum total bilirubin. Dominant-negative FGFR pathway inhibition reduced the number
of periportal cells exhibiting AKT-dependent activation of β-catenin, and changed the
distribution of cells co-expressing A6 and HNF4 α by reducing dual positive cells near the central
vein of the hepatic lobule while increasing them near the portal vein. Co-treatment of Fgf10
over-expressing DDC-treated mice with AKT inhibitor Wortmannin similarly changed the
distribution of A6/HNF4 α co-positive cells. Conclusion: FGF signaling regulates the expansion
of early postnatal hepatic progenitor cells and DDC-induced A6-expressing hepatocytes partly
through AKT-dependent activation of β-catenin.
1
Chapter 1: Introduction
1 Liver Development
The liver performs many diverse and necessary functions in both fetal development and
postnatal life. During embryonic development, the liver is the primary site of hematopoiesis.
Postnatally, the liver is the key site of metabolism, detoxification, and hormone production. The
adult liver is comprised of many cell types including hepatocytes, biliary epithelial cells (BECs),
Kupffer cells, sinusoidal endothelial cells, and hepatic stellate cells.
1.1 Hepatic Specification of Endoderm
Development of the liver in the mouse begins at embryonic day 8-8.5 (E8-8.5) when the
ventral foregut endoderm interacts with the adjacent septum transversum and cardiac mesoderm
[1]. Before morphological changes are seen, hepatic differentiation/specification is seen by
mRNA expression of Albumin (Alb) and α-Fetoprotein (Afp) [1]. The liver bud forms at E9.5
when specified hepatic cells migrate as cords into the septum transversum mesenchyme [2, 3].
These cords of liver cells surrounded by mesenchymal cells will eventually form the hepatic
sinusoids of the liver. The liver mass increases throughout gestation and becomes the site of
hematopoiesis until birth [4-6]. This process is regulated by many complex signaling pathways
which include, but is not restricted to: bone morphogenic protein (BMP), Hepatocyte Growth
Factor (HGF), Transforming Growth Factor (TGF)- β, Fibroblast Growth Factor (FGF), Notch,
and Wingless (WNT) signaling.
2
1.2 Fibroblast Growth Factor Signaling
Fibroblast Growth Factor signaling is an established regulator of embryonic
development, cell fate determination, migration, and proliferation as well as postnatal tissue
repair and wound healing. In the liver, FGF signaling plays a key role in hepatic specification of
the endoderm, liver morphogenesis, and postnatal liver regeneration.
1.2.1 Protein Structure and Function
FGFs comprise a family of 22 polypeptide ligands which bind to 4 promiscuous
transmembrane receptor tyrosine kinases (RTKs) in mice [7]. The extracellular domain of FGF
Receptors (FGFRs) consists of three IgG domains, seven conserved amino acids, and a heparin
binding domain (Figure 1A,B) [8, 9]. Alternative splicing of the Ig domain III is a major
determinant of ligand binding specificity for FGFRs 1-3 [10, 11]. Splice variants are regulated
in a tissue specific fashion with “b” variants expressed in epithelial cells and “c” in mesenchymal
cells [12, 13]. Activation of the FGFRs results in dimerization of FGFRs which requires binding
of heparin or heparin sulfate proteoglycans (HSPG) [14].
While the Fibroblast Growth Factors were named originally for their mitogenic potential
for fibroblasts, not all FGF ligands regulate fibroblast growth. FGF7 specifically is expressed by
fibroblasts to regulate epithelial cell growth and has no mitogenic potential for fibroblasts cells
[15]. FGF10, which has a high protein sequence similarity to FGF7 [16], has higher binding
affinity to heparin than FGF7 [17] and is mitogenic for fibroblasts at high concentrations [18].
Both FGF7 and FGF10 are expressed by stromal cells [19], and have been proposed as
3
stimulatory signals from the progenitor cell niche [20, 21].
Figure 1.1: Generic FGF Ligand and Receptor Protein Structure. A) Generic FGF ligand
protein structure with conserved core region for FGF receptor and heparin sulfate proteoglycan
binding sites. B) Structural domains of FGFRs including Ig domains, transmembrane domain,
and tyrosine kinase domain. FRS-2, fibroblast growth factor receptor substrate 2; PKC, protein
kinase C.
1.2.2 Downstream Signaling Targets
The Mitogen Activated Protein Kinase (MAPK) pathway, which consists of three core
proteins, is one of many signaling pathways downstream of RTK activation [22]. First,
MAPKKK, a serine threonine protein kinase ,phosphorylates a dual specificity protein kinase
(MAPKK). MAPKK in turn phosphorylates another serine/threonine protein kinase (MAPK).
One example of the MAPK pathway consists of RAF as the equivalent MAPKKK, MEK as the
4
MAPKK, and extracellular signal-regulated kinase (ERK) as the MAPK . The
RAS/RAF/MEK/ERK signaling cascade links RTK activation to cytosolic protein kinase cascade
and activation of downstream targets (Figure 1.2).
Downstream signaling activation of the MAPK signaling is controlled by Fibroblast
Growth Factor Receptor Substrate 2 (FRS2). FGF Receptor activation results in recruitment of
the downstream mediator FRS2 α [23] and subsequent assembly of Growth Factor Receptor
Bound Protein 2 (GRB2) and GRB2 Associated Binder 1 (GAB1) [24]. GRB2 also complexes
with Son of Sevenless (SOS), a guanine nucleotide exchange factor, and activates RAS by GTP
exchange after binding to FRS2. RAS then interacts with effecter proteins such as RAF to
activate the downstream signaling cascade (Figure 1.2).
Phosphatidylinositol 3-kinase (PI3K)/protein kinase B (AKT) signaling, another
signaling cascade downstream of FGFR activation, can be activated by several methods. First,
GAB1 can be activated by FRS2 via interaction with GRB2 leading to tyrosine phosphorylation
and downstream activation of PI3K/AKT signaling through the PI3K regulatory subunit p85 [24,
25]. Second, p85 may directly interact with phosphorylated tyrosine residues on FGFRs [26].
Also, RAS may activate the PI3K catalytic subunit p110 (Figure 1.2) [27, 28]. PI3K activation
results in activation of AKT-dependent anti-apoptotic pathways [29].
5
Figure 1.2: FGFR Activated Intracellular Signaling Pathways. Signaling pathways were
simplified to include only those discussed in the text. FGFR activation occurs with formation of
the FGF-heparin-FGFR ternary complex. Activation and auto-phosphorylation of FGFR leads to
activation of intracellular signaling cascades: RAS/MAPK pathway (shown in orange), and
PI3K/AKT pathway (shown in blue). RAS/MAPK pathway activation occurs after GRB2 binds
to FRS2. PI3K/AKT pathway activation is the product of three different routes of activation.
FRS2 can form a complex with Grb2 and Gab1 to phosphorylate p85, or p85 can be directly
phosphorylated by the FGFR. Lastly, RAS can activate the catalytic unit p110. FGF signaling
activation regulates, but is not limited to, cell survival, proliferation, and differentiation.
6
1.2.3 FGFs in Liver Specification
FGF1, 2, and 8 are expressed by the cardiac mesoderm which activates FGFR1 and
FGFR4 expressed on the adjacent ventral foregut endoderm [30]. FGF1 and 2 expression was
sufficient for induction of the endoderm to liver cell fate, while FGF8 contributed to the
morphogenetic outgrowth of the hepatic mesoderm [30]. In the absence of FGF signaling from
the pre-cardiac mesoderm, the ventral foregut endoderm progenitors default is to differentiate
into the pancreas [31]. Working in parallel with cardiac mesoderm expression of FGFs, the
septum transversum mesenchyme expresses BMP4 to induce liver genes and inhibit cells from
differentiating towards the pancreatic lineage [32]. Downstream of FGF signaling, the initiation
of liver specific genes in the foregut endoderm is regulated by Forkhead box proteins A1 and A2
(Foxa1 & Foxa2) [33]. Induction of hepatic genes is regulated by FGF signaling activation of
MAPK independent of PI3K signaling, which regulates tissue growth but is not necessary for the
induction of hepatic gene expression [34].
1.3 Cross-talk between FGF and Wnt/ β-catenin Signaling
Cell cycle regulation is often a result of a complex network of cross-talk between
signaling pathways. Cross-talk between FGF and Wingless (Wnt)/ β-catenin signaling pathways
has been previously established in both homeostatic cell cycle regulation as well as cancer
progression.
1.3.1 Canonical Wnt Signaling
Canonical Wnt signaling is a key pathway regulating embryogenesis, liver development
and liver regeneration. Humans and mice have 19 Wnt ligands and 10 Frizzled (Fz) receptors
7
with canonical β-catenin mediated downstream transcriptional regulation, or non-canonical β-
catenin independent downstream signaling. In the absence of Wnt signaling, β-catenin is
phosphorylated and targeted for degradation by the Axin/Glycogen Synthase Kinase-3 (GSK-
3)/Adenomatous Polyposis Coli (APC) destruction complex. With canonical signaling
activation, the Fz receptor and its co-receptors Lrp5/6 recruit Dishevelled (Dvl) which inhibits β-
catenin’s destruction complex. Disruption of the destruction complex leads to β-catenin
translocation to the nucleus where it interacts with the T-cell transcription factor/Lymphocyte
enhancer factor (Tcf/Lef) transcription factors (Figure 1.3). Non-canonical Wnt signaling is β-
catenin independent but involves many of the same signaling components.
8
Figure 1.3: Canonical WNT Signaling Pathway. Schematic of Wnt signaling components
discussed in text. In the absence of Wnt ligands, the GSK3 β/APC/Axin phosphorylates β-
catenin, targeting it for degradation. In the presence of Wnt ligands, Disheveled (Dsh) inhibits
GSK3 β activity, allowing β-catenin to accumulate in the cytoplasm and translocate to the
nucleus to induce downstream gene transcription via interaction with the TCF/LEF transcription
factors.
1.3.2 MAPK Activation of β-catenin
β-catenin activation downstream of RAS/MAPK pathways regulates various processes
such as proliferation, differentiation, and cell death. Aberrant signaling of these pathways has
been implicated in tumor progression and is therefore a point of investigation for potential
therapeutic targets. Cross-talk between these pathways have been seen in skin [35, 36], prostate
[37], bladder [38], and liver [39].
9
β-catenin signaling activation may be achieved through both direct and indirect methods.
Downstream of FGFR2 and FGFR3, ERK directly phosphorylates β-catenin at Y142,
dissociating it from its membrane-bound protein complexes which increases its cytoplasmic
levels leading to nuclear translocation and activation of canonical WNT signaling [40]. Also,
FGF2/MAPK signaling has been shown to inhibit GSK3 β, a component of β-catenin’s inhibitory
complex, leading to subsequent induction of β-catenin signaling in endothelial cells [41].
Inactivation of GSK3 β is achieved by ERK1/2 directly phosphorylating Serine 9 (Ser9)-GSK3 β
(Figure 1.4) [42, 43].
1.3.3 AKT Activation of β-catenin
Wnt signaling has also been shown to regulate survival of osteoblasts and their
progenitors through activation of the PI3K/AKT signaling cascade via p-Ser9-GSK3 β [42]. In
gliomas, an aggressive brain tumor, AKT phosphorylates and inactivates GSK3 β, leading to up-
regulation of β-catenin activation (Figure 1.4) [44]. PI3K/AKT signaling also increases β-
catenin stabilization and nuclear localization by phosphorylating β-catenin at Serine 552 (p-
Ser552- β-catenin) and reducing N-terminal β-catenin phosphorylation, which targets β-catenin
for degradation (Figure 1.4) [45, 46]. FGF-mediated AKT-dependent β-catenin activation
regulates HPC proliferation during hepatogenesis as well as tumor initiating HPCs in vitro [47].
10
Figure 1.4: Non-Canonical β-catenin Activation. FGF receptor activation leads to activation
of downstream MAPK and PI3K/AKT signaling cascades, which can non-canonically activate β-
catenin. MAPK and AKT inhibit through phosphorylation of GSK3β on Serine 9. AKT can also
directly phosphorylate β-catenin at Serine 552 to stabilize β-catenin. Each of these pathways
results in nuclear translocation of β-catenin and subsequent transcription of downstream targets.
11
1.3.4 Wnt/ β-catenin Signaling in Hepatic Specification
While the role of Wnt/ β-catenin signaling is well established in the expansion of injury
induced HPCs, its role in hepatic specification has not been definitively determined. Cell-
autonomous Wnt/ β-catenin signaling can specify non-hepatic endoderm to a hepatic cell fate
[48]. In zebrafish, wnt2bb expression from the adjacent mesoderm, and subsequent β-catenin
activation is essential for hepatic induction of the endoderm [49]. In contrast, in Xenopus
development, inhibition of β-catenin is important for induction of liver genes and then activation
later on is essential for expansion of the hepatoblast population and liver morphogenesis [50].
