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Determination of the source of myofibroblasts and therapeutic strategy against fibrosis

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Determination of the source of myofibroblasts and therapeutic strategy against fibrosis

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Content DETERMINATION OF THE SOURCE OF MYOFIBROBLASTS
AND THERAPEUTIC STRATEGY AGAINST FIBROSIS

by
Ingrid Ahim Lua

Dissertation
Presented to the Faculty of the Graduate School of
University of Southern California
in Partial Fulfillment
of the Requirements
for the Degree of

Doctor of Philosophy
Genetic, Molecular and Cellular Biology

University of Southern California
Aug 2016

Copyright
by
Ingrid Ahim Lua
2016

The Dissertation Committee for Ingrid Ahim Lua Certifies that this is the approved
version of the following dissertation:

DETERMINATION OF THE SOURCE OF MYOFIBROBLASTS
AND THERAPEUTIC STRATEGY AGAINST FIBROSIS

Committee:

Dr. Kinji Asahina, Supervisor
Dr. Hidekazu Tsukamoto
Dr. Cheng-Ming Chuong
iv
Dedication
To my lovely family members, Ahai Lua, Mei-Jin Fang, Evelyn Lua, and my partner,
Yuan-Li Tsai, for their unconditional support along the way.

v
Acknowledgements
This dissertation would not have been possible without all the support of many people.
First of all, I would like to wholeheartedly thank my advisor, Dr. Kinji Asahina, for his
tremendous guidance during my Ph.D. study. He dedicated himself to teaching and had
great patients with students. More importantly, he was approachable and willing to share
his knowledge generously. We also had a great time outside of laboratory. We attend
several conferences and went hiking and sightseeing together. To me, his was not only a
mentor, but also a good friend. He was certainly the best mentor I ever had. I am also
grateful for my committee members throughout the years. I greatly appreciate Dr.
Hidekazu Tsukamoto for his insightful discussions in every joint lab meeting and
research support for my project. At the same time, I would like to thank Dr. Cheng-Ming
Chuong for his support when I joined the NICHD T32 Development, Stem Cells and
Regeneration Program. It was an important fellowship for my Ph.D. study. In addition, I
would also like to thank the members from Southern California Research Center for
ALPD & Cirrhosis and former members including Yuchang Li for making the laboratory
such a friendly environment. I would thank the technical support and core facilities from
USC Norris Bioinformatics Core and Flow Cytometry Core Facility. Moreover, I
appreciate the collaboration with various labs including Dr. Kasper Wang from
Children’s Hospital Los Angeles. Last but not least, I would like to express my deepest
gratitude and love towards my loving family members, my partner and friends in Los
Angeles and Taiwan. They have always been standing by me and gave me strength to
excel through this journey.
vi
Abstract
Organ fibrosis is a worldwide health care problem as it leads to end-stage organ failure in
patients. Currently, there is no medical treatment for organ fibrosis, such as in the liver
and peritoneum. Myofibroblasts have been suggested to participate in fibrogenesis
through synthesizing extracellular matrices and proinflammatory cytokines.
Accumulating evidence suggest that there are multiple cellular sources of myofibroblasts.
However, it remains elusive how different cell types contribute to myofibroblasts in
organ fibrosis. Thus, identification of cell types responsible for fibrogenesis and
understanding molecular mechanisms underlying generation of myofibroblasts are
essential to determine therapeutic targets for suppression of fibrogenesis. Prolonged
injury to the liver often results in liver fibrosis. Similarly, patients who undergo
peritoneal dialysis for kidney failure also cause peritoneal fibrosis. In the present study,
we focused on the liver and peritoneum as visceral and parietal organs in the peritoneal
cavity and identified the cellular sources of myofibroblasts in liver and peritoneal
fibrosis. Moreover, we examined molecular mechanisms underlying fibrogenesis in these
organs. Mesothelial cells (MCs) form a single epithelial cell layer, called mesothelium,
on the surface of the both organs in the peritoneal cavity. We found that MCs give rise to
myofibroblasts in both organs. Although MCs partly contributed to myofibroblasts in
peritoneal fibrosis, protection of the mesothelium ameliorated fibrosis. In liver fibrosis,
hepatic stellate cells (HSCs) reside in the space of Disse and are known to differentiate
into myofibroblasts. In addition, portal fibroblasts (PFs) around the bile duct are believed
to be the major source of myofibroblasts in biliary fibrosis. However, contribution of PFs
vii
to myofibroblasts remains elusive due to insufficient availability of markers and isolation
methods. We established a unique protocol to isolate MCs, HSCs, and PFs from normal
or injured mouse livers and demonstrated that all three cell types convert into
myofibroblasts and transforming growth factor-  (TGF- ) stimulates their differentiation.
Our data indicate that HSCs are the dominant source of myofibroblasts in liver fibrosis
caused by different etiology. Although PFs and MCs partly contributed to myofibroblasts
in liver fibrosis, they uniquely participated in biliary fibrosis and capsular fibrosis,
respectively. Lastly, we identified lysophosphatidic acid (LPA) as a profibrogenic factor
in liver fibrosis. We found that LPA signaling stimulates nuclear localization of TAZ
coactivator and induces differentiation of HSCs and MCs toward myofibroblasts.
Treatment with an LPA inhibitor reduced liver fibrosis in mice. In conclusion, we
demonstrated that MCs act as a source of myofibroblasts in liver and peritoneal fibrosis.
In liver fibrosis, HSCs are the major source of myofibroblasts and both MCs and PFs
uniquely contribute to fibrosis beneath the liver surface and around the bile duct,
respectively. Our data indicate that LPA acts as a profibrogenic factor and its signaling
pathway will be a new therapeutic target for suppression of liver fibrosis.

viii
Table of Contents
Dedication .............................................................................................................. iv
Acknowledgements ..................................................................................................v
Abstract .................................................................................................................. vi
Abbreviations ....................................................................................................... xiv

Chapter 1 Introduction
1.1 Fibrosis and myofibroblasts ......................................................................1
1.2 Peritoneal fibrosis .....................................................................................3
1.3 Liver fibrosis .............................................................................................5

Chapter 2 Conversion of mesothelial cells into myofibroblasts in peritoneal
and liver fibrosis
2.1 Introduction ...............................................................................................8
2.2 Materials .................................................................................................12
2.3 Results .....................................................................................................21
Phenotype of peritoneal MCs...............................................................21
Differentiation of MCs to myofibroblasts in peritoneal fibrosis .........24
Regeneration of peritoneal MCs ..........................................................28
Isolation and culture of peritoneal MCs ...............................................32
Induction of MMT by TGF-  ..............................................................36
Conditional deletion of Tgfbr2 gene in MCs .......................................36
Role of Tgfbr2 gene in regeneration ....................................................41
CG causes liver injury ..........................................................................44
Distince origins of peritoneal and liver MCs .......................................45
2.4 Discussion ...............................................................................................50

ix
Chapter 3 Isolation of hepatic stellate cells, portal fibroblasts, and mesothelial
cells from mouse livers and quantification of their contribution to
myofibroblasts in liver fibrosis
3.1 Introduction .............................................................................................58
3.2 Materials .................................................................................................60
3.3 Results .....................................................................................................63
Broad expression of GFP in the Col1a1
GFP
mouse liver......................63
Separating HSCs with FACS ...............................................................66
The presence of PFs and MCs in the VitA-GFP+ population .............70
Enrichment of PFs by FACS ...............................................................74
Myofibroblastic conversion of HSCs, PFs, and MCs ..........................76
Expression of HSC, PF and MC markers in BDL-induced biliary
fibrosis.........................................................................................79
Expression of HSC, PF and MC markers in DDC diet-induced
biliary fibrosis .............................................................................82
Expression of HSC, PF and MC markers in CCl
4
-induced liver
fibrosis .......................................................................................85
Isolation of HSCs, PFs, and MCs from fibrotic livers .........................88
3.4 Discussion ...............................................................................................91