This is further supported by in vitro studies which showed that repression of Wnt and Notch
signaling from the endothelial cell niche is essential for hepatic specification of murine
endodermal cells [51]. CBP/P-300 interacting trans-activator-1 (CITED1), a non-DNA binding
transcriptional co-activator of the Wnt signaling pathway, is expressed from E10.5 to E14.5 in
the liver bud and early hepatoblasts before hepatocyte differentiation [52] implicating a key role
for Wnt/ β-catenin signaling during hepatogenesis after liver specification. Further study will be
required to fully understand the temporal regulation of β-catenin activation during liver
specification and hepatogenesis.
1.4 Liver Morphogenesis
Upon specification and liver bud formation, the embryonic progenitor cell population,
called hepatoblasts, proliferate and eventually differentiate into hepatocytes and BECs to form
the liver. Temporal and spatial expression of growth factors and cytokines regulates the
morphogenesis and differentiation of the developing liver. E11.5 expression of Sonic Hedgehog
(SHH) increases hepatoblast proliferation, but down-regulation of its expression thereafter is
12
essential for hepatocyte differentiation [53]. HGF has also been shown to induce hepatocyte
differentiation of hepatoblasts in vitro [54].
After induction of the liver bud, FGFs continue to play a key role in regulating liver
morphogenesis. Fgf10 and Fgfr2b knockout results in reduced liver size, hypoproliferation of
hepatoblasts, and reduced β-catenin activation, indicating a continued essential role for FGF
signaling and non-canonical β-catenin activation for proper liver morphogenesis [55]. FGFR1
and FGFR2 expression is seen in hepatogenesis from E17 through postnatal development [56].
Activation of FGFRs was detected only during early hepatogenesis (E10-E12) and ex vivo
culture of E10 livers with FGF ligands activated β-catenin and regulated hepatoblast potentiality
(FGF 1 and 4), proliferation/survival (FGF8), and differentiation towards hepatocyte cell fate
(FGF8) [57].
After repeated self-renewal to expand the hepatoblast cell population, hepatoblasts
differentiate into hepatocytes or BECs. Hepatoblast cell fate decisions are regulated by Wnt and
Notch signaling. Current study indicates that Wnt activation inhibits Notch signaling and
subsequent BEC differentiation. Hepatoblast-specific knockout of APC, one of the components
of the β-catenin destruction complex, resulted in hypoplasia of the liver concurrent with
reduction in cell proliferation and up-regulation of genes related to growth arrest [58]. Wnt9a,
expressed in sinusoidal endothelial and stellate cells, regulates proliferation of both hepatoblasts,
and hepatocytes, and later maturation of hepatocytes as seen by glycogen accumulation [59].
1.5 Hepatoblasts
While the mid-gestational liver contains the bipotential hepatic stem/progenitor cell
population, their phenotype is not completely elucidated. Cell surface markers on mid-
13
gestational hepatoblasts include: Aminopeptidase N (CD13) [60], C-Met [61], Delta-like 1
homolog (Dlk) [62], Prominin 1 (CD133) [63], Integrin α6 (CD49f)[47], and Epithelial
Cadherin (E-Cadherin) [64]. CD13
+ive
Dlk
+ive
E9.5-10.5 fetal liver cells can form colonies of
Cytokeratin (CK) 19 positive biliary epithelial cells and Albumin-positive hepatocytic cells [65].
Hepatoblasts derived from the ventral foregut endoderm proliferate to form the liver bud
and expanding liver during embryonic development. These cells express markers for
hepatocytes (AFP and ALB), as well as markers for mature biliary epithelial cells (CK19) [66].
In the developing liver, a subset of hepatoblast express SRY-related HMG box transcription
factor 9 (SOX9) which are the precursors to biliary epithelial cells [67]. SOX9
-ive
hepatoblasts
outside of the ductal plate differentiate towards hepatocytes [68, 69]. Zonation of the
hepatocytes occurs during postnatal development, in which the expression of metabolic enzymes
expressed in hepatocytes varies along the porto-central axis [70]. The SOX9
+ive
cells which lie
around the portal mesenchyme form the ‘ductal plate’ which will differentiate into BECs,
forming 1-2 bile ducts per portal tract [71]. Ductal plate cells which do not form BECs
demonstrate HPC properties and differentiate into periportal hepatocytes as well [72].
1.5.1 Hepatoblast Cell Niche
The embryonic liver contains a variety of cells that provide mitogens and cellular niche
for hepatoblasts. After liver bud induction, cords of hepatic progenitor cells expand and
intercalate with the septum transversum mesenchyme. The developing liver is also the site of
hematopoiesis. Hematopoietic stem cells (HSCs) migrate to the fetal liver around E11.5 where
they undergo self-renewal and differentiation between E12.5-16.5 [73]. After birth, HSCs then
migrate from the liver to the bone marrow. During this early developmental stage, fetal liver
progenitors provide the critical stem cell niche and supportive cytokines for HSC expansion [74].
14
While hepatoblasts have been established as the stem cell niche for HSCs the mitogenic potential
of HSCs for hepatoblasts has yet to be determined.
The mesenchymal cell populations have been the main focus of study as the embryonic
hepatic progenitor cell niche. Hepatic mesothelial cells, which are precursors to hepatic stellate
cells, in periphery provide mitogens for hepatoblasts [75]. HPCs co-cultured with mouse
embryonic fibroblasts showed improved survival indicating a potential niche role [65]. Fgf10 is
expressed from the mesenchymal cell compartment which activates β-catenin in the epithelial
hepatic progenitor cells to regulate cell proliferation and survival [55]. Interestingly, Fgf10 is
expressed during the early stages of hepatogenesis, through E13.5, and then Fgf7 is expressed
from E14.5 through E16.5 which could indicate that these growth factors may be regulating
different aspects of hepatoblast proliferation and differentiation [55]. The Fgf10-expressing cells
are located in close proximity to the hepatoblast cell population in the developing liver, and in
the extrahepatic bile duct. Interestingly, regulation of β-catenin in the mesenchymal cell
compartment also plays a key role in regulating the hepatoblast proliferation as seen by
hypoplasia of the liver when β-catenin is knocked out in the mesenchymal cells [76]. This is also
coupled with a decrease in PAN-CYTOKERATIN
+ive
cells indicating disruption of the
hepatoblast cell niche and perhaps a delay or disruption of BEC differentiation.
1.6 Biliary Epithelial Cell Differentiation
In both murine and human hepatogenesis, Notch signaling is a critical regulator of
hepatoblast and postnatal hepatic progenitor cell differentiation to biliary epithelial cells. Notch
signaling is an intracellular signaling pathway comprised of four transmembrane Notch receptors
(Notch 1-4) which bind mainly to transmembrane Delta-like ligand (DLL) or Jagged ligands.
15
Ligand-receptor binding catalyzes a series of proteolytic events mediated by γ-secretase to cleave
the Notch intracellular domain (NICD) (Fig 1.5). The NICD translocates to the nucleus where it
binds to recombination signal binding protein immunoglobulin kappa J (RBP-J) to activate
transcription of target genes such as the Hes or Hey family of genes (Figure 1.5) [77]. In the
pancreas, the Notch pathway is induced by Fgf10, maintaining epithelial progenitor cells in a
progenitor-like state [78, 79].
Intrahepatic bile ducts (IHBDs) which are located adjacent to portal veins within the liver
develop as a consequence of hepatoblast differentiation and morphogenesis. Periportal
hepatoblasts first form the “ductal plate” which is a periportal sheet of biliary precursor cells
around the portal vein mesenchyme (PVM). The ductal plate cells form BECs, adult HPCs, and
periportal hepatocytes [80]. Although the exact time point of hepatoblast commitment to the
biliary lineage is still unclear, periportal hepatoblasts express the biliary marker SOX9 as early
as E11.5 which indicates that specification of these cells occurs earlier [67]. Differentiation of
the ductal plate cells occurs sequentially with the portal cells differentiating before the cells on
the parenchymal side [67, 81].
16
Figure 1.5: Notch Signaling Pathway. The Notch signaling pathway consists of two Delta and
four Jagged transmembrane ligands which bind to four transmembrane Notch receptors. Ligand-
receptor binding results in proteolytic cleavage of the extracellular domain by Metalloprotease
and intracellular domain by γ-secretase. Cleavage of the Notch intracellular domain (ICD) leads
to its nuclear translocation where it interacts with RBP-J to activate transcription of target genes
such as those of the Hes and Hey family.
Notch signaling is both necessary and sufficient to differentiate hepatoblasts into biliary
epithelial cells. Conditional deletion of RBP-J reduces the number of biliary epithelial cells at
E16.5, indicating an essential role for Notch signaling in hepatoblast differentiation [81].
17
Another study followed the long term effects of in vivo knockout of RBP-J which resulted in loss
of intact IHBDs during postnatal life [82]. This is further supported by the clinical disease
Alagille syndrome, an autosomal dominant disorder caused by mutations in JAGGED1 (JAG1)
or occasionally NOTCH2 which is characterized by paucity of the IHBDs [83-85]. Notch
signaling has also been shown to be sufficient to differentiate hepatoblasts into BECs.
Constitutive expression of NOTCH1 and NOTCH2 induced ectopic biliary cell differentiation
and tubule formation [81, 82, 86]. Interestingly, Notch2 activation regulates hepatoblast
differentiation into BECs independent of Hes1 transcriptional regulation, one of the downstream
targets of Notch signaling [87].
Notch activation occurs via cell-cell contact with the portal vein mesenchyme which
expresses JAG1 early on [81, 88-93]. Activation of Notch signaling in the first layer of ductal
plate cells differentiates them towards BECs and subsequently induces their own expression of
JAG1 which activates the adjacent layer of ductal plate hepatoblasts [81]. The signaling
pathways which regulate the position of bile duct formation have yet to be determined.
TGF β [67] and Wnt/ β-catenin [58, 94] have also been shown to regulate duct
morphogenesis. TGF β ligands are expressed from the portal mesenchyme and bind to TGF β
receptor positive parenchymal ductal cells which regulates the down-regulation of hepatocyte
genes, and up-regulation of biliary epithelial cell genes to promote differentiation [67]. During
normal development β-catenin expression is seen in the ductal plate during BEC differentiation
[58]. Hepatoblast specific knockout of the β-catenin destruction complex protein APC results in
partial differentiation of hepatoblasts towards the BECs independent of the portal vein [58].
Knockout of β-catenin from hepatoblasts results in paucity of the primitive bile ducts, as well as
disruption of hepatocyte maturation [94].
18
1.7 Hepatocyte Differentiation
Hepatocyte differentiation and maturation requires a complex network of signaling as
seen with biliary cell differentiation. Ex vivo culture of E10 livers with FGF8 showed a decrease
in the BEC marker, CK19, which indicates that FGF signaling may promote differentiation
towards hepatocyte precursors [57]. Inhibition of Notch signaling via γ-secretase inhibitors
promotes hepatoblast differentiation into hepatocytes [95]. Murine embryonic stem cells (ESCs)
are treated with HGF, FGF, and TGF in culture to differentiate them towards hepatocytes [96].
Exogenous expression of E-cadherin in ESCs resulted in earlier or higher expression of
hepatocyte markers, indicating that stabilization of E-cad enhances hepatic differentiation [96].
HNF4 α is considered the master regulator of hepatocyte differentiation by regulating hepatocyte
specific-gene expression for many of the metabolic proteins expressed in hepatocytes [97-100].
Wnt/ β-catenin is a key regulator of hepatocyte differentiation and maturation. Knockout
of APC in the liver leads to hyper-activation of β-catenin resulting in loss of hepatocyte
differentiation during embryonic development [58]. Hepatoblast-specific knockout of β-catenin
results in increased hepatocyte apoptosis due to oxidative stress and impairs hepatocyte
proliferation and maturation [94]. This is further supported by β-catenin anti-sense studies which
showed that disruption of β-catenin signaling results in a more immature hepatocyte phenotype
as seen by increased c-kit expression in hepatocytes [101]. A more in depth study of β-catenin
regulation of hepatocyte differentiation was performed by Matrigel® induction of HPCs to
hepatocytes [102]. This study found that differentiating hepatocytes had decreased β-catenin
degradation but did not have a subsequent increase in β-catenin activation. Instead, an increased
association of β-catenin with the HGF receptor, c-Met, was seen, indicating that an increase in
membrane localization of β-catenin is important for hepatocyte differentiation [102]. Also,
19
APC/ β-catenin signaling is an established regulator of liver zonation, regulating the expression
of metabolic proteins across the hepatic lobule [103].
2 Hepatic Progenitor Cells in Liver Disease
In the adult liver, hepatic progenitor cells, otherwise termed ‘oval cells’ reside in the
space between terminal biliary duct branches and adjacent hepatocytes, called the Canals of
Herring. During postnatal liver injury, damaged liver mass is restored via the enormous
proliferative capacity of hepatocytes. In the event that hepatocytes cannot proliferate in response
to injury, the normally quiescent HPCs are activated.
2.1 Cholestatic Liver Disease
Cholestasis is characterized by the inability of bile to flow from the liver into the
duodenum. Cholestasis is generally a product of either obstructive cholestasis in which there is a
mechanical blockage of bile flow, or metabolic cholestasis in which bile formation is disrupted
by either a genetic defect or acquired as a medication side effect. The buildup of bile in the liver
results in fibrosis, and eventually cirrhosis.
Primary sclerosing cholangitis (PSC) which is associated with inflammatory bowel
disease results from inflammation and progressive obstruction of both intra- and extra-hepatic
bile ducts [104]. The cause of primary biliary cirrhosis (PBC) is still unknown, but has been
characterized as an autoimmune disorder resulting in the progressive destruction of the small
intrahepatic bile ducts [105]. Human patients with fulminant hepatic failure and acute hepatitis
have elevated serum levels of FGF7, which may play a key role in regulating the HPC response
seen in both human and rodent liver diseases [21].
20
One avenue of potential therapeutic targets for treatment of cholestatic liver disease is to
target bile acid synthesis and bile acid receptors. FGF signaling is a potential therapeutic target
since studies have shown FGF15 (or FGF19 in humans) inhibits transcription of cholesterol
7alpha hydroxylase (CYP7A1), a key regulator of bile acid synthesis, resulting in inhibition of
bile acid synthesis [106].
2.1.1 Developmental Cholestatic Disease
The majority of pediatric liver diseases progress to liver fibrosis and eventually cirrhosis
[107-109]. Liver fibrosis is the phenomenon in which the liver parenchyma is destroyed and
replaced by scar tissue [110, 111]. Pediatric liver diseases are characterized by different patterns
of fibrosis which expands around the pericentral areas towards the portal spaces, or extends from
the portal vein towards the central vein [112, 113]. Herein, we will discuss biliary atresia which
is a cholestatic liver disease seen in infants.
2.1.1.1 Biliary Atresia
Biliary atresia (BA) is a multifactorial fibro-obliterative pediatric disease of the extra-
hepatic bile duct. BA is uncommon, occurring in only 1 in 10,000-15,000 live births, but is the
leading cause of death for liver failure and liver transplantation in children. Surgical intervention
via the Kasai procedure is performed to improve bile acid drainage from the liver. The cause of
BA is currently unknown, but potential etiologies include viral infection, exposure to toxins,
immunologic dysregulation, and defects in prenatal circulation, or biliary morphogenisis [114].
Left untreated, BA leads to liver failure and its associated complications is uniformly fatal. The
molecular mechanisms which regulate BA progression still remain unclear, but histological
21
analysis of the extra hepatic bile ducts (EHBD) revealed persistent HES1 expression, indicating
that Notch signaling may play a key role [115].
The histological features of BA include cholestasis, biliary epithelial cell proliferation,
and eventually loss of ducts and malformation of ducts to a ductal plate like formation. The
ductular reaction seen with BA is also associated with expansion of ductal HPCs [116]. A single
case-report of transplanted human fetal liver cells in a BA patient showed improved serum
bilirubin and increased liver cell function two months after treatment, indicating that HPCs may
be utilized to manage biliary atresia [117].
2.1.2 Animal Models of Cholestatic Liver Disease
A number of animal models have been established to study cholestatic liver diseases
which utilize feeding of toxins, surgical obstruction of the extra hepatic bile duct, or genetic
manipulation to induce bile duct injury and cholestasis. Although the mechanism of cholestatic
liver injury is not yet fully understood, lineage tracing of biliary cells and fibroblasts with
surgically and chemically induced cholestatic injury revealed epithelial-to-mesenchymal
transition (EMT) and mesenchymal-to-epithelial transition (MET) does not contribute to the
progression of liver fibrosis [118].
2.1.2.1 Murine Rotavirus-Induced Biliary Atresia
Neonatal mice injected with rhesus rotavirus (RRV) are utilized to study the molecular
mechanisms which regulate BA progression. The RRV model of BA is performed by
intraperitoneal injection of virus at P0 in Balb/c mouse pups. Analysis of RRV presence in
extrahepatic BECs 3 days after injection revealed RRV uptake [119]. While RRV can destroy
BECs, it is not the sole mechanism of injury with this model. Interferon-gamma (IFN γ) is a key
22
pro-inflammatory cytokine mediator of RRV-induced BA. Mice lacking IFN γ develop jaundice
after RRV infection, but resolve their symptoms with long-term improved weight gain and
survival [119]. In vitro immortalized and primary human BECs showed similar infectivity as
murine BECs indicating that rotavirus infection may play a key role in the pathogenesis of BA
[120]. RRV-induced BA correlates with MAPK activation and in vitro analysis of MAPK
inhibitors on RRV infected cholangiocytes reduced cholangiocyte injury and viral replication
[121]. The signaling mechanisms and contribution of hepatic progenitor cells in biliary atresia
pathogenesis and repair still require much study in the RRV-induced BA model.
2.1.2.2 Bile Duct Ligation
Bile duct ligation (BDL) is a surgical model of liver fibrosis and cholestatic liver injury
in which the common extrahepatic bile duct is surgically tied closed to prevent drainage of bile
into the intestines. BDL induces BEC proliferation, periportal fibrosis, inflammation, and
expansion of HPCs. Omori, M et al. found the bile duct epithelium of young rats had a
phenotypically similar HPC response to BDL compared to older rats, indicating that younger
rodents should be utilized to explore the HPC reaction to cholestatic liver injury [122]. The
molecular mechanisms which regulate the activation and expansion of these cell populations are
not fully understood.
Expansion of the HPC population is partly dependent upon Kuppfer cell activation
independent of BEC proliferation and myofibroblast transformation with BDL [123]. Recent
study showed FGF7 expression in the Thy1
+ive
mesenchymal cells after BDL in the periportal
HPC niche [21]. Lineage tracing demonstrated that Forkhead Box 1 (FOXL1) expressing cells
which arise in the ductular reaction replace both BECs and hepatocytes, indicating contribution
23
of a bipotential HPC to cholestatic liver injury [124]. Wnt signaling is a key regulator of
hepatices stellate cell activation and thus the progression of liver fibrosis during BDL induced
cholestatic liver injury [125].
2.1.2.3 DDC-Induced Cholestatic Liver Injury
Feeding of 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC) to mice is an efficient
model in which to study cholestatic liver injury and biliary fibrosis. Treatment with this
porphyrinogenic drug increases biliary secretion of porphyrin IV resulting in small bile duct
obstruction as a result of protoporphyrin plug formation. The model is non-lethal, and removal of
the diet allows for recovery/regeneration and resolution of disease, making it an easy model to
work with to study both cholestatic liver injury and mechanisms of regeneration.
Obstruction leads to bile duct proliferation, infiltration of inflammatory cells, and biliary
fibrosis [126]. DDC liver injury also expands HPCs expressing SOX9 [127], CD133 [128], and
A6 [129]. Lineage tracing of the SOX9
+ive
cells which originate in the biliary ductal reaction
indicates that these cells expand out into the parenchyma to restore lost liver mass [127]. While
the precise signaling pathways which regulate the HPC expansion is not yet fully understood,
there is evident redundancy of signaling to regulate the HPC reaction in response to DDC liver
injury.
Similar to what is seen with BDL, FGF7 is expressed in the Thy1
+ive
periportal
mesenchymal cells and regulates the HPC expansion with DDC injury [21]. Over-expression of
Fgf7 reversed hepatocyte injury and cholestasis and increased the number of CK19
+ive
and
A6
+ive
CK19
-ive
HPC populations with DDC injury [21]. Canonical Wnt/ β-catenin is also up-
regulated with DDC injury [130] and hyper-activation of β-catenin reverses hepatic injury and
24
cholestasis while increasing expansion of the A6
+ive
hepatocytes [131]. The expansion and
distribution of the A6
+ive
hepatocytes throughout the hepatic lobule is also partly mediated
through HGF/c-met signaling [132]. The potential cross-talk between all of these signaling
pathways has not yet been addressed, but may provide insight into future therapeutic targets for
cholestatic liver disease.
2.2 Current Liver Disease Therapy
Currently the only treatment for end-stage liver disease is whole liver transplantation.
Liver transplantation is costly, requires lifelong immunosuppression, and is limited by the lack of
donor organs. Research has turned to cell therapies as a potential treatment or as a way to bridge
patients until they can receive a transplant. Primary hepatocytes are being considered as
potential sources for cell therapy [133], as well as mesenchymal stem cells [134]. Current cell
therapies are restricted by cell viability, cell number, as well as sufficient hepatic differentiation
of stem cells in order to be safe for clinical application. Greater understanding of the signaling
pathways which regulate hepatic progenitor cells and hepatocytes during development and liver
injury may provide insight into therapeutic targets and improve future cell therapies.
25
Chapter 2: Over-Expression of Fgf10 Expands the
Periportal Hepatic Progenitor Cell Population in Early
Postnatal Liver
2.1 Abstract
Background: Fibroblast Growth Factor (FGF) signaling is an established key regulator of
induction of hepatogenesis and embryonic hepatic progenitor cell survival and proliferation. To
study the effects of FGF signaling on early postnatal HPC expansion, we utilized Fgf10 over-
expressing mice. Hypothesis: We hypothesized that early postnatal over-expression of Fgf10
would expand the hepatic progenitor cell (HPC) population. Methods: Rosa26
rtTA/-
tet(o)-
Fgf10
+/-
mice were induced with Doxycycline from postnatal day 7 (P7) to 21 (P21).
Immunohistochemistry (IHC) and immunofluorescence (IF) were performed to analyze the
periportal cell phenotype at P7, P21 Control, and P21 Fgf10 induced livers. Results:
Histologically, a cluster of small periportal cells persists postnatally until P7 and thereafter
disappear. A subset of P7 and Fgf10 induced periportal cells express FGFR1 and FGFR2. The
periportal cell population is a heterogeneous mix of CD45
+ive
and E-CADHERIN
+ive
cells which
lack mesenchymal cell markers (DESMIN, αSMA). A subset of the cell population is
ALBUMIN
+ive
, HNF4 α
-ive
, and CK19
+ive
. IF staining for HPC markers revealed a subset of the
cell population to be CD49f
+ive
and CD133
+ive
.
Immunohistochemistry of the NOTCH-1
intracellular domain showed marked nuclear localization within the Fgf10-induced periportal
cells suggesting possible cross-talk between the FGF and Notch signaling pathways in the
regulation of this cell population. Conclusion: The fetal HPC population persists for a week
after birth and its attenuation can be disrupted by Fgf10 over-expression.
26
2.2 Introduction
Induction of the liver bud from the ventral foregut endoderm begins at E8.5 which is
mediated by expression of Fibroblast Growth Factors (FGFs) from the adjacent pre-cardiac
mesoderm [30]. In the absence of FGF signaling, the foregut endoderm’s default differentiation
is towards the pancreatic lineage [31]. Definitive cell surface markers for the hepatic progenitor
cell population are still a point of much debate. Currently A6 [129], CD49f [135], CD133 [128],
Epithelial Cell Adhesion Molecule (EpCAM) [136, 137] , SOX9 [127], and 1C3 [127] have been
established as markers of the hepatic progenitor cells under various homeostatic and injury
conditions.
Fibroblast Growth Factor (FGF) signaling is comprised of 22 murine ligands and 4
transmembrane tyrosine kinase receptors each which has two splice isoforms [7]. We previously
demonstrated that FGF10/FGF Receptor (FGFR)2b signaling regulates the survival of embryonic
hepatic progenitor cells partially through β-catenin activation [55]. In vitro analysis of
hepatoblast and tumor initiating cells revealed FGF7/10-mediated AKT-dependent β-catenin
activation regulates cell survival and proliferation [47].
Recent studies have shown that FGF7 plays a key role in regulating both hepatic
progenitor cells and hepatocytes during postnatal liver injury. In cholestatic liver injury models,
FGF7 is expressed from the mesenchymal cell population in the periportal region to regulate the
expansion of the hepatic progenitor cells [21]. Over-expression of FGF7 reduced hepatocyte
damage and cholestasis with 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC) induced
cholestatic liver injury [21]. During liver regeneration following partial hepatectomy, FGF7
regulates the proliferation of the hepatocyte cell population [138]. While these data show that
27
FGF signaling plays a key role in liver development and regeneration, the de novo role of FGF
signaling on hepatic progenitor cells has yet to be determined.