Chapter 4 No transdifferentiation of mesenchymal cells to epithelial cells in
liver fibrosis
4.1 Introduction .............................................................................................97
4.2 Materials .................................................................................................99
4.3 Results ...................................................................................................101
MesP1+ mesoderm gives rise to HSCs, PFs, SMCs, and MCs in
the adult liver ............................................................................101
Rare contribution of the MesP1+ mesoderm to Kupffer cells ...........105
Mesodermal origin of HSCs ..............................................................107
Activation of mesodermal HSCs in culture .......................................110
No contribution of HSCs and PFs to hepatocytes and
cholangiocytes in liver fibrosis .................................................114
x
No contribution of HSCs and PFs to oval cells .................................117
No contribution of mesodermal mesenchymal cells to epithelial
cells in the regenerating liver ....................................................120
4.4 Discussion .............................................................................................123

Chapter 5 Lipid mediators in liver fibrosis
5.1 Introduction ...........................................................................................128
5.2 Materials ...............................................................................................134
5.3 Results ...................................................................................................137
Cell-cell contacts counteract TGF- -induced MMT .........................137
TAZ regulates MMT ..........................................................................141
F-actin formation enhances MMT .....................................................144
LPA enhances TGF- 1-induced MMT ..............................................146
LPA induces MMT ............................................................................148
LPA activates HSCs through TAZ ....................................................150
Inhibition of LPA reduces liver fibrosis ............................................152
5.4 Discussion .............................................................................................154

Chapter 6 Conclusions & Perspectives
6.1 Conclusions ...........................................................................................157
6.2 Future directions ...................................................................................150

Bibliography .......................................................................................................160
xi
List of Tables
Table 1: List of antibodies for immunostaining ..............................................15
Table 2: List of primers for QPCR ..................................................................18
Table 3: Summary of microarray analysis ......................................................69

xii
List of Figures
Figure 1: Characterization of peritoneal MCs on the mouse body walls .........22
Figure 2: Peritoneal MCs give rise to myofibroblasts in peritoneal fibrosis ....26
Figure 3: Lineage tracing of MCs during recovery from fibrosis. ...................30
Figure 4: Induction of peritoneal MMT by TGF-  in vitro ..............................34
Figure 5: Essential role of TGF-  signaling in peritoneal MMT in fibrosis ....38
Figure 6: Deletion of Tgfbr2 gene in MCs .......................................................42
Figure 7: Lineage tracing of liver and peritoneal MCs in CG-induced injury .47
Figure 8: A proposed model of CG-induced fibrosis .......................................56
Figure 9: Expression of GFP in HSCs, PFs, SMCs, SLCs, CFs, and MCs in
Col1a1
GFP
mouse livers.....................................................................64
Figure 10: Separation of VitA+ HSCs and VitA-GFP+ population from Col1a1
GFP

livers ..................................................................................................67
Figure 11: Expression of MC markers in mouse and human livers ...................72
Figure 12: Separation of HSCs, PFs, and MCs from the Col1a1
GFP
livers by FACS
...........................................................................................................75
Figure 13: Differential responses of HSCs, PFs, and MCs against TGF- 1 and
PDGF-BB ..........................................................................................77
Figure 14: Expression of markers in biliary fibrosis induced by BDL ..............80
Figure 15: Expression of markers in biliary fibrosis induced by the DDC diet .83
Figure 16: Expression of markers in CCl
4
-induced liver fibrosis ......................86
Figure 17: Isolation of HSCs, PFs, and MCs from Col1a1
GFP
fibrotic livers ....90
Figure 18: Summary of the contribution of HSCs, PFs, and MCs in liver fibrosis
...........................................................................................................96
xiii
Figure 19: Contribution of MesP1+ mesoderm to HSCs, PFs, SMCs, and MCs, but
not hepatocytes and cholangiocytes, in the adult liver ...................103
Figure 20: Few Kupffer cells express GFP in MesP1
Cre/+
; R26TG
fl/fl
mouse liver
.........................................................................................................106
Figure 21: In vitro activation of mesodermal HSCs .........................................108
Figure 22: Similar phenotypes of GFP+ and GFP- HSCs in the MesP1
Cre/+
;
R26TG
fl/fl
liver ................................................................................112
Figure 23: No contribution of mesodermal mesenchymal cells to hepatocytes and
cholangiocytes in liver fibrosis .......................................................115
Figure 24: No contribution of mesodermal mesenchymal cells to oval cells,
hepatocytes and cholangiocytes in injured liver .............................118
Figure 25: No contribution of mesodermal mesenchymal cells to hepatocytes and
cholangiocytes in the regenerating liver .........................................121
Figure 26: A propose model of MesP1+ mesoderm in the liver ......................127
Figure 27: Chemical structure of major species of LPA ..................................129
Figure 28: LPA synthesis pathways. ................................................................132
Figure 29: TGF-   does not fully induce MMT at high cell density ...............139
Figure 30: TAZ regulates MMT .......................................................................143
Figure 31: F actin formation, but not MST1/2, is required for MMT ..............145
Figure 32: LPA enhances TGF-  -induced MMT via TAZ ............................147
Figure 33: LPAR expression pattern in liver cells ...........................................149
Figure 34: LPA induces HSC activation ..........................................................151
Figure 35: Inhibition of LPA signaling in liver fibrosis ...................................153
Figure 36: A proposed model of LPA-TAZ signaling .....................................156

xiv
Abbreviations
-smooth muscle actin (ACTA2); Bile duct ligation (BDL); Bromodeoxyuridine (BrdU);
Capsular fibroblasts (CFs); Carbon tetrachloride (CCl
4
); Chlorhexidine gluconate (CG);
Collagen1a1 promoter-green fluorescent protein (Col1a1
GFP
); Connective tissue growth
factor (CTGF); Cytokeratin 8 (KRT8); Cytokeratin 19 (KRT19; CK19); 3,5-
diethoxycarbonyl-1,4-dihydrocollidine (DDC); Desmin (DES); E-cadherin (CDH1);
Elastin (ELN); Ectonucleoside triphosphate diphosphohydrolase-2 (ENTPD2); Epithelial-
mesenchymal transition (EMT); Fluorescence-activated cell sorting (FACS);
Glycoprotein M6A (GPM6A); Hepatic stellate cells (HSCs); Lysophosphatidic acid
(LPA); Magnetic-activated cell sorting (MACS); Mesothelin (MSLN); Mesothelial cells
(MCs); Mesothelial-mesenchymal transition (MMT); Nonparenchymal cells (NPCs);
Platelet-derived growth factor (PDGF); Podoplanin (PDPN); Portal fibroblasts (PFs);
Quantitative polymerase chain reaction (QPCR); Reelin (Reln); Rosa26mTmG
flox

(R26TG
fl
); Second-layer cells (SLCs); Sinusoid endothelial cells (SECs); Smooth muscle
cells (SMCs); Transforming growth factor-  (TGF- ); TGF-  type II receptor
(TGFBR2); Type I collagen (COL I); Type IV collagen (COL IV); Uroplakin 1b
(Upk1b); Vimentin (VIM); Vitamin A (VitA); Wilms tumor 1 (WT1)