Further investigation into FGF signaling regulation of hepatic progenitor cells could
provide insight into potential therapeutic targets for liver disease as well as for the expansion of
these cells in culture for cell therapy. In this study, we investigate the role of FGF10 signaling as
a regulator of early postnatal hepatic progenitor cells.
2.3 Methods and Materials
2.3.1 Experimental Animals
Wild type (WT) mixed background mice were utilized for the early postnatal time course
analysis. CMV
cre
;Rosa26
rtTA/-
;tet(O)-Fgf10
+/-
(Fgf10 induced) and CMV
cre
;Rosa26
rtTA/-
;tet(O)-
sFgfr2-IIIb
+/-
(dnFGFR) mice were bread on a mixed background as well. Non-inducible
mothers were fed Doxycycline chow (Harlan Teklad) so that pups received Dox through the
breast milk postnatal day 7 (P7) through P17 for Fgf10 induced mice and P0 through P7 for
dnFGFR mice. Lineage tracing of CD133 cells was performed with Tamoxifen injection of
Prom1-creER
T2
-IRES-nlacZ;Rosa26-YFP (Prom1
+/C-L
;Rosa26-YFP) mice [139] during early
postnatal development.
2.3.2 Tissue Collection
After carbon dioxide euthanasia, 1X phosphate buffered saline (PBS) was flushed
through the portal vein. Portions of the right lobe were collected for histology, RNA, and
western blot analysis. Hepatocyte/non-hepatocyte single cell fractionation was performed by
mechanically mincing the remaining liver followed by enzymatic digestion and serial
centrifugation to isolate the hepatocyte and non-hepatocyte fractions (NHF).
2.3.3 Immunohistochemistry and Immunofluorescence Analysis
28
Tissue collected for histological analysis was fixed in 4% paraformaldehyde (PFA) or
30% sucrose for four hours. 5 μm tissue sections were cut using a Leica RM 2235 microtome,
dried, deparaffinized with histochoice, and rehydrated with ethanol and water. Antigen retrieval
was performed with Tris EDTA unmasking solution. Tissue was permeabilized with 1x TBST
with 0.05% Triton at room temperature before overnight primary antibody incubation (Table
2.1). Immunohistochemistry was performed using EnVision+ Dual Link System HRP (DAB+)
(DAKO Cytomation, CA, USA) according to manufacturer protocol. Immunofluorescence
utilized secondary antibodies (Table 2.1) incubated for 1 hour at room temperature. Slides were
mounted with Vectashield containing DAPI (Vector Laboratories, CA, USA) and images were
taken with a Leica DM 5500 microscope with LAS-AF software (Leica, Vetzlar, Germany).
Table 2.1: Antibodies for Immunostaining
1
o
Antibody Company Species Dilution
A6 Gift from Dr. Factor rat 1:100
Albumin Sigma rabbit 1:100
alpha-SMA Sigma rabbit 1:100
BEK (FGFR2) Santa Cruz Biotech rabbit 1:100
CD45 eBiosciences rat 1:50
CD49f eBiosciences rat 1:100
CD133 eBiosciences rat 1:50
CK19 Gift from Dr. Friedman rabbit 1:100
Desmin Sigma rabbit 1:100
E-Cadherin Santa Cruz Biotech rabbit 1:100
FLG (FGFR1) Santa Cruz Biotech rabbit 1:100
HNF4alpha Santa Cruz Biotech rabbit 1:100
Pan-Cytokeratin Sigma mouse 1:100
pHistoneH3 Cell Signaling rabbit 1:100
PCNA Vector mouse 1:100
2.3.4 Statistical Analysis
29
ANOVA-Post hoc Fisher’s PLSD test or Mann-Whitney Rank Sum was performed using
Statview software (SAS Institute, Inc. Cary, NC) to calculate statistical significance. p<0.05 was
considered as significant.
2.4 Results
2.4.1 The periportal cell population which normally declines during early postnatal
development is retained and expands in the presence of Fgf10 over-expression
During embryonic development, the ductal plate forms a two cell layer in which a subset
of the hepatoblasts differentiates into the biliary epithelium. During early postnatal development,
a single cell layer is retained around the developing biliary epithelium which expands through
postnatal day 7 (P7) before it declines and is gone histologically by P21 (Figure 2.1). During
embryonic development, FGF signaling has been shown to be a key regulator of hepatoblast
proliferation and survival [55].
Figure 2.1: Early Postnatal Periportal Cell Population Peaks at Postnatal Day 7. Early
postnatal periportal cells start out as a two layer of cells called the ductal plate seen at embryonic
day 18.5 (E18.5). The periportal cell population around the bile ducts (BD) increases throughout
postnatal day 7 (P7). This cell population characterized by a high nuclei-to-cytoplasmic ratio
declines after P7 and is gone histologically by P21 (40x).
Transgenic CMV
cre
;Rosa26
rtTA/-
;tet(O)-Fgf10
+/-
(Fgf10 induced) mice were induced from
P7 through P21 and compared to littermate controls and P7 livers to assess the effects of ectopic
Fgf10 expression on the periportal cell population (Figure 2.2A). Histological analysis revealed
30
that the periportal cell population which normally attenuates from P7 to P21 is retained and
expanded (Figure 2.2B). Assessment of the periportal cell population was quantified along the
portal tree as quantification of the ratio of the periportal cell area (PCA) to portal vein area
(PVA). The PCA/PVA ratio was and significantly increased in P21 Fgf10-induced livers
compared to P21 Control and P7 livers (Figure 2.2C, p<0.05).
Figure 2.2: Fgf10 Over-Expression Retains the P7 Periportal Cell Population.
A) Schematic of Doxycycline treatment of CMV
cre
;Rosa26
rtTA/-
;tet(O)-Fgf10
+/-
mice from P7
through P21. B) H&E of the periportal region of P7, P21 Control, and P21 Fgf10 Induced
animals show the periportal cell population seen at P7 normally declines by P21, but is retained
with Fgf10 over-expression (40x). C) The ratio of periportal cell area to portal vein area revealed
a statistically significant increase Fgf10 induced livers compared to P7 and P21 Controls
(p<0.05). Statistical significance was determined by ANOVA with a Fisher’s Post Hoc test
(n=3). PV, Portal Vein; BD, Bile Duct.
31
Inhibition of FGF signaling was performed with CMV
cre
;Rosa26
rtTA/-
;tet(O)-sFGFR2-
IIIb
+/-
mice (dnFGFR) which were induced from P0 to P7 (Figure 2.3A). No histological change
in the periportal cell population was seen between dnFGFR and their littermate controls (Figure
2.3B). Thus FGF signaling is not necessary to regulate this early postnatal periportal cell
population, but it is sufficient to retain it past P7 in the presence of ectopic over-expression of
Fgf10.
Figure 2.3: Dominant Negative FGFR2b Inhibition Does Not Disrupt the Expansion of the
P7 Periportal Cell Population. A) Schematic showing Doxycycline treatment of mice from
postnatal day 0 through postnatal day 7. B) Representative H&E staining from control
littermates and dnFGFR livers shows no change in the periportal cell expansion (20x).
32
2.4.2 The FGF responsive periportal cell population lacks mesenchymal marker expression
In order to confirm that these cells are capable of responding directly to FGF ligands,
non-isoform specific FGF receptor staining was performed. Immunohistochemistry revealed
FGFR2 expression in BECs as well as a subset of the periportal cells at P7 and in the Fgf10
induced animals (Figure 2.4A). Immunofluorescence for FGFR1 revealed a similar expression
pattern of FGFR1 positive BECs and periportal cells at P7 and in Fgf10 induced livers (Figure
2.4B).
Immunofluorescence analysis of the early postnatal periportal cell population has
revealed that is a heterogeneous cell population. Analysis of mesenchymal marker DESMIN
revealed expression of DESMIN
+ive
portal fibroblasts at P7, while Desmin expression after Fgf10
induction revealed cells positive for DESMIN restricted to the portal artery endothelial cells
(Figure 2.5A). Alpha Smooth Muscle Actin ( αSMA) expression was restricted exclusively to the
portal mesenchyme and portal artery (Figure 2.5B). These data combined indicate that Fgf10
over-expression does not induce the expansion of the periportal mesenchymal cell population. A
subset of the periportal cells is CD45
+ive
, indicating infiltration of either hematopoietic cells or of
endothelial progenitor cells (Figure 2.5C). IF staining for epithelial marker Epithelial Cadherin
(E-Cadherin) confirmed that a subset of the P7 and Fgf10 induced cells are epithelial cells
(Figure 2.5D).
33
Figure 2.4: Periportal Cells Express FGF Receptors. A) Immunohistochemistry of FGFR2 in
biliary epithelium and periportal cells (40x). B) IF of PAN-CYTOKERATIN (green) and FGFR1
(red) shows expression of FGFR1 expression in bile duct epithelial cells and periportal cells
(40x). PV, Portal Vein; BD, Bile Duct.
34
Figure 2.5: Periportal Cells are a Heterogeneous Cell Population of Epithelial and
Hematopoietic Cells. A) IF of αSMA (red) and CK19 (red) was restricted to the endothelial
cells. B) IF for DESMIN (red) and PCNA (green) indicates that the mesenchymal cell population
is not expanding in response to Fgf10 (40x). C) IF for hematopoietic marker CD45 (red) with
CK19 (green) shows a subset of CD45
+ive
cells in the periportal region (40x). D) IF for epithelial
cell marker E-CADHERIN (red) marks a subset of the periportal cell population (40x). PV,
Portal Vein; BD, Bile Duct.
35
2.4.3 The FGF responsive periportal cell population expresses hepatic progenitor cell
markers and lack markers for differentiated biliary epithelial cells and hepatocytes
Next, we performed immunofluorescence analysis for hepatocyte (Albumin, HNF4 α),
and biliary epithelial markers (CK19, HNF1 β). P7 and Fgf10 induced periportal cells express
Albumin, but not HNF4 α which indicates they have not fully differentiated into hepatocytes
(Figure 2.6A, B).
Figure 2.6: Periportal Cells Express Albumin but not HNF4 α. A) IF analysis of ALBUMIN
(green) and CK19 (red) expression in the periportal cell population (40x). B) IF analysis of
HNF4 α (green) and CK19 (red) shows the periportal cells are HNF4 α negative (40x).
Hepatic progenitor cell markers analyzed included CD49f, CD133, and A6. CD49f
positive cells were present in the bile ducts and in a subset of the periportal cell population
(Figure 2.7A). The progenitor cell marker CD133 was found in a subset of the P7 and P21 Fgf10
induced periportal cells as well (Figure 2.7B). A6, an established marker of hepatic progenitor
cells was restricted to the BECs at P7 and in P21 control livers (Figure 2.7C). Fgf10 induction
resulted A6
+ive
periportal cells (Figure 2.7C).
36
Figure 2.7: Fgf10 Induced Periportal Cells Express Hepatic Progenitor Cell Markers. A)
Immunofluorescence analysis of CD49f (green) with PCNA (red) reveal a subset of biliary
epithelial cells and periportal cells which are proliferating at P7 and in P21 Fgf10 Induced livers
(40x). B) IF for CD133 (green) with PCNA (red) revealed a subset of cells are positive for
CD133 in P7 and P21 Fgf10 Induced livers (40x). C) IF for the HPC marker A6 (green) overlays
with CK19 (red) ductal cells, and expands in the periportal cell population of Fgf10 induced
livers (40x). PV, Portal Vein; BD, Bile Duct.
2.4.4 Lineage tracing the CD133 positive cell population reveals that they contribute to the
adult hepatocyte cell population
Lineage tracing of the CD133 positive cell population was performed in order to
determine the role of HPCs and their progeny during early postnatal development. The
contribution of hepatic progenitor cells to the hepatocyte cell population during development and
liver injury models is currently under debate. Tamoxifen injection of Prom1
+/C-L
;Rosa26-GFP
37
postnatal day 0 pups was performed to lineage trace the progeny of CD133
+ive
cells during
postnatal development. LacZ staining of the CD133
+ive
cells at P0 revealed them to be in the
biliary epithelial cells as well as a subset of undefined cells around the portal vein which may be
periportal hepatocytes or periportal HPCs (Figure 2.8A). Very few GFP positive cells were
detectable at P1 (Figure 2.8B), but a marked increase CD133 derived CD49f cells were seen at
P5 (Figure 2.8C). CD49f expression in 6 month old livers is minimal, but immunofluorescence
revealed a large number of progeny from the CD133
+ive
cell population (Figure 2.8D).
Further characterization of the CD133
+ive
cells and their progeny was performed to
identify them as hepatocytes or biliary epithelial cells. Co-localization with CK19, a biliary
epithelial cell marker, revealed a subset of the bile duct epithelium at P5 and 6 months to be
derived from CD133
+ive
cells (Figure 2.9A). Hepatocyte Nuclear Factor 4 alpha (HNF4 α), a
hepatocyte transcription factor, co-localizes with the parenchymal CD133 progeny seen in P5
and 6 month old livers (Figure 2.9B).