1
Chapter 1
Introduction
1.1 Fibrosis and myofibroblasts
Wound healing is a complicated biological process in repairing injured organisms
(Reinke and Sorg, 2012). In response to injury and stress, the synthetic machinery of
extracellular matrix proteins responds to injury and helps heal the wound in a tissue
homeostasis manner. Physiological wound healing is accomplished by the precise balance
between the rate of synthesis of extracellular matrix proteins by myofibroblasts and their
degradation by proteolytic enzymes. However, prolonged inflammation and sustained
activation of myofibroblasts lead to excessive collagen deposition in the wound area. This
pathological wound healing often results in fibrosis. Fibrosis is a scaring process
characterized by excessive accumulation of extracellular matrix proteins along with
massive inflammation (Ghosh et al., 2013). Multiple organs suffer from fibrosis, such as
liver fibrosis, peritoneal fibrosis, idiopathic pulmonary fibrosis, systemic sclerosis,
myocardial infarction, and glomerulosclerosis (Gabrielli et al., 2009; D'Agati et al., 2011;
Korte et al., 2011; Hinz, 2012; Jaffe and Apple, 2012; Schuppan and Kim, 2013). Fibrosis
causes loss of tissue homeostasis and eventually leads to end-stage organ failure.
Currently, there is no effective therapy for treatment of fibrosis, and thus novel
therapeutic inventions are urgently needed. In experimental models, lines of evidence
suggest that control and removal of the causative etiology can delay fibrosis progression
and lead to fibrosis regression (Issa et al., 2004; Zareie et al., 2005; Nishioka et al., 2008),
2
yet such treatment options have not been succeeding in clinical (Hauff et al., 2015;
Koyama and Brenner, 2015). Treatment strategies should not only focus on eliminating
the causative etiology, but also target the downstream effector-myofibroblasts.
Myofibroblasts are the primary source for producing extracellular matrix proteins
and were first identified in 1971 (Gabbiani et al., 1971). In general, there are no
myofibroblasts in normal health tissues. Upon injury, local or recruited fibroblasts
become activated and differentiate into myofibroblast (Wynn, 2008). One well known
feature of the myofibroblast is the expression of-smooth muscle actin ( C  ) that
confers their contractile activity (Klingberg et al., 2013). Although myofibroblasts have
been extensively studied, accumulating evidence suggest that there are multiple origins of
myofibroblasts in organ fibrosis. Therefore, identification the origin of myofibroblasts
and molecular mechanism underlying myofibroblastic conversion are a prerequisite in
developing anti-fibrosis therapy.

3
1.2 Peritoneal fibrosis
Approximately 400,000 patients are being treated with hemodialysis or peritoneal
dialysis for treatment of end-stage renal disease in the United States (Weinhandl et al.,
2015). Compared with hemodialysis, peritoneal dialysis is a cost effective therapy and
shows a lesser mortality rate for the initial 1-2 years. In peritoneal dialysis, the
mesothelium on the body wall is used as a semipermeable barrier in patients. However,
long-term exposure to dialysis solution causes injury to peritoneal mesothelial cells
(MCs) and often leads to peritoneal fibrosis and encapsulating peritoneal sclerosis
characterized by excessive fibrotic thickening of the peritoneum (Korte et al., 2011). In
the body wall peritoneum, a single layer of MCs is separated from underlying fibroblasts
by a basal lamina. Injury to the body wall causes the disappearance of the MC layer from
the body wall surface, massive development of connective tissue consisting of
myofibroblasts and blood vessels, and deterioration of the barrier function (Aroeira et al.,
2007; Devuyst et al., 2010; Korte et al., 2011). In patients who underwent peritoneal
dialysis, the dialysis effluent from the peritoneal cavity is known to contain floating MCs
and these MCs are shown to undergo epithelial-mesenchymal transition (EMT) and
differentiate into fibroblastic cells in culture (Yanez-Mo et al., 2003). Since this finding,
different laboratories have studied EMT of MCs and it has generally accepted that MCs
are the major source of myofibroblasts in peritoneal fibrosis. In addition to MCs,
fibroblasts beneath MCs are also suggested to be the source of myofibroblasts in
peritoneal fibrosis (Beavis et al., 1997; Sakai et al., 2013; Chen et al., 2014).
4
Nevertheless, the origin of myofibroblasts in peritoneal fibrosis remains to be determined
by rigorous cell lineage tracing.
Different signaling pathways have been identified to be inducers of EMT of MCs.
For example, the renin-angiotensin-aldosterone system has been suggested to contribute
to peritoneal fibrosis (Ersoy et al., 2007), yet clinical trials targeting this system failed to
improve fibrosis in patients (Vazquez-Rangel et al., 2014). Thus, more critical signaling
pathways that involved in peritoneal fibrosis need to be identified for suppression of
peritoneal fibrosis. In addition to the development of fibrosis, several studies have
reported that peritoneal fibrosis is reversible (Nishioka et al., 2008). However, cellular
sources of regenerated MCs are not known due to the lack of lineage tracing methods. In
the present study, we conducted rigorous cell lineage tracing and demonstrated that
peritoneal MCs partially contribute to myofibroblasts in peritoneal fibrosis through
transforming growth factor-  (TGF- ) signaling pathway. In addition, survived
peritoneal MCs are the source of regenerated mesothelium from peritoneal fibrosis. For
details results, we will discuss more in Chapter 2.