38
Figure 2.8: CD133
+ive
Cell Progeny Form a Subset of the Liver Parenchymal Cells During
Postnatal Development. A) LacZ staining of CD133
+ive
cells at P0 indicates that biliary
epithelial cells and a subset of cells outside of the bile ducts are CD133 positive. Lineage tracing
of the P0 CD133
+ive
cells (green) was co-localized with hepatic progenitor cell marker CD49f
(red) (B-D,40x). B) Lineage tracing of this CD133
+ive
cell population indicates that the CD49f
+ive
(red) cells at P1 are not derived from CD133
+ive
cells. C) By P5, a subset of the CD49f
+ive
(red)
cell population is derived from the CD133
+ive
cells (green). D) 6 month old adult livers have very
low CD49f expression, but the CD133
+ive
progeny form a large subset of the parenchymal cells.
39
Figure 2.9: CD133 Progeny Form a Subset of Biliary Epithelial Cells and Hepatocytes in
Postnatal Development. A) CD133 progeny (green) co-localized with biliary epithelial cell
marker CK19 (red) reveals a subset of BECs are derived from P0 CD133
+ive
cells (40x). B) A
subset of hepatocytes are also derived from the CD133
+ive
cell population (white arrowhead) as
seen by co-localization with HNF4 α (red) (40x). PV, Portal Vein; BD, Bile Duct.
40
2.4.5 Fgf10 over-expression results in downstream activation of NOTCH1 and β-catenin
Immunohistochemical analysis of the Notch1 Intracellular Domain (NICD), revealed
nuclear localization, an indicator of Notch activation, in periportal cells and periportal
hepatocytes at P7 and in Fgf10 induced models (Figure 2.8A). These data support a potential
cross-talk between FGF signaling and Notch signaling, as similarly seen in the pancreas where
FGF10 resulted in Notch activation and subsequent maintenance of the pancreatic progenitor cell
population [78].
Figure 2.10: Notch1 Signaling is Activated with Fgf10 Over-Expression.
Immunohistochemistry of the Notch1 intracellular domain (brown) shows nuclear localization in
P7 livers and in Fgf10 induced livers indicating activation of Notch1 signaling (20x).
Previous study found that FGF10 and FGF7 act upstream of β-catenin activation to
regulate HPC proliferation and survival [55] by AKT-mediated phosphorylation of Serine-552- β-
catenin [47]. We therefore performed immunofluorescence analysis of pSer552- β-catenin to
determine whether this signaling pathway contributes to the proliferation of the periportal cell
population in the early postnatal model. Interestingly, we found that there are no pSer552- β-
catenin
+ive
periportal cells at P7 nor in the P21 Fgf10 induced livers (Figure 2.11A). P7 livers
demonstrated a small subset of pSer552- β-catenin
+ive
parenchymal cells near the central vein, and
41
even fewer cells in P21 Control and P21 Fgf10 induced livers (Figure 2.11B). This indicates that
while AKT-mediated β-catenin activation is important in the regulation of tumor initiating HPCs
and hepatoblasts [47], this signaling pathway is not activated with Fgf10 induced periportal
HPCs during early postnatal development.
Figure 2.11: AKT-mediated β-catenin activation is not regulated in the early postnatal
periportal cell population. A) IF analysis of pSer552- β-catenin
+ive
(red) cells in the periportal
region shows there are no positive cells during early postnatal development (20x). B) IF analysis
of pSer552- β-catenin
+ive
(red) cells reveals a small subset of positive cells at P7, and loss of
activation of this signaling pathway by P21 in both Control and Fgf10 Induced livers (40x).
Arrows designate positive cells. PV, Portal Vein; BD, Bile Duct; CV, Central Vein.
2.5 Discussion
Specification of the endoderm towards hepatic lineage is regulated by FGF/MAPK
signaling [34]. Expression of FGF ligands from the adjacent pre-cardiac mesoderm activates
FGF receptors expressed on the ventral foregut endoderm to induce liver bud formation [30].
After ventral foregut endoderm specification and liver bud formation, FGF signaling is a key
42
regulator of hepatoblast proliferation and survival as seen by knockout of Fgfr2b and Fgf10 [55].
Knockout of Fgfr2b is an embryonically lethal knockout, so we utilized inducible dominant-
negative FGFR2b expression to study the role of FGF signaling on early postnatal development.
We did not observe a reduction in the expansion of the early postnatal periportal cell population
with FGF signaling inhibition. This indicates either redundant signaling to regulate this cell
population, or that FGFR2b signaling is not necessary for the expansion of this early postnatal
periportal cell population. Fgf10 over-expression from P7 did expand the HPC marker positive
periportal cell population indicating that while FGF signaling is not necessary for their expansion
from P0 to P7, it is sufficient to maintain these cells after P7.
Characterization of the periportal cell population revealed them to express HPC markers
CD49f and CD133. Lineage tracing of the CD133
+ive
cell population starting at P0 showed that
they differentiate into both biliary epithelial cells and hepatocytes. The streaming hepatocyte
theory postulates that hepatic progenitor cells in the periportal regions differentiate into
hepatocytes which then subsequently migrate out towards the central vein in the parenchyma.
Current research suggests that the SOX9
+ive
HPCs contribute to the hepatocyte population in
during postnatal development [140]. Interestingly, lineage tracing of the hepatocytes as well as
SOX9
+ive
HPCs with DDC injury indicates that hepatic progenitor cells do not contribute to
hepatocytes [140, 141]. This is most likely due to the fact that the periportal cells are surrounded
by a thick laminin extracellular matrix which would prevent them from entering the hepatic
cords [142]. Lineage tracing of the CD133 along with the SOX9 study indicates that during early
postnatal development, HPCs which originate in the periportal region are not restricted by the
laminin extracellular matrix and can differentiate into hepatocytes to expand into the
parenchyma.
43
Notch signaling has been previously established as an important regulator of biliary
epithelial cell differentiation [81, 82, 86]. In the pancreas, ectopic FGF10 expression results in
Notch1 activation and subsequent maintenance of pancreatic progenitor cell population by
preventing their differentiation [78]. In our early postnatal liver model, Fgf10 over-expression
also results in nuclear localization of the NOTCH1 ICD, indicating activation of the Notch1
pathway. We also observed expression of CK19 in a subset of the expanded cell population, as
well as expression of hepatic progenitor cell markers: CD49f, CD133, and A6. Further
investigation will be required to determine whether these cells are differentiating towards a
biliary epithelial cell fate, or whether the FGF and Notch signaling is maintaining them in a
progenitor cell like state as seen in the pancreas. Previous study indicatedthat FGF10 acts
upstream of β-catenin activation to regulate hepatoblast proliferation and survival [47, 55], but
this study found that AKT-mediated β-catenin activation does not play a role in regulating the
Fgf10 Induced periportal cell population. Further investigation into the regulation of other
pathways to mediate β-catenin activation should be addressed in the future.
In conclusion, we demonstrate that ectopic Fgf10 expression can expand the early
postnatal hepatic progenitor cell population through modulation of Notch signaling. Further
investigation these signaling pathways and how they regulate hepatic progenitor cells in liver
injury and regeneration could provide insight into improved therapies for liver diseases.
44
Chapter 3: Fibroblast Growth Factor signaling regulates the
expansion of A6-expressing hepatocytes via AKT-dependent
β-catenin activation during DDC-induced liver injury
3.1 Abstract
Background & Aims: We previously established that Fibroblast Growth Factors (FGFs)
promote the proliferation and survival of embryonic hepatic progenitor cells (HPCs) and a
transformed murine HPC line via AKT-dependent β-catenin activation. Recent studies show that
postnatal expansion of A6-expressing hepatocytes during 3,5-diethoxycarbonyl-1,4-
dihydrocollidine (DDC)-induced liver injury is mediated partly by FGF and canonical Wnt
signaling. Herein, we investigate the role of FGF signaling and AKT-mediated β-catenin
activation in acute DDC-induced HPC expansion. Methods: Transgenic mice were fed 0.1%
DDC chow for 14 days concurrent with either inducible hyper-activation of FGF signaling by
Fgf10 over-expression or inhibition of FGF signaling via dominant-negative expression of
soluble FGF Receptor (R)-2IIIb. Results: After 14 days of DDC treatment, we observed
expansion of cells expressing FGFR1, FGFR2, and AKT-dependent β-catenin activation in the
periportal region of DDC-induced ductular reaction. Quantitative PCR analysis of the non-
hepatocyte cell fraction demonstrated significant up-regulation of genes encoding FGFR2IIIb
ligands: Fgf7, Fgf10, and Fgf22. The percentage of periportal cells demonstrating AKT-
dependent β-catenin activation and the number of cells co-positive for HPC marker A6 and
hepatocyte marker Hepatocyte Nuclear Factor-4 α (HNF4 α) increased with over-expression of
Fgf10 and decreased with dominant-negative FGFR pathway inhibition. Inhibition of FGF
signaling also changed the intralobular distribution of A6
+ive (positive)
HNF4 α
+ive
cells resulting in
more cells near the portal vein than near the central vein. The AKT inhibitor, Wortmannin,
decreased Fgf10-induced AKT-dependent β-catenin activation and A6
+ive
HNF4 α
+ive
cell
45
expansion. Conclusion: FGF signaling regulates the expansion of A6-expressing hepatocytes
during DDC-induced liver injury partly through AKT-dependent activation of β-catenin.
3.2 Introduction
When hepatocyte proliferation is impaired during liver injury, there is activation and
expansion of the hepatic progenitor cell (HPC) compartment [143-146]. Upon activation,
previously quiescent HPCs, which reside within the Canals of Hering at the interface between
hepatocytes and biliary ducts, proliferate and expand into the parenchyma to replace lost
hepatocyte mass [147-149]. Human diseases such as congenital biliary atresia and primary
biliary cirrhosis are marked pathologically with expanding biliary ductular reactions, the cells of
which exhibit characteristics consistent with those of epithelial progenitor and stem cells [116,
150]. Treatment of rodents with 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC) induces
similar ductular reactions populated with HPCs expressing Sox9 [127], Prominin-1 (CD133)
[128], and A6 [129].
Fibroblast Growth Factor (FGF) signaling regulates hepatogenesis [30, 34, 55], HPC
expansion, and liver regeneration [21, 151]. The FGF family comprises 22 polypeptide ligands
which bind to 4 promiscuous transmembrane tyrosine kinase receptors [152]. Greater ligand-
receptor binding diversity is achieved via alternative splicing of the third immunoglobulin
domain of FGF Receptors (FGFR)-1 through -3, heterodimerization of FGFR molecules within
the cell membrane, and variable association with heparin sulfate moieties [11]. We previously
demonstrated during early hepatogenesis that FGF10 promotes fetal HPC proliferation via β-
catenin activation [55]. Hepatocyte proliferation following postnatal liver injury is regulated in
part by activation of FGF signaling via FGFR2IIIb [151]. FGF7 regulates HPC expansion and
46
over-expression of Fgf7 reduces hepatocyte damage and cholestatic liver injury induced by DDC
treatment [21].
Wnt/ β-catenin signaling also promotes HPC-mediated liver regeneration [130, 153]. The
binding of Wnt ligand to Frizzled receptor leads to dephosphorylation, activation, and nuclear
translocation of transcriptional regulator β-catenin. Non-canonical activation of β-catenin can
occur via receptor tyrosine kinase (RTK) activation through AKT-dependent phosphorylation of
β-catenin at Serine-552 (pSer552- β-catenin) [40, 46]. FGF7/10-mediated AKT-dependent β-
catenin activation regulates HPC proliferation during hepatogenesis as well as tumor initiating
HPCs in vitro [47]. Postnatal HPC proliferation induced by DDC treatment is mediated in part
via β-catenin activation through increased expression of Wnt ligands [130]. Moreover, hyper-
activation of liver-specific β-catenin during DDC-induced liver injury leads to increased
expansion of A6-expressing hepatocytes from the periportal region into the hepatic lobule in
association with improved hepatic repair and resolution of cholestasis [131]. Recent study has
shown that Notch signaling activation during DDC injury regulates the expression of biliary
epithelial cell markers (A6, Sox9, CK19, and OPN) in hepatocytes [154]. This indicates that
under injury conditions and in the presence of Notch activation hepatocytes transdifferentiate
towards biliary cell lineage.
In this study, we investigate the role of FGF signaling with periportal HPCs and the
distribution of A6
+ive
HNF4 α
+ive
hepatocytes in the hepatic lobule. Furthermore, we elucidate the
relationship between FGF signaling and β-catenin activation in the regulation of the A6
+ive
cell
population with DDC induced liver injury.