5
1.3 Liver fibrosis
Chronic liver injury caused by excessive alcohol consumption, drugs, obese/diabetes, and
hepatitis virus infection leads to liver fibrosis and cirrhosis (Hernandez-Gea and
Friedman, 2011; Schuppan and Kim, 2013; Wree et al., 2013; Pellicoro et al., 2014;
Louvet and Mathurin, 2015). Cirrhosis is the 12th leading cause of death in the United
States, accounting for nearly 32,000 deaths each year (Murphy et al., 2013). Currently, no
drugs cure patients with cirrhosis except liver transplantation. Although cirrhosis is
irreversible, recent clinical cases and animal studies indicate liver fibrosis is reversible
(Iredale et al., 1998; Ellis and Mann, 2012; Kisseleva et al., 2012; Troeger et al., 2012).
Therefore, suppression of liver fibrosis is a key target for preventing cirrhosis progression.
Hepatic stellate cells (HSCs) reside in the space of Disse in the liver and store vitamin A
(VitA) lipids as retinylester in their cytoplasm (Friedman, 2010). HSCs extend dendritic
processes along the sinusoidal wall and express desmin (DES) and type I collagen (COL I)
(Yin et al., 2013). Upon liver injury, various insults including reactive oxygen species
generated from damaged hepatocytes and TGF-  from Kupffer cells induce the activation
of HSCs that express ACTA2. Activated HSCs acquire myofibroblasts phenotype and
synthesize extracellular matrix proteins and proinflammatory cytokines. Although HSCs
are suggested to be the major source of myofibroblasts (Mederacke et al., 2013),
myofibroblasts seem to be heterogeneous and they might derive from different sources
during liver fibrosis (Knittel et al., 1999; Cassiman et al., 2002; Lemoinne et al., 2013;
Iwaisako et al., 2014). Bone marrow-derived cells have been shown to differentiate into
HSCs or myofibroblasts, yet several studies reported that their contribution is negligible
6
in liver fibrosis (Baba et al., 2004; Forbes et al., 2004; Kisseleva et al., 2006;
Higashiyama et al., 2009). EMT of hepatocytes or cholangiocytes into myofibroblasts
was also proposed in liver fibrosis, but cell-lineage tracing studies have refuted this
possibility in mice (Zeisberg et al., 2007; Nitta et al., 2008; Chu et al., 2011; Xie and
Diehl, 2013).
Asahina’s laboratory previously reported that MCs covering the liver surface give
rise to HSCs and portal fibroblasts (PFs) in liver development. In liver fibrosis, MCs
migrate inward and give rise to HSCs or myofibroblasts near the liver surface depending
on the etiology (Li et al., 2013). HSCs, PFs, and MCs are known to be the source of
myofibroblasts; however, it remains elusive the relative contribution of these cells in liver
fibrosis caused by different etiology due to insufficient availability of markers and
isolation methods. Furthermore, several studies reported that HSCs differentiate into
epithelial cell lineage, including progenitor cell types known as oval cells which act as
stem cells in the liver regeneration (Yang et al., 2008; Michelotti et al., 2013). In Chapter
3, we established a new isolation protocol to separate MCs, HSCs, and PFs by
fluorescence-activated cell sorting (FACS) from normal or injured mouse livers and
quantified the relative contribution of these cells in liver fibrosis caused by different
etiology. In Chapter 4, we traced cell lineages of liver mesenchymal cells including HSCs,
PFs, and MCs and refuted the notion that HSCs act as liver progenitor cells.
TGF-is a profibrogenic factor in organ fibrosis and stimulates differentiation of
fibroblasts to myofibroblasts (Hinz et al., 2012). TGF-  binds to a heterodimer of two
TGF-  type II (TGFBR2) and two type I receptors, activates phosphorylation of
7
SMAD2/3, and regulates target gene expression (Lamouille et al., 2014). We found that
TGF-  induces myofibroblastic conversion of HSCs, PFs, and MCs equally. Although
many tools are available to suppress TGF-  signaling, it is difficult to selectively
suppress fibrosis due to its wide range of biological functions including inhibition of cell
proliferation, immunity, and inflammation (Akhurst and Hata, 2012). In addition,
outcome of clinical trials using TGF-  neutralizing antibody has no effect or even
adverse effect on glaucoma or systemic sclerosis patients (van Meeteren and ten Dijke,
2012). Thus, it is essential to identify other signaling pathways that may modulate TGF- 
signaling or regulate myofibrogenesis independently from TGF-  signaling, for targeting
liver fibrosis. In Chapter 5, we identified that lysophosphatidic acid (LPA) acts as a
profibrogenic factor and its signaling pathway is a novel therapeutic target for repression
of liver fibrosis.

8
Chapter 2
Conversion of mesothelial cells into myofibroblasts in peritoneal and
liver fibrosis
2.1 Introduction
MCs form single epithelial sheets that cover the surface of visceral organs, including the
heart, lung, digestive tract, omentum, and ovaries. They also cover the parietal wall of the
pleura, pericardium, and peritoneum (Mutsaers, 2004). Their major function is to
synthesize lubricating fluids to create a slippery surface that facilitates movement
between organs without friction. For example, MCs are the major source of
proteoglycans in the peritoneal fluid (Yung et al., 1995). MCs express epithelial cell
markers and form epithelial sheets with tight junctions, adherence junctions, and gap
junctions on the surface of organs (Simionescu and Simionescu, 1977). Interestingly,
MCs also express mesenchymal cell markers. MCs in the normal adult mouse liver
express Wilms tumor 1 (WT1), cytokeratin 8 (KRT8), cytokeratin 19 (KRT19), and
vimentin (VIM) but not E-cadherin (CDH1) (Li et al., 2013), indicating that MCs have an
intermediate phenotype of epithelial and mesenchymal cells. In addition to these markers,
MCs express CD200, glycoprotein M6A (GPM6A), mesothelin (MSLN), and podoplanin
(PDPN) (Li et al., 2013).
In chick embryos, MCs originate from the lateral plate mesoderm (Thomason et
al., 2012). After gastrulation, the lateral plate mesoderm delaminates and coelomic cavity
develops in the mesoderm. The dorsal mesoderm associated with the ectoderm gives rise
to somatic mesoderm and forms the parietal mesothelium of the body wall. In contrast,
9
the mesoderm associated with the endoderm becomes splanchnic mesoderm and
contributes to the visceral mesothelium of the respiratory and digestive tracts. In mouse
embryogenesis, MCs in the liver were shown to originate from MesP1+ mesoderm
(Asahina et al., 2009). MESP1 is a transcription factor that is transiently expressed in
nascent mesoderm including the lateral plate mesoderm during gastrulation (Saga et al.,
1999). Although Mesp1+ mesoderm contributes to liver MCs in embryonic mouse livers
(Asahina et al., 2009; Lua et al., 2014), it remains elusive whether both visceral and
parietal MCs are derived from the same Mesp1+ mesoderm in mice.
The multi-differentiation potential of MCs was reported in mouse development.
Conditional cell lineage tracing using Wt1
CreERT2
mice revealed that Wt1+ MCs migrate
inward from the liver surface and give rise to HSCs, PFs and smooth muscle cells (SMCs)
during mouse liver development (Asahina et al., 2011). Lineage tracing of Msln+ MCs
also indicated their contribution to fibroblast and SMCs of the internal organs in mouse
embryos (Rinkevich et al., 2012). During lung development, MCs were shown to migrate
inward and give rise to SMCs and mesenchymal cells (Que et al., 2008; Cano et al., 2013;
Dixit et al., 2013). Similar contributions have been reported in developing heart and gut
(Wilm et al., 2005; Zhou et al., 2008; Carmona et al., 2013; Winters et al., 2014). In
addition, Wt1+ MCs were shown to differentiate into visceral fat in mice (Chau et al.,
2014). Thus, MCs are the source of mesenchymal cells in organ development.
Recent studies indicate that this process is recapitulated in organ fibrosis such as
in the liver and lung. In liver fibrosis, MCs convert into HSCs or myofibroblast
depending on the etiology (Li et al., 2013; Li et al., 2016). In idiopathic pulmonary
10
fibrosis, MCs also gave rise to myofibroblasts (Karki et al., 2014). Because MCs have an
intermediate phenotype of epithelial and mesenchymal cells in normal condition, we
defined the change of MCs into myofibroblasts as mesothelial-mesenchymal transition
(MMT) instead of epithelial-mesenchymal transition (EMT) (Li et al., 2013). TGF- 
plays a critical role in generation of myofibroblast in organ fibrosis (Hinz et al., 2012).
We previously reported that TGF-  induces liver MCs into ACTA2+ myofibroblasts in
vitro (Li et al., 2013). Inhibition of TGF-  signaling by treatment with a soluble form of
TGF-  type II receptor resulted in suppression of MMT in liver fibrosis, suggesting that
TGF-  is involved in MMT. However, it remains unclear whether direct TGF-  signaling
is involved in MMT in both visceral and parietal MCs in organ fibrosis.
The mesothelium on the body wall is used as a semipermeable barrier in patients
who undergo peritoneal dialysis for end-stage renal disease (Aroeira et al., 2007; Devuyst
et al., 2010; Korte et al., 2011). Prolonged exposure of a dialysis solution to the
peritoneal cavity causes injury to peritoneal MCs and induces their conversion into
myofibroblasts which lead to the development of peritoneal fibrosis. Peritoneal fibrosis
can be reproduced in mice or rats by injection of dialysis solution or chemicals. For
example, administration of chlorhexidine gluconate (CG) or sodium hypochlorite to the
peritoneal cavity causes injury to MCs on the body wall and induces peritoneal fibrosis
(Subeq et al., 2011; Chen et al., 2014). Effluent of the dialysis solution from the
peritoneal cavity contains floating MCs (Carozzi et al., 1997) and these floating MCs
were shown to undergo EMT through TGF-  signaling and differentiate into
11
myofibroblasts in culture (Yanez-Mo et al., 2003). After this finding, conversion of MCs
to mesenchymal cells has been extensively studied and several signaling pathways, such
as TGF-   IL-1  and angiotensin II, have been demonstrated to induce this conversion
(Patel et al., 2010; Strippoli et al., 2010; Xie et al., 2010). Although MCs are believed to
be the major source of myofibroblasts in peritoneal fibrosis, several studies suggest that
fibroblasts beneath MCs give rise to myofibroblasts in peritoneal fibrosis as well (Beavis
et al., 1997; Sakai et al., 2013; Chen et al., 2014). Sakai et al. reported that peritoneal
MCs secrete connective tissue growth factor (CTGF) that induces fibroblast proliferation
in peritoneal fibrosis (Sakai et al., 2013). The origin of myofibroblasts in peritoneal
fibrosis remains to be clarified by rigorous cell lineage tracing.
MCs exhibit different phenotypes in different organs. Visceral MCs express
autotaxin, and they have greater migration ability compared to parietal MCs (Shelton et
al., 2013). We are interested in whether visceral and parietal MCs behave similarly during
development, tissue injury, and regeneration. The aim of our study is to characterize and
determine the fate of MCs in fibrosis by comparing liver MCs and peritoneal MCs as
representatives of visceral and parietal organs, respectively. Injections of CG into the
peritoneal cavity also induce fibrosis beneath the liver surface. In the present study, we
defined this fibrosis pattern as capsular fibrosis in the liver. Using Wt1
CreERT2
and
Rosa26mTmG
flox
(R26TG
fl
) mice, we traced peritoneal MCs and liver MCs in the CG
model and demonstrated for the first time that Wt1+ MCs partly contribute to ACTA2+
myofibroblasts in both peritoneal and liver fibrosis.
12
2.2 Materials
Mice. Mesp1
Cre
and Wt1
CreERT2
knock-in mice were obtained from Drs. Henry Sucov and
William Pu (Saga et al., 1999; Zhou et al., 2008). Tgfbr2
fl
, R26TG
fl
, and Uroplakin 1b
(Upk1b)
RFP
knock-in mice were purchased from the Jackson Laboratory (Leveen et al.,
2002; Muzumdar et al., 2007). Collagen1a1 promoter-green fluorescent protein
(Col1a1
GFP
) transgenic mice were obtained from Dr. David Brenner (Yata et al., 2003).
Tamoxifen (Sigma-Aldrich) dissolved in ethanol was emulsified in sesame oil at 12.5
mg/mL and injected intraperitoneally into Wt1
CreERT2
mice (10 to 12 weeks old) at 100
mg/g body weight twice during a 3-day interval. Two weeks after the last injection, the
mice were injected with 0.1% CG solution (Sigma-Aldrich) in 15% ethanol/phosphate
buffered saline at 1.5 mL/100 g body weight to the right part of the peritoneum every
other day (Kushiyama et al., 2011). After 10 injections of CG, the mice developed
peritoneal fibrosis. We collected the left part of the peritoneum for histology analyses to
avoid the area damaged by intraperitoneal injections. For the recovery model, the mice
that were treated 10 times with CG were maintained for 1 month without treatment. For
bromodeoxyuridine (BrdU) incorporation assay, mice were injected with BrdU labeling
reagent (Life Technologies; 10 mL/g body weight; n=3) by intraperitoneal injection 4
hours before sacrifice. For inhibition of TGF-  signaling, mice were treated with STR
(TGF-bR2 Fc chimera; R&D Systems) or mouse IgG2a isotype control (0.1 mg/kg body
weight; n=3) with five intraperitoneal injections of CG every other day. Mice were used
in accordance with protocols approved by the Institutional Animal Care and Use
Committee of the University of Southern California.
13
Immunohistochemistry, Histology, and Quantification. The peritoneal body wall or
liver tissues were fixed with 4% paraformaldehyde or 70% ethanol, incubated with 30%
sucrose in PBS overnight, and embedded in OCT compound (Sakura Finetech). Using a
cryostat (CM1900; Leica), 7- m cryosections were cut. After blocking with 5% goat or
donkey serum, the sections were incubated with primary antibodies for 1 hour. The
primary and secondary antibodies for immunohistochemistry are listed in Table 1.
TOMATO fluorescence was bleached with 3% H
2
O
2
in methanol for 10 min. The
sections were counterstained with DAPI (Life Technologies). Fluorescence signals were
captured with Nikon 90i microscope and DS-Qi1 digital camera (Nikon). For histology,
sections were stained with hematoxylin and eosin (H&E) or Sirius red.
To quantify the labeling efficiency of tamoxifen to MCs, we injected tamoxifen
twice into Wt1
CreERT2/+
; R26TG
fl/fl
mice and sacrificed the mice 2 weeks after the second
injection. After immunostaining the body wall and liver with anti-GFP antibodies (3
sections for each mouse; n=3), digital images were randomly captured using a 20X
objective (60 images for each mouse, 180 images in total), and the length of the GFP+
MCs was measured with imaging software (NIS-Element, Nikon). To quantify MC-
derived ACTA2
+
myofibroblasts in the CG model, sections were stained with antibodies
against ACTA2 and GFP (3 sections for each mouse; n=3). After counterstaining with
DAPI, images were captured from the connective area, and 18,825 myofibroblasts were
counted in the CG model. Similarly, GFP+ MCs were counted in the CG reversal model
(3,237 MCs total; n=3). In the Tgfbr2
fl/fl
model, sections were stained with antibodies
against GPM6A and the length of GPM6A+ MCs was measured (30 images for each
14
mouse; n=3). To quantify the area of connective tissue in the body wall, at least 3
sections from each mouse (3 mice for each group) were stained with antibodies against
ACTA2. After counterstaining with DAPI, at least 20 digital images were randomly
captured from a section using a 20X objective. Each image contained musculature length
in the full-screen mode, and areas of connective tissue were measured by circling the
ACTA2+ area between the surface and musculature of the body wall. Similarly, the area
of connective tissue, the number of GFP+ MCs, or BrdU+ cells were quantified in
Tgfbr2
fl/fl
CG reversal model (n=3).