47
3.3 Materials and Methods
3.3.1 Experimental Animals and Wortmannin Inhibition
Six-week old, C57BL/6J (wild-type, WT) male mice (Jackson Laboratories) were fed
either a standard diet or 0.1% DDC diet (Test Diet, Richmond). Inducible transgenic and
littermate control mice were given water with 1% doxycycline (Clontech) 2 days prior to and
throughout 14 days of DDC treatment. CMV
cre
;Rosa26
rtTA/-
;tet(O)-sFGFR2-IIIb
+/-
mice
(dnFGFR) induced ubiquitous expression of dominant-negative soluble Fibroblast Growth Factor
Receptor 2-IIIb [155]. CMV
cre
;Rosa26
rtTA/-
;tet(O)-Fgf10
+/-
mice (Fgf10-induced) induced Fgf10
over-expression [156]. In a separate experiment, Fgf10-induced mice were injected daily via
intra-peritoneal route with AKT inhibitor Wortmannin (Fgf10+Wort, 0.7mg/kg dose) or vehicle
control (Fgf10+Vehicle) in DMSO with 1x phosphate buffered saline (PBS) (Sigma-Aldrich),
from two days prior to and throughout 14 days of DDC treatment. Lineage tracing of CD133
cells was performed with Prom1-creER
T2
-IRES-nlacZ;Rosa26-YFP (Prom1
+/C-L
;Rosa26-YFP)
mice [139] treated for 2 months with DDC. All procedures were done in compliance with the
IACUC of Children’s Hospital Los Angeles/Saban Research Institute guidelines for use of
laboratory animals.
3.3.2 Tissue Collection
After carbon dioxide euthanasia, 1x PBS was flushed through the portal vein. Portions of
the right lobe were collected for histology, and total RNA. Hepatocyte/non-hepatocyte
fractionation into a single cell suspension was accomplished by mincing the remaining liver
followed by enzymatic digestion and serial centrifugation as previously described [128] to isolate
the hepatocyte and non-hepatocyte fractions (NHF).
48
3.3.3 Immunohistochemistry and Immunofluorescence Analysis
Collected tissues were immediately fixed in 4% paraformaldehyde (PFA) or 30% sucrose
for 4 hours. 5 μm sections of PFA-fixed, paraffin-embedded and sucrose-fixed, OCT-embedded
tissues were utilized for immunostaining (Table 3.1). Secondary antibodies used were goat-anti-
rabbit-Cy3 (G αRb-Cy3), G αRb-Cy5, G αRat-Cy3, and G αMouse-Cy5 (1:200, Jackson
ImmunoResearch Laboratories, West Grove, PA).
Table 3.1: Antibodies for Immunostaining
1
o
Antibody Company Species Dilution
A6 Gift from Dr. Factor rat 1:100
BEK (FGFR2) Santa Cruz Biotech rabbit 1:100
pSer552- β-catenin Gift from Dr. Li mouse 1:100
FLG (FGFR1) Santa Cruz Biotech rabbit 1:100
HNF4alpha Santa Cruz Biotech rabbit 1:100
pHistone H3 Cell Signaling rabbit 1:100
PCNA Vector Mouse 1:100
Hematoxylin & Eosin (H&E) was utilized for morphological analysis. Quantification of
A6
+ive
HNF4 α
+ive
cells per lobular region was performed by separating each lobular structure into
periportal (0-96 μm), mid (97-194 μm), and central regions (195-290 μm). Immunohistochemistry
was performed with the Dako EnVision+ Dual Link System-HRP (DAB+) kit (Dako, Denmark)
according to manufacture protocol. Immunofluorescence images were taken using a Leica
DM5500B microscope (Leica, Buffalo Grove, IL). All IF quantification was performed on three
images per animal (n=3).
49
3.3.4 Quantitative PCR Analysis of Gene Expression
Total RNA from the NHF was isolated using the Qiagen RNeasy Mini kit (Qiagen;
Valencia, CA). RNA concentration and purity were determined by Nanodrop spectrophotometry
(ND-1000, Thermo Scientific, Wilmington, DE). Complementary DNA (cDNA) was prepared
with iScript cDNA synthesis kit (Bio-Rad, Hercules, CA). Quantitative real-time PCR (qPCR)
was performed using Light-Cycler Taqman Master (Roche Applied Science, Indianapolis, IN)
and probes from the Universal Probe Library (Roche Applied Science, Indianapolis, IN) against
intron spanning, gene specific primers (Table 3.2). 18s was selected as the optimal internal
control for all analyses due to more stable expression levels between treatment groups.
Table 3.2: qRT-PCR Primers
Gene Forward Primer 5'-3' Reverse Primer 5'-3'
Fgfr1b atccgcagcctcacattc aattcccgaatgcttcag
Fgfr2b cctacctcaaggtcctgaagc catccatctccgtcacattg
Fgf1 cagcctgccagttcttcag ggctgcgaaggttgtgat
Fgf3 tgagaacagcgcctatagca gtaccgcccagaaaagagc
Fgf7 tctcatcaatctccagttcacaa cttgcgttggattgctactcct
Fgf10 cgggaccaagaatgaagact aacaactccgatttccactga
Fgf22 gacacgacggcaccaact aggcccttcaagacgagac
18s gcaattattccccatgaacg gggacttaatcaacgcaagc
50
3.3.5 Statistical Analysis
ANOVA-Post hoc Fisher’s PLSD test or Mann-Whitney Rank Sum was performed using
Statview software (SAS Institute, Inc. Cary, NC) to calculate statistical significance. p<0.05 was
considered as significant.
3.4 Results
3.4.1 Expression of FGF ligands and receptors is up-regulated during acute DDC liver
injury
FGF ligands expressed by the mesenchymal cell population regulate HPCs and liver
regeneration during chemically induced liver injury (16, 17). qPCR was performed on RNA
isolated from the non-hepatocyte cell fraction (NHF) to determine FGF ligand and receptor
expression during acute DDC injury. The NHF was collected in order to exclude large
hepatocytes and enrich for HPCs [128], and Fgf-expressing mesenchymal cells [21]. By qPCR,
we observed 11.6-fold and 12.4-fold increase in Fgfr1IIIb and Fgfr2IIIb expression respectively
(Figure 3.1A, p<0.05) suggestive of FGFR1IIIb and FGFR2IIIIb activation [47]. FGF1, FGF2,
FGF3, and FGF10 possess high binding affinity for FGFR1IIIb while FGF1, FGF3, FGF7,
FGF10, and FGF22 exhibit high binding affinity for FGFR2IIIb [157] qPCR was performed in
lieu of western blot analyses given lack of available of specific antibodies against FGF ligands.
Fgf3 was not detectable by qPCR. Fgf1 expression was transiently down-regulated at 3 days but
was significantly upregulated by 14 days of DDC treatment compared to baseline (Figure 3.1B,
p<0.05). Fgf10 expression was significantly up-regulated 14 days after DDC injury while Fgf7
and Fgf22 were upregulated throughout the entire observation period (Figure 3.1B, p<0.05).
51
Using non-isoform specific antibodies for FGFR1 and FGFR2, we observed increased
FGFR1 and FGFR2 expression in expanding populations of periportal cells and hepatocytes at 7
days and 14 days DDC injury (Figure 3.1C,D). Quantification after 3 days and throughout DDC
treatment revealed increased FGFR1
+ive
cells, and increase in the fraction of FGFR1
+ive
PCNA
+ive
cells in the periportal region (Figure 3.1D-F, p<0.05).
Figure 3.1: FGF Signaling is Up-Regulated During DDC Injury. A) Relative expression
levels of Fgfr1IIIb and Fgfr2IIIb from the non-hepatocyte fraction. B) Relative expression
levels of Fgf1, Fgf7, Fgf10, and Fgf22. C) Immunohistochemistry for FGFR2 (brown) in the
52
periportal region at 0d, 7d, and 14d DDC injury (20x) and representative negative control (10x).
D) Immunofluorescence staining for PCNA (green) and FGFR1 (red) at 0d, 7d, and 14d DDC
injury and representative negative control (40x). E) Quantification of the number of FGFR1
+ive
cells per periportal area at 0d, 3d, 7d, and 14d DDC injury. F) Quantification of the % of
FGFR1
+ive
PCNA
+ive
cells per periportal area. Statistical significance was determined by Mann-
Whitney Rank Sum test (n=3, *p<0.05).
3.4.2 Fgf10 promotes the expansion of A6
+ive
HNF4 α
+ive
cell populations
In order to assess the effects of FGF signaling inhibition on HPC expansion, dnFGFR
mice and their non-inducible, control littermates (WT) were co-treated with doxycyline 2 days
prior to and throughout 14 days of DDC diet. Hyper-activation of FGF signaling using Fgf10-
induced using the same treatment scheme. There were neither significant changes in liver-to-
body weight ratios, serum AST, and ALT levels (Table 3.3) nor were there histological changes
in terms of DDC-induced ductular reactions (Figure 3.2A) between WT, dnFGFR, and Fgf10-
induced groups. DDC-treated Fgf10-induced mice did, however, exhibit partial rescue of
cholestasis with a significant reduction in total serum bilirubin levels (Table 3.3).
Table 3.3: Effects of dnFGFR and Fgf10 over-expression on DDC liver injury
Treatment
Liver/Body Weight
Ratio AST (U/L) ALT (U/L)
Total
Bilirubin
(mg/dL)
WT 0.08±0.003 2171.7± 230.3 2336.8± 342.5 17.4± 3.6
dnFGFR 0.07±0.005 2245.2 ± 112.2 2404.3± 458.0 15.5± 2.8
Fgf10 Induced 0.08±0.002 1446.5± 383.8 2588.8± 639.0 5.2± 1.2*
Values are given as mean ± standard error of mean (SEM). Statistical significance was
determined using ANOVA with Fisher’s post-hoc test. * p<0.05 compared to WT and dnFGFR.
Abbreviations: AST, asparatate transaminase; ALT, alanine aminotransferase.
53
β-catenin [130, 131] and Hepatocyte Growth Factor (HGF) [132] signaling are known
mediators of DDC-induced A6
+ive
progenitor-derived hepatocyte expansion into the parenchyma.
The contribution of HPCs to hepatocytes and the subsequent streaming hepatocyte theory has
been a topic of much debate, lineage tracing of SOX9+ HPCs gives rise to a minimal number of
hepatocytes after DDC treatment [127, 140]. Recent evidence suggests that Notch activation and
cholestatic liver injury results in reprogramming of the hepatocytes to express biliary epithelial
cell markers [154]. We utilized the hepatocyte marker Hepatocyte Nuclear Factor-4 α (HNF4 α)
and the HPC marker A6 to investigate the HPC population and hepatocytes transdifferentiating
towards biliary epithelial cells in response to FGF signaling. We observed increases in
A6
+ive
HNF4 α
-ive
HPCs and A6
+ive
HNF4 α
+ive
hepatocytes in Fgf10-induced livers with DDC
treatment (Figure 3.2B-D, p<0.0001 and p<0.05 respectively). Analysis of the distribution of the
A6
+ive
HNF4 α
+ive
hepatocytes within the hepatic lobule was performed by quantifying the cells in
the periportal, mid, and central regions (Figure 3.2E). Although there was an increase in the total
number of A6
+ive
HNF4 α
+ive
cells in Fgf10-induced livers, there was no change in the overall
distribution of A6
+ive
HNF4 α
+ive
cells across the periportal, mid, and central regions of hepatic
lobule compared littermate controls (Figure 3.2F).
Inhibition of FGF signaling during DDC treatment did not affect the number of
A6
+ive
HNF4 α
-ive
HPCs or A6
+ive
HNF4 α
+ive
hepatocytes compared to littermate controls (Figure
3.2B-D). However, dnFGFR mice exhibited altered intralobular distribution of A6
+ive
HNF4 α
+ive
cells with an increased fraction of A6
+ive
HNF4 α
+ive
cells in the periportal region and a reduction
in the central region in comparison to WT and Fgf10-induced mice (Figure 3.2F, p<0.05). This
54
observation suggests impaired transdifferentiation of A6
+ive
HNF4 α
+ive
hepatocytes in the central
region during DDC injury with inhibition of FGF signaling.
55
56
Figure 3.2: FGF signaling regulates A6
+ive
HNF4 α
+ive
cell expansion. A) WT, dnFGR, and
Fgf10-induced H&E staining of periportal regions (20x). B) IF staining for A6 (green), and
HNF4 α (red) for each treatment (10x). C) Quantification of the mean number of A6
+ive
cells per
high powered field (HPF). D) Quantification of mean number of A6
+ive
HNF4 α
+ive
cells per high
powered field (HPF). E) Schematic for quantification of distribution of A6
+ive
HNF4 α
+ive
cells
within the hepatic lobule. The hepatic lobule was broken into thirds: the periportal, mid, and
central regions. Dual positive A6 (green) HNF4 α (red) cells were counted per region to analyze
cell distribution. F) Quantification of percent of A6
+ive
HNF4 α
+ive
cells in periportal, mid, and
central lobular regions. Statistical significance was determined by ANOVA with Fisher’s post-
hoc test (n=3, * p<0.05).