15
Table 1. List of antibodies for immunostaining

Antibodies Source (catalog no.) Dilution Notes
Goat anti-GFP Rockland (600-101-215) 600
Goat anti-human HNF4 Santa Cruz (sc-6556) 200
Goat anti-mouse RELN R&D Systems (AF3820) 200 ProK*
Hamster anti-mouse PDPN eBioscience (14-5381) 100
Mouse anti-ACTA2-Cy3 Sigma (C6198) 200
Rabbit anti-human ACTA2 Proteintech (14395-1-AP) 200 Paraffin†
Rabbit anti-COLI Millipore (AB755P) 200
Rabbit anti-COL IV Millipore (AB756P) 200
Rabbit anti-Cytokeratin Dako (Z0622) 100
Rabbit anti-human DES Thermo (RB-9014) 300
Rabbit anti-rat Elastin Cedarlane (CL55041AP) 200
Rabbit anti-human MSLN Sigma (HPA017172) 200 Paraffin†
Rabbit anti-rat ELN Cedarlane (CL55041AP) 200
Rabbit anti-mouse ENTPD2 Jean Sévigny 200
Rabbit anti-GFP LifeTechnologies (A11122) 800
Rabbit anti-RFP Rockland (600-401-379) 300
Rabbit anti-P-SMAD3 Abcam (ab52903) 100 Triton-X‡
Rabbit anti-P-SMAD2/3 Santa Cruz (sc-11769-R) 50
Rabbit anti-TAZ Sigma (T4077) 600
Rabbit anti-Vimentin Epitomics (2707-1) 200
Rat anti-A6 Valentina M. Factor, NCI 100 Acetone¶
Rat anti-BrdU Abcam (ab6326) 200
Rat anti-GFP Nacalai Tesque (04404-84) 1,000
Rat anti-mouse CD31 BD Pharmingen (550274) 100
Rat anti-mouse 45 eBioscience (13-0451) 200
Rat anti-mouse 90.2 BD Pharmingen (553011) 200
Rat anti-mouse 133 eBioscience (14-1331) 100
Rat anti-mouse CD200 eBioscience (14-5200) 100
Rat anti-Cytokeratin 8 DSHB (Troma-I) 50 Ethanol§
Rat anti-Cytokeratin 19 DSHB (Troma-IIII) 50 Ethanol§
Rat anti-mouse E-cadherin Zymed (13-1900) 100 Ethanol§
Rat anti-mouse EPCAM DSHB (G8.8) 1
Rat anti-mouse F4/80 eBioscience (13-4801) 1000
Rat anti-mouse GPM6A MBL (D055-3) 500
Rat anti-mouse LYVE1 eBioscience (14-0443) 200
Rat anti-mouse KRT19 DSHB (TROMA-III) 50 Ethanol§
Rat anti-mouse MSLN MBL (D233-3) 200 Acetone¶
Rat anti-mouse THY1 BD Pharmingen (553011) 600

16
* Pretreated cryosections with 20 g/ml Proteinase K for 3 minutes before blocking.
† Pretreated paraffin sections with 10 mM citrate buffer pH 6.0 at 95
o
C for 20 minutes
for antigen retrieval.
‡ Pretreated cells with 0.5% Triton-X in PBS for 10 minutes.
§ Fixed cryosections with ethanol at -20
o
C for 10 minutes.
¶ Fixed cryosections with acetone at -20
o
C for 10 minutes.

17
Quantitative Polymerase Chain Reaction (QPCR). Total RNA was extracted with
RNAqueous Micro (Life Technologies) and cDNA was synthesized using a SuperScript
III kit as previously reported (Asahina et al., 2009). QPCR was performed with SYBR
Green in a ViiA 7 Real-Time PCR System (Life Technologies). The samples were run in
triplicate. The relative mRNA levels per samples were calculated by subtracting the
detection limit (40 Ct) from the cycle threshold value (Ct) of each gene in the same
sample to obtain the Ct value. Taking the log
2
of - Ct resulted in the relative expression
value of each gene for each sample expressed in arbitrary units. Each value was
normalized to Gapdh. The primer sequences are listed in Table 2.

18
Table 2. List of primers for QPCR.
Genes Forward primer (5’-3’) Reverse primer (5'-3’)
Acta2 Ctgagcgtggctattccttc cttctgcatcctgtcagcaa
Alb Tgctgctgattttgttgagg agagttggggttgacacctg
Atx Gtcagaaaggaatggggtca gtcggtgaggaaggatgaaa
Ccnd1 Ggcacctggattgttctgtt cagcttgctagggaacttgg
Cd31 Gaatgacacccaagcgttt ggcttccacactaggctcag
Cd68 Ccaattcagggtggaagaaa ttgcatttccacagcagaag
Cd200 Aacgtcaccgaaatcaggag ggcactgcattgctctacaa
Col1a1 Caccctcaagagcctgagtc gttcgggctgatgtaccagt
Ctgf Atcccaccaaagtgagaacg acagctggactcagcctcat
Des Caggacctgctcaatgtgaa gtagcctcgctgacaacctc
Eln Ctgctaaggctgcccagtat ccacctggataaatgggaga
Entpd2 Gcgctgtagccatgttcata aagagcagcaggagagcaac
Epcam Cggctcagagagactgtgtc gatccagtaggtcctcacgc
Gapdh Cgtcccgtagacaaaatggt gaatttgccgtgagtggagt
Gfap Cacgaacgagtccctagagc ccttctgacacggatttggt
Gfp Cctacggcgtgcagtgcttcagc cggcgagctgcacgctgccgtcctc
Gpm6a Caaggactgctggagacaca acgcagatccaagcagagat
Hgf Ttcccagctggtctatggtc tggtgctgactgcatttctc
Krt8 Atcgagarcaccacctaccg ctcattccgtagctgaagcc
Krt19 Ctcggattgaggagctgaac tcacgctctggatctgtg
Lpar1 Tcttctgggccattttcaac tgcctgaaggtggcgctcat
Lpar2 Accacactcagcctagtcaagac ctgagtaacgggcagacttg
Lpar3 Acaccagtggctccatcag gttcatgacggagttgagcag
Lpar4 Aggcatgagcacattctctc caacctgggtctgagacttg
Lpar5 Aggaagagcaaccgatcatcacag accaccatatgcaaacgatgtg
Lpar6 Tgtttccaactgctgctttg gagcagtcccagtggcttag
Lrat Ctgggagtcatttgcaaggt cagattgcaggaagggtcat
Msln Cttggtcgcctgctatcttc acggacagggcttttatcct
Pdgfra Acagagactgagcgctgaca ctcgatggtctcgtcctctc
Pdgfrb Atcatccccttacctgaccc ctctgcttcagccagaggtc
Pdpn Gtgaccccaggtacaggaga atggctaacaagacgccaac
Reln Gaaaccgagaagcaaagctg caggtgatgccattgttgac
Taz Gcgctcgtctacgtcttctc cctgatgagtctgtgctgga
Tgfb1 Ttgcttcagctccacagaga tggttgtagagggcaaggac
Tgfb2 Ccgctgcatatcgtcctgt agtggatggatggtcctattaca
Tgfbr1 Tcccaactacaggacctttttca gcagtggtaaacctgatccaga
Tgfbr2 Ccgctgcatatcgtcctgt agtggatggatggtcctattaca
Thy1 Gggcgactacttttgtgagc gagggctcctgtttctcctt
Timp1 Cagtaaggcctgtagctgtgc ctcgttgatttctggggaac
Yap Aggagagactgcggttgaaa cccaggagaagacactgcat
Upk1b Agttgcctggtttggatttg tgcagcatcttgaaagccta
19
Isolation and culture MCs. MCs were isolated from the body wall from five mice with
the use of magnetic-activated cell sorting (MACS) as previously described (Li et al.,
2013). In brief, the body wall was digested with 1mg/mL pronase for 20 minutes. After
washing, the cells were incubated with anti-GPM6A antibodies (MBL) and GPM6A+
MCs were separated with anti-rat microbeads and autoMACS (Miltenyi Biotech). The
yield of MCs was approximately 1 x 10
5
MCs from five mice. MCs (2 x 10
4
cells) were
plated on a collagen-coated 24-well plate in Dulbecco’s modified Eagle’s medium with
low glucose that contained 5% fetal bovine serum. The MCs were treated with 10 ng/mL
TGF- 1 (Sigma-Aldrich) from day 4 to 8. MCs were also isolated from five R26TG
fl/fl
;
Tgfbr2
fl/fl
mice, treated with an Adenovirus vector carrying LacZ or Cre at multiplicity of
infection 50 (Kerafast) from day 4, and treated with TGF- 1 from day 6 for 3 or 12 hours.
Cultured MCs were immunostained with antibodies against GFP, GPM6A, ACTA2- Cy3,
and phosphorylated-SMAD3. The primary antibodies were detected with secondary
antibodies conjugated with Alexa Fluor dyes (Life Technologies). Nuclei were
counterstained with DAPI. All experiments were replicated twice.

FACS. After isolation of MCs from five mice from each genotype by MACS using the
anti-GPM6A antibody, the GPM6A+ MCs were further incubated with anti-rat IgG
Alexa Fluor 647 (Life Technologies). Then, the cells were analyzed using FACS Aria
sorter (BD Bioscience) in USC-CSCRM/NCCC Flow Cytometry Core. After excluding
cell debris, GFP+ Alexa Fluor 647+ and GFP- Alexa Fluor 647+ MCs were sorted and
were subjected to QPCR. All experiments were replicated twice.
20
Statistical Analysis. Statistical significance was assessed by ANOVA followed by post-
hoc Tukey HSD test among multiple samples or Student’s t-test between two samples. A
P value of less than 0.05 was considered statistically significant.

21
2.3 Results
Phenotype of Peritoneal MCs
We had previously identified Cd200, Gpm6a, Pdpn, and Upk1b as markers for mouse
liver MCs (Li et al., 2013). Using immunohistochemistry, we examined whether MCs on
the body wall similarly express these markers. In this study, we referred to MCs on the
body wall as "peritoneal MCs" and compared their phenotype with that of "liver MCs" on
the liver surface. Similar to liver MCs, peritoneal MCs expressed GPM6A, PDPN, and
CD200 on the body wall (Fig. 1A-C, arrowheads). Upk1b
RFP
mice expressed RFP
exclusively in PDPN+ MCs (Fig. 1B). No expression of these markers was observed in
fibroblasts beneath MCs (Fig. 1A-C, arrows). MCs expressed both PDPN and VIM,
whereas fibroblasts beneath MCs only expressed VIM (Fig. 1D). Peritoneal MCs
expressed the epithelial markers KRT8 and KRT19 and the mesenchymal markers type I
collagen (COL I) but not CDH1 and ACTA2 (Fig. 1E-I) in the normal body wall. To test
whether peritoneal MCs synthesize Col1a1 mRNA, we analyzed the body wall of a
Col1a1
GFP
transgenic mouse. As shown in Fig. 1J, both PDPN+ MCs and PDPN-
fibroblasts expressed GFP. Peritoneal MCs and fibroblasts were separated by the basal
lamina composed of type IV collagen (COL IV) in the body wall, and both cell types
expressed GFP in normal Col1a1
GFP
mice (Fig. 1K). These results indicate that both
peritoneal and liver MCs share the same marker expression in normal mice.

22
Figure 1

Figure 1. Characterization of peritoneal MCs on the mouse body walls. The body walls
from normal adult (A,C-I), Upk1b
RFP
(B) and Col1a1
GFP
(J,K) mice were immunostained
with GPM6A (Ai), PDPN and RFP (B), CD200 (C), VIM and PDPN (D), KRT8 (E),
KRT19 (F), COL I (G), CDH1 (H), ACTA2 (I), GFP and PDPN (J), and GFP and COL
IV (K). Aii shows a bright field image of Ai. ct; connective tissue, sm; skeletal muscle.
Arrowheads indicate peritoneal MCs on the body wall. Fibroblasts beneath MCs are
indicated by arrows. Peritoneal MCs express GPM6A, PDPN, CD200, KRT8, KRT19,
VIM, and COL I but not CDH1 and ACTA2 in the normal mouse (n=3). The Upk1b
RFP

mouse shows specific expression of RFP in PDPN+ MCs (n=3). MCs and fibroblasts are
23
separated by the basal lamina composed of COL IV, and both cell types express GFP in
the Col1a1
GFP
mouse (n=3). Nuclei were counterstained with DAPI. Scale bar: 10 m.

24
Differentiation of MCs into Myofibroblasts in Peritoneal Fibrosis
To determine whether peritoneal MCs are the source of myofibroblasts in peritoneal
fibrosis, we traced the MC lineage using Wt1
CreERT2
and R26TG
fl
mice. Upon tamoxifen
treatment, the Wt1
CreERT2/+
; R26TG
fl/fl
mouse changes the expression of TOMATO into
membrane-tagged GFP in Wt1+ cells by CreERT2 (Fig. 2A) Although WT1 expression
was not detectable in the peritoneum by immunohistochemistry using available
antibodies, tamoxifen injection resulted in specific GFP expression in 12.5±2.1% of
peritoneal MCs expressing GPM6A or PDPN (Fig. 2B, arrowheads). Importantly, no
GFP expression was observed in fibroblasts beneath MCs in the peritoneum (Fig. 2B,
arrows). The Wt1
+/+
; R26TG
fl/fl
control mouse treated with tamoxifen (Fig. 2B, Cre-) did
not show GFP signals in MCs after tamoxifen injection, indicating specific activity of the
CreERT2 in MCs. Two weeks after tamoxifen injection into Wt1
CreERT2/+
; R26TG
fl/fl
mice,
peritoneal fibrosis was induced by repeated injections of CG, which is known to cause a
thickening of the connective tissue (Kushiyama et al., 2011; Yokoi et al., 2012).
Immunohistochemistry showed that GFP+ MCs begin to express ACTA2 one day after a
single CG injection (Fig. 2Ci, arrowheads). After 3 CG injections, GFP+ cells were
observed in the connective tissue, and the cells showed fibroblastic morphology and
expressed ACTA2 (Fig. 2Cii, double arrows). After 10 CG injections, ACTA2+ GFP+
myofibroblasts were observed in the connective tissue, and 2.1±1.2% of ACTA2+
myofibroblasts expressed GFP (Fig. 2Ciii, double arrows). Considering that all MCs
contribute equally to ACTA2+ myofibroblasts and that the labeling efficiency of
25
tamoxifen to MCs is 12.5%, we estimated that 16.8% of ACTA2+ myofibroblasts are
derived from MCs in peritoneal fibrosis.
One day after a single CG injection, GFP+ MCs still expressed GPM6A+ and
were located above the basal lamina that was composed of COL IV (Fig. 2Di and 2Ei,
arrowheads). After 3 CG injections, the basal lamina began to be disrupted, and the
expression of GPM6A decreased on the surface (Fig. 2Dii and 2Eii, arrowheads). After
10 CG injections, a few GPM6A+ MCs survived, and no expression of COL IV was
found on the surface (Fig. 2Diii and 2Eiii, arrowheads). In addition, no expression of
KRT8 and KRT19 was observed on the surface of the body wall (Fig. 2F, arrowheads),
indicating the disappearance of the MC layer from the body wall. GFP+ myofibroblasts
in peritoneal fibrosis expressed PDPN, COL I, and DES, but not KRT8 and KRT19 in the
connective tissue (Fig. 2F, double arrows). These GFP+ myofibroblasts did not express
F4/80, a marker of macrophages, or CD31, a marker of endothelial cells, in the
connective tissue (Fig. 2F, double arrows). These results demonstrate that MCs undergo
MMT and give rise to myofibroblasts in peritoneal fibrosis.