To analyze whether FGF10 signaling regulates expression of A6 in hepatocytes in
uninjured livers, Fgf10 inducible animals were treated with Doxycycline for two weeks in 6
week old mice. Immunofluorescence analysis of A6 (green) and HNF4 α (red) staining revealed
that a small subset of HNF4 α
+ive
cells express A6 following 2 weeks of Fgf10 over-expression
which are not seen in non-inducible control littermates (Figure 3.3).
Figure 3.3: Fgf10 over-expression in non-injured adult livers induces A6 expression in
hepatocytes. Immunofluorescence staining of HPC and BEC marker A6 (green) and hepatocyte
marker HNF4 α (red) revealed dual positive cells in Fgf10 induced livers (denoted by white
arrowheads) (20x). 40x inset shows dual positive cells in Fgf10 induced livers.
57
3.4.3 FGF signaling mediates periportal cell proliferation and AKT-dependent β-catenin
activation
Cell proliferation was quantified by phospho-HistoneH3 (pHisH3) staining. Analysis of
pHisH3
+ive
cells from 20x images around central vein revealed no significant change between
treatment types (data not shown). However, proliferation of the periportal cells (Figure 3.2A,
within the yellow dotted line) exclusive of large hepatocytes but inclusive of A6
+ive
cells, biliary
epithelial cells, mesenchymal cells, and small A6
+ive
HNF4 α
+ive
hepatocytes, was impaired with
inhibition of FGF signaling during DDC injury compared to controls (Figure 3.4A, B, p<0.01).
In contrast, over-expression of Fgf10 during DDC treatment promoted periportal cell
proliferation coinciding with the increase in A6
+ive
HNF4 α
-ive
and A6
+ive
HNF4 α
+ive
hepatocytes
(Figure 3.4A, B, p<0.01).
Given that β-catenin activation regulates the expansion of the A6
+ive
cell population with
DDC injury [153] and that FGF promotes AKT-dependent β-catenin activation of embryonic and
tumor initiating HPC in vitro [47], we sought to determine if FGF signaling induces AKT-
mediated β-catenin activation occurs in DDC liver injury. We observed a ~10% increase in the
fraction of pSer552- β-catenin
+ive
periportal cells in Fgf10-induced livers compared to WT
(Figure 3.4C,D, p<0.0001). In contrast, there was a marked reduction in the fraction of pSer552-
β-catenin
+ive
periportal cells with inhibition of FGF signaling in dnFGFR livers compared to WT
and Fgf10-induced livers (Figure 3.4C,D, p<0.0001, and p<0.01 respectively). Collectively,
these observations indicate that FGF signaling promotes AKT-dependent β-catenin activation in
periportal cells.
58
Figure 3.4: FGF signaling regulates heterogeneous periportal cell population proliferation
and AKT-dependent β-catenin phosphorylation. A) IF staining for cell proliferation by
phospho-Histone H3 (red) (pHisH3) and DAPI (blue) (40x). B) Quantification of the percent of
pHisH3
+ive
cells per periportal area. C) IF staining for pSer552- β-catenin
+ive
(red) cells with
DAPI (blue) per periportal area (40x). D) Quantification of the percent of pSer552- β-catenin
+ive
cells per periportal area. Statistical significance was determined by ANOVA with Fisher’s post-
hoc test (n=3, * p<0.05, # p<0.0001).
3.4.4 Inhibition of PI3K signaling disrupts A6
+ive
HNF4 α
+ive
cell expansion
To further validate the role of FGF signaling in β-catenin-dependent expansion of A6
+ive
cells, Fgf10-induced animals were injected daily with the AKT inhibitor Wortmannin
(Fgf10+Wort) (0.7mg/kg). No significant difference was seen histologically (data not shown) or
with serum AST, ALT, and total bilirubin between Fgf10+Wort and Fgf10+vehicle controls
(Table 4). There was no observed change in the number of A6
+ive
cells between Fgf10+Wort
livers and vehicle controls, indicating that disruption of pAKT signaling does not affect the
overall expansion of the A6
+ive
cell population (Figure 3.5A,B). Fgf10+Wort livers exhibited a
significant reduction in the number of A6
+ive
HNF4 α
+ive
cells indicating that pAKT signaling may
regulate the transdifferentiation of the hepatocytes towards a biliary lineage (Figure 3.5C,
59
p<0.05). The distribution of the A6
+ive
HNF4 α
+ive
cells within the hepatic lobule was also altered,
showing a pattern similar to that seen in dnFGFR livers in which the percent of cells near the
portal vein is greater than that near the central vein (Figure 3.5D, p<0.05). Collectively these
data indicate that FGF/AKT signaling disrupts hepatocyte expression of A6
+ive
cells, an
indication of hepatocyte transdifferentiation towards biliary cells as seen with DDC injury [154]
without affecting the expansion of the A6
+ive
HNF4 α
-ive
HPC population.
Table 3.4: Effects of pAKT inhibition with Fgf10 over-expression on DDC liver injury
Treatment
Liver/Body
Weight Ratio AST (U/L) ALT (U/L)
Total
Bilirubin
(mg/dL)
Fgf10/Vehicle 0.07± 0.004 1173.0± 315.8 2238.3± 15.5 4.6± 1.6
Fgf10/Wort 0.11± 0.028 1604.7± 282.8 1987.7± 439.1 6.7± 2.1
Values are given as mean ± standard error of mean (SEM). Statistical analysis was determined
using the Mann-Whitney Rank Sum test. Abbreviations: AST, asparatate transaminase; ALT,
alanine aminotransferase.
60
Figure 3.5: Wortmannin inhibition of Fgf10 induced livers disrupts the A6+HNF4 α+ cell
differentiation and migration. A) IF staining for A6 (green), and HNF4 α (red) (10x). B)
Quantification of mean number of A6
+ive
cells per high powered field (HPF). C) Quantification
of mean number of A6
+ive
HNF4 α
+ive
cells per high powered field (HPF). D) Quantification of %
of A6
+ive
HNF4 α
+ive
cells in periportal, mid, and central lobular regions. E) IF staining for
pSer552- β-catenin
+ive
(red) cells with DAPI (blue) in the heterogeneous periportal cell population
(40x). F) Quantification of the % of pSer552- β-catenin
+ive
cells per periportal area. Statistical
significance was determined by Mann-Whitney Rank Sum test (n=3, * p<0.05).
61
To determine the effect of Wortmannin treatment on the periportal cell proliferation and
AKT-dependent β-catenin activation, we quantified the percent of pHistoneH3
+ive
and pSer552-
β-catenin
+ive
periportal cells. We observed a ~30% reduction in the percentage of pSer552- β-
catenin
+ive
periportal cells in the Fgf10+Wort livers compared to Fgf10+Vehicle (Fig. 3.5F,
p<0.05). We also observed a 50% reduction in periportal cell proliferation in Fgf10+Wort
compared to Fgf10+Vehicle (Figure 3.6A,B, p<0.01). This data indicates that AKT-dependent
β-catenin activation plays a key role in regulating the A6
+ive
HNF4 α
+ive
cell population during
cholestatic liver injury.
Figure 3.6: Wortmannin inhibition decreases periportal cell proliferation. A) IF staining for
cell proliferation by phospho-Histone H3 (red) (pHisH3) and DAPI (blue) (40x). B)
Quantification of the % pHisH3+ive cells per periportal area. Statistical significance was
determined by Mann-Whitney Rank Sum test (n=3, * p<0.05).
3.4.5 CD133
+ive
Cells Form a Subset of Biliary Epithelial Cells and Hepatocytes with DDC
Injury
Lineage tracing of the CD133
+ive
cells and their progeny was performed on 2 month old
adult livers. In uninjured adult livers, a small subset CD133
+ive
cells are present in the bile ducts
(Figure 3.7A) as well as a small number of hepatocytes (Figure 3.7B). When looking for
CD133
+ive
progeny treated at 2 months of age and collected at 4 months, we found CD133
+ive
biliary epithelial cells and hepatocytes only make up a very small subset of cells (Figure 3.7C,
D). With 2 months of DDC liver injury, the CD133
+ive
progeny are seen to form a small subset
62
of the A6
+ive
HNF4 α
+ive
cell population, indicating that the CD133
+ive
hepatocytes seen at
2months of age may proliferate and may transdifferentiate towards a biliary epithelial cell type
(Figure 3.7E, F).
Figure 3.7: CD133
+ive
cells form a subset of biliary epithelial cells and hepatocytes in non-
injured livers and DDC injured livers. Lineage tracing of in CD133
+ive
cells and their progeny
(green) in 2 month old livers was overlayed with biliary epithelial marker CK19 (red) (A) or
hepatocyte marker HNF4 α (red), and BEC/HPC marker A6 (white) (B). CD133
+ive
progeny in 4
month old adult livers from 2 month old mice (C, D). E) Lineage tracing CD133
+ive
cells in the
CK19
+ive
atypical ductular reaction (red) after 2months DDC treatment. F) Lineage tracing
CD133
+ive
cells in the A6
+ive
(white) HNF4a
+ive
(red) hepatocytes after 2months DDC treatment.
63
3.5 Discussion
FGF signaling is a key regulator of endoderm specification toward a liver-specific fate
[34] as well as embryonic HPC survival and proliferation [47, 55]. Postnatally, FGF signaling
regulates liver regeneration following injury [21, 138, 151]. In this study, we show that FGF
signaling promotes expansion of DDC A6
+ive
HPCs and intralobular distribution of
A6
+ive
HNF4 α
+ive
hepatocytes via AKT-mediated β-catenin activation.
Takase et al. showed that FGF7, expressed by the periportal Thy1
+ive
mesenchymal cells,
induces the expansion of periportal HPCs during adult cholestatic liver injury [21]. The authors
further demonstrated that over-expression of Fgf7 during 3 weeks of DDC injury reversed
hepatocyte injury and cholestasis while increasing the CK19
+ive
HPC population and the A6
+ive
CK19
-ive
periportal cell population [21]. Here we show a similar enhancement of A6
+ive
cell
expansion with over-expression of Fgf10, a member of the FGF7 ligand sub-family [18]. We
observe a reduction in hyperbilirubinemia along with an increased number of pSer552- β-
catenin
+ive
periportal cells with Fgf10 over-expression, consistent with the published observation
that hyper-activation of β-catenin during DDC injury results in reduced bilirubin levels [131]. In
contrast, inhibition of FGF signaling or inhibition of AKT by Wortmannin in Fgf10-induced
mice reduces the number of pSer552- β-catenin
+ive
periportal cells during DDC injury. Over-
expression of Fgf10 during DDC injury does not alter serum transaminase levels indicating a
lack of effect overall on hepatocyte injury although it does increase the total number of
A6
+ive
HNF4 α
+ive
cells, which are likely newly transdifferentiating hepatocytes. We speculate
that the reduction in DDC-induced hyperbilirubinemia is due to the emergence of these
A6
+ive
HNF4 α
+ive
cells, although the mechanism is unclear.
64
Knockout of Fgf7 in the liver leads to impairment of CK19
+ive
HPC proliferation during
DDC injury and is lethal in 80% of mice by 8 weeks compared to nearly zero mortality in WT
mice [21]. In comparison, our study focuses on the role of FGF signaling during the initial two
week induction of HPC. Although inhibition of FGF signaling for two weeks in dnFGFR mice
does not reduce the overall number of either A6
+ive
HNF4 α
+ive
hepatocytes or A6
+ive
HNF4 α
-ive
HPCs, the observed change in intralobular distribution of A6
+ive
HNF4 α
+ive
cells and reduction in
cell proliferation and p-Ser552- β-catenin
+ive
cells indicates a potential disruption of the
transdifferentiation of hepatocytes near the central vein. Similarly, inhibition of AKT by
Wortmannin in Fgf10-induced mice disrupts expansion of A6
+ive
HNF4 α
+ive
hepatocytes. These
data are consistent with published observation of reduced Phosphoinositide 3-Kinase
(PI3K)/AKT activation and A6
+ive
cell expansion with knockout of c-met, the HGF Receptor
gene, [132] or liver-specific conditional knockout of β-catenin [153]. Our data indicate that FGF
and PI3K/AKT signaling are important regulators of β-catenin-mediated A6
+ive
HPC expansion
postnatally as well as the distribution of the A6
+ive
HNF4 α
+ive
hepatocytes in a manner that
recapitulates embryonic HPC and tumor initiating hepatic stem cell expansion [47]. The
incomplete block of A6
+ive
cell expansion in our model of FGF inhibition along with the
aforementioned knockout models may also indicate redundancy in the pathways regulating HPC
expansion and hepatocyte transdifferentiation.
Lineage tracing of SOX9
+ive
HPCs, which originate in the bile ducts, show a hepatocyte
fate as they migrate downstream from the periportal region toward the central vein during normal
homeostasis [140] in a manner consistent with the “streaming hepatocyte” hypothesis [158]. In
response to DDC injury, these SOX9
+ive
cells give rise to biliary epithelial cells, but only
minimally contribute to the hepatocyte cell population [127, 140]. Lineage tracing of hepatocytes
65
with DDC injury revealed that ductal cells did not contribute to newly formed hepatocytes [141]
which may be due to the thick laminin basement membrane seen with DDC injury preventing
HPCs from differentiating escaping into the parenchyma [142]. Similarly, using OPN, the
biliary epithelial cell marker, lineage tracing showed that these cells do not contribute to the
hepatocyte cell population with DDC injury [159]. Instead, the concept of hepatocytes
transdifferentiating towards the biliary lineage has been recently proposed in the DDC model and
in response to Notch activation [154], the signaling pathway which is required for biliary
epithelial cell differentiation during hepatogenesis. Fgf10 over-expression resulted in an
increase in the A6
+ive
HNF4 α
+ive
hepatocytes [55] indicating that FGF signaling also plays a role
in regulating the transdifferentiation of the hepatocyte cell population. Inhibition of FGF
signaling and downstream target AKT also changed the intralobular distribution of these cells
which further supports that FGF signaling regulates the reprogramming of hepatocytes with
DDC injury. Further investigation will be required to analyze the potential cross-talk between
Notch signaling, FGF siganling, and β-catenin activation.
β-catenin activation plays a complex and dynamic role in regulating liver cell
proliferation and metabolic zonation patterning (reviewed in [160]). In vitro canonical Wnt-
mediated β-catenin activation in newly differentiated periportal hepatocytes regulates HNF4 α-
driven transcription to switch cells to a perivenular hepatocyte phenotype [161]. Modulation of
β-catenin activation by FGF signaling may regulate the differentiation and distribution of the
A6
+ive
HNF4 α
+ive
cells in DDC injury.
In conclusion, we demonstrate that FGF signaling regulates the expansion of the A6
+ive
HPCs and A6
+ive
HNF4 α
+ive
hepatocyte-fated cells via AKT-mediated β-catenin activation during
DDC induced liver injury. Further investigation into the signaling pathways which regulate
66
differentiation and migration of the hepatocyte cells from HPCs could provide better
understanding to regulate HPCs and hepatocytes for cell-based therapies for liver diseases.
67
Concluding Remarks and Future Perspectives
4.1 Concluding Remarks
Our previous work on Fibroblast Growth Factor signaling in hepatogenesis showed that
knockout of Fgf10 and Fgfr2b results in atypical morphogenesis and reduced hepatoblast
proliferation and survival [55]. Primary cell culture of hepatoblasts and tumor initiating cells
revealed that FGF10 treatment regulates cell proliferation partly through AKT-dependent β-
catenin activation [47]. In this study, we found a novel early postnatal hepatic progenitor cell
population which expands in the periportal region shortly after birth before it disappears
(Chapter 2). Inhibition of FGF signaling during early postnatal development did not disrupt the
expansion of the periportal HPC population from postnatal day 0 to postnatal day 7 (Chapter 2).
Fgf10 over-expression results in proliferation of this HPC periportal cell population after P7.
Interestingly, ectopic Fgf10 expression in adult uninjured liver does not result in expansion of
HPCs from the Canals of Herring (Chapter 3). Instead, a small subset of hepatocytes express A6,
a HPC marker, in response to Fgf10 over-expression, indicating that ectopic FGF signaling
activation results in reprogramming of hepatocytes towards a biliary cell fate in adult uninjured
liver. Further investigation will be required to determine the downstream signaling mechanisms
which regulate this early postnatal HPC cell population. Preliminary data indicates that Notch
signaling and β-catenin activation may play a key role in regulating this cell population (Chapter
2).
The role of FGF signaling in the regulation of liver disease and regeneration has been
investigated in several mouse models. Following partial hepatectomy, FGF signaling is activated
to regulate the hepatocyte proliferation [56]. In DDC induced cholestatic liver injury, FGF7 is
expressed in the periportal mesenchymal cell population to regulate the atypical ductular reaction
68
and expansion of hepatic progenitor cells [21]. Current research indicates that the periportal HPC
population in DDC induced cholestatic liver injury does not contribute to the hepatocyte cell
population [140, 141]. Instead, hepatocytes most likely proliferate, and undergo reprogramming
towards a biliary epithelial cell fate [154]. Our own study contributed to these findings by
showing Fgf10 over-expression and FGF signaling inhibition in the DDC model regulates the
distribution of A6-expressing hepatocytes (Chapter 3). We found that AKT inhibition and
dominant negative FGFR2b inhibition resulted in a change in the distribution of A6
+ive
HNF α
+ive
cells such that more cells were seen in the periportal region and less near the central vein.
Coupled with a decrease in periportal cell pSer552- β-catenin, this data indicates that downstream
FGF signaling modulates the reprogramming of the hepatocyte cell population in cholestatic
liver injury. Further investigation into cross-talk between FGF, Notch, and β-catenin signaling
activation will provide greater insight into the reprogramming of hepatocytes in DDC induced
cholestatic liver injury.
4.2 Future Perspectives
The studies presented in this thesis on Fibroblast Growth Factor signaling regulation of
hepatic progenitor cells and hepatocytes may lead to additional insights on the following areas of
research: 1) regulation of hepatic progenitor cells and hepatocytes in liver disease 2) cross-talk
between FGF signaling and β-catenin activation.
Hepatoblastoma (HB) is the most common pediatric liver cancer thought to be derived
from hepatic progenitor cells which arrest at various stages of liver development due to the
morphological patterns which are similar to hepatoblasts [162]. Analysis of HepG2 cells, a
hepatoblastoma-derived cell line, revealed mRNA expression of all FGFRs, FGF1, and FGF2,
indicating that FGF signaling may play a role in liver tumor transformation [163]. Classification
69
of HBs is segregated according to histological patterning of the cells [164]. Regardless of type,
HBs are associated with up-regulation of the HPC marker, Dlk [165]. During hepatogenesis,
FGF signaling regulates β-catenin activation as well as the proliferation and survival of
hepatoblasts [55]. The findings from the present study indicate that aberrant FGF signaling can
expand the hepatic progenitor cell population (Chapter 2). Further analysis of the tumorigenic
potential of Fgf10-induced HPCs may provide key insight into the regulation of HPCs in
diseases such as hepatoblastoma.
FGF signaling is an established regulator of hepatogenesis. Previous work in our lab has
demonstrated that FGF10/FGFR2b signaling acts upstream of β-catenin activation on the
hepatoblast cell population in vivo [55]. More in depth in vitro analysis of primary hepatoblasts
and tumor initiating cells revealed that FGF signaling resulted in AKT-mediated β-catenin
activation [47]. Subsequent β-catenin interaction with CBP regulates the proliferation and
survival of the hepatic progenitor cell population [47]. In concert with genetic screening of HBs
has revealed that 50-90% of tumors have a genetic mutation related to the Wnt pathway
including CTNNB1, the gene which encodes β-catenin [166]. Non-canonical β-catenin
activation downstream of HGF activation is also found with hepatoblastoma [167]. In vitro
studies of pharmaceutical β-catenin inhibitors shows reduced HB cell viability, suggesting this
may be a good therapeutic target for high risk HB patients [168]. Berg et al found that FGF7 and
FGF10 up-regulate beta-catenin activation during hepatogenesis, suggesting a link between these
two signaling pathways in the progression of HB [55]. Ectopic Fgf10 over-expression in the
adult results in AKT-dependent β-catenin activation and A6 expression in periportal hepatocytes
(Chapter 3). The present study supports the expansion of progenitor cells in the presence of
Fgf10 over-expression during early postnatal development (Chapter 2). During early postnatal
70
development, we characterized a periportal cell population as expressing hepatic progenitor cell
markers (Chapter 2). This cell population which normally abrogates by murine postnatal day 21
is seen to proliferate in response to Fgf10 over-expression. Further analysis of this model and
subsequent activation of β-catenin may provide greater insight into the pathways which regulate
postnatal hepatic progenitor cells.
In postnatal DDC induced cholestatic liver injury, FGF7 signaling has been shown to be a
key regulator of the HPC population [21]. FGF7 is expressed from the mesenchymal cell
population in the periportal region to regulate the atypical ductular reaction and HPC expansion
[21]. Knockout of Fgf7 resulted in impairment of the CK19
+ive
HPC proliferation following
DDC injury [21]. Also, canonical WNT signaling has been shown to regulate the HPC
proliferation induced by DDC liver injury [130].Hyper-activation of β-catenin signaling in
conjunction with DDC liver injury also resulted in increased A6-expressing hepatocytes from the
periportal region towards the central vein [131]. Our own study showed that FGF signaling
modulates AKT-dependent β-catenin activation as well as the A6
+ive
HPCs and distribution of
A6
+ive
hepatocytes (Chapter 3). These data indicate that there may be cross-talk between these
FGF signaling and β-catenin activation to regulate both cholestatic liver injury induced HPC
expansion as well as reprogramming of hepatocytes towards a biliary epithelial cell fate. Further
investigation into the cross-talk between these signaling pathways and how they regulate
hepatocyte reprogramming will be the next step to improve the current field of research on
hepatocyte reprogramming in vivo. Hepatocytes are currently being considered as a source for
cell therapy in liver disease, thus improved understanding of de-differentiation of these cells will
be crucial to clinical applications.
71
This work overall will provide insight into the regulation of “stem-ness” in the liver of
not only the resident hepatic progenitor cell population, but also hepatocytes. Greater
understanding of how hepatic progenitor cells are regulated and their subsequent differentiation
will provide greater insight in to how to regulate these cells and utilize them for liver cell
therapies in the future.
72
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Abstract (if available)
Abstract
Fibroblast Growth Factor (FGF) signaling is an established regulator of endoderm specification to the hepatic fate. We have previously determined that Fibroblast Growth Factors (FGFs) promote the proliferation and survival of embryonic hepatic progenitor cells (HPCs), termed hepatoblasts, and a transformed murine HPC line via AKT-dependent β-catenin activation. Recent studies have shown that postnatal expansion of A6-expressing HPCs during 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC) induced liver injury is partly mediated by β-catenin activation and FGF signaling. Herein, we examine the role of FGF signaling in early postnatal HPCs and acute DDC-induced HPC expansion. Methods: The effects of FGF10 on early postnatal HPCs was studied by inducible transgenic pups over-expressing Fgf10 (P21 Fgf10 Induced) from postnatal day 7 (P7) to P21. For postnatal liver injury, inducible transgenic mice were fed 0.1% DDC chow for 14 days concurrent with either over-activation of FGF signaling by Fgf10 over-expression or inhibition of FGF signaling via dominant-negative expression of soluble FGF Receptor (R)-2IIIb. Results: During early postnatal development, a resident population of cells with a high nuclei-to cytoplasmic ratio increases from postnatal day 0 (P0) through P7 before it disappears histologically. Over-expression of Fgf10 from P7 to P21 retains and expands the periportal cell population which normally declines after P7. Characterization of this cell population revealed they express HPC markers CD49f, and CD133, but lack expression of markers for mesenchymal cells, differentiated hepatocytes and biliary epithelial cells. This heterogeneous cell population expressed epithelial cell marker E-Cadherin, and a subset of the cells was CD45 positive, indicating infiltration of hematopoietic cells as well. During acute postnatal DDC injury, we observed expansion of cells expressing FGFR1 and FGFR2 in the periportal ductular reaction. Quantitative PCR analysis of the non-hepatocyte cell fraction demonstrated significant up-regulation of genes encoding FGFR2IIIb ligands: Fgf7, Fgf10, and Fgf22. Over-expression of Fgf10 increased the number of cells co-positive for HPC marker A6 and hepatocyte marker HEPATOCYTE NUCLEAR FACTOR-4α (HNF4α) while reducing serum total bilirubin. Dominant-negative FGFR pathway inhibition reduced the number of periportal cells exhibiting AKT-dependent activation of β-catenin, and changed the distribution of cells co-expressing A6 and HNF4α by reducing dual positive cells near the central vein of the hepatic lobule while increasing them near the portal vein. Co-treatment of Fgf10 over-expressing DDC-treated mice with AKT inhibitor Wortmannin similarly changed the distribution of A6/HNF4α co-positive cells. Conclusion: FGF signaling regulates the expansion of early postnatal hepatic progenitor cells and DDC-induced A6-expressing hepatocytes partly through AKT-dependent activation of β-catenin.
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The role of fibroblast growth factor signaling on postnatal hepatic progenitor cell expansion
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