26
Figure 2

Figure 2. Peritoneal MCs give rise to myofibroblasts in peritoneal fibrosis. (A)
Experimental design. After two tamoxifen injections into Wt1
CreERT2/+
; R26TG
fl/fl
mice,
MCs change from the expression of TOMATO to membrane-tagged GFP. Two weeks
27
after tamoxifen injection, the mice were treated with CG by 10 intraperitoneal injections.
(B) Immunohistochemistry of GFP (green) and GPM6A or PDPN (red). After tamoxifen
injection into Wt1
CreERT2/+
; R26TG
fl/fl
mice, only GPM6A+ or PDPN+ MCs express GFP
(arrowheads). Note that no GFP expression in fibroblasts (arrows) is evident. No
induction of GFP was observed in the control Wt1
+/+
; R26TG
fl/fl
mice (Cre-) after
tamoxifen treatment (n=3). (C-E) Phenotypic changes of MCs in peritoneal fibrosis. The
body wall tissues 1 day after CG injections (1, 3, or 10 times) were immunostained with
GFP (green) and ACTA2, GPM6A, or COL IV (red). After a single CG injection (i;
CGx1, n=2), GFP+ MCs begin to express ACTA2 (arrowheads). After 3 CG injections
(ii; CGx3, n=2), GFP+ cells in the connective tissue show a myofibroblastic morphology
with ACTA2 expression but not with GPM6A (double arrows). After 10 CG injections
(iii; CGx10, n=3), a few MCs expressing GPM6A survives on the body wall
(arrowheads) and no COL IV is observed on the surface of the body wall, indicating
denudation of the MC layer. (F) Immunohistochemistry of GFP (green) and cell markers
(red) after 10 CG injections. Arrowheads indicate the surface of the body wall. MC-
derived GFP+ myofibroblasts express PDPN, COL I, and DES but not KRT8, KRT19,
F4/80, and CD31 (double arrows). Immunostaining without primary antibodies does not
show signals (Negative). Nuclei were counterstained with DAPI. Scale bar: 10 m.

28
Regeneration of Peritoneal MCs
We investigated whether the peritoneal mesothelium regenerates if CG treatment is
discontinued. We labeled MCs as GFP+ cells in Wt1
CreERT2/+
; R26TG
fl/fl
mice, induced
peritoneal fibrosis with 10 CG injections, and allowed the mice to recover by
discontinuing the CG treatment (Fig. 3A). After 10 CG injections, the connective tissue
area was significantly increased from 3,982 m
2
to 62,146 m
2
in sections captured
under a 20X objective, and the area decreased to 18,665 m
2
at 4 weeks after the last CG
injection (Fig. 3B,C), indicating degeneration of the connective tissue. QPCR showed
increased expression of Acta2 and Col1a1 mRNAs by CG injections, and that the mRNA
expression returned to the baseline level 4 weeks after the last injection (Fig. 3D).
Immunohistochemistry showed that GPM6A+ MCs reappeared on the surface of the
body wall, and 17.6±4.2% of MCs co-expressed GFP (Fig. 3E, arrowheads), which is
similar to the original labeling efficiency of MCs (12.5%) in the normal body wall (Fig.
2B). Both GFP+ and GFP- MCs expressed GPM6A, PDPN, KRT8, and KRT19, and
COL IV+ basal lamina reappeared beneath MCs (Fig. 3E-I, arrowheads). Some of the
regenerated MCs still expressed ACTA2 (Fig. 3J, arrowhead). GFP+ ACTA2+
myofibroblasts were rarely observed in the connective tissue (Fig. 3J). To test whether
survived MCs proliferate and participate the regeneration of the mesothelium, we
measured the proliferation by BrdU incorporation assay. One week after last CG
injection, we injected BrdU to mice and collected the liver tissues 4 hours later.
Incorporation of BrdU in nuclei was observed in 2.2±0.9% of MCs (Fig. 3K).
Myofibroblasts in the connective tissue rarely showed the BrdU incorporation
29
(0.2±0.07%). These results indicate that the MC layer regenerates and that peritoneal
fibrosis is reversible in the mouse model.

30
Figure 3

Figure 3. Lineage tracing of MCs during recovery from fibrosis. Regeneration of the MC
layer on the body wall during recovery from peritoneal fibrosis. (A) By tamoxifen
injection into Wt1
CreERT2/+
; R26TG
fl/fl
mice, MCs begin to express GFP. Two weeks after
tamoxifen injection, peritoneal fibrosis was induced by 10 CG injections, and the mice
were allowed to recover for 4 weeks by discontinuing the CG treatment. (B) H&E
staining of the body wall before (normal), after 10 CG injections (CG), and 4 weeks after
the last CG injection (CG+4W). An arrowhead and arrow indicate MCs and fibroblasts,
respectively. (C) Quantification of the areas of connective tissue in the body wall in B
(n=3). **P < 0.01. (D) QPCR of mRNA expression in body wall tissues from the normal
31
(N), fibrosis (CG), and recovered 4 weeks after the CG injection (CG+4W) mice. **P <
0.01. (E-J) Immunohistochemistry of the body wall with GFP (green) and GPM6A (E),
PDPN (F), KRT8 (G), KRT19 (H), COL IV (I), or ACTA2 (J, red) at 4 weeks after the
last CG injection. Arrowheads indicate regenerated peritoneal MCs expressing GFP and
markers. An arrow indicates GFP+ fibroblasts in the connective tissue. (K) BrdU
incorporation in the body wall. One week after the last CG injection, the peritoneum was
analyzed 4 hours after BrdU treatment. Immunohistochemistry for cytokeratin (Keratin,
green) and BrdU (red) showed incorporation of BrdU in 2.2% MCs (arrowheads, n=3,
910 MCs were counted). An arrow indicates rare myofibroblasts showing nuclear BrdU
staining. Nuclei were counterstained with DAPI. Scale bar: 50 m (B) and 10 m (E-K).

32
Isolation and Culture of Peritoneal MCs
To examine the mechanisms underlying MMT, we isolated peritoneal MCs from the
body wall using anti-GPM6A antibodies and MACS. The separated GPM6A+ cells
expressed mRNAs for MC markers (Gpm6a, Cd200, Msln, Pdpn, Upk1b, Wt1) and
epithelial cell markers (Krt8, Krt19) compared with GPM6A- cells (Fig. 4A), validating
the successful purification of peritoneal MCs. In culture, peritoneal MCs formed
epithelial colonies up to 7 days (Fig. 4B). On day 14, some MCs lost the epithelial cell
phenotype and showed a mesenchymal morphology. QPCR showed increased expression
of Acta2 in cultured MCs (Fig. 4C).
TGF-  was shown to be a key factor for MMT in the liver (Li et al., 2013). TGF-
 binds to a TGF-  type II receptor (TGFBR2) dimer, induces phosphorylation of TGF- 
type I receptor (TGFBR1), and stimulates cells via downstream effectors including
SMAD2/3 (Lamouille et al., 2014). In primary peritoneal MCs, TGF-  induced a
morphological change of MCs into myofibroblasts (Fig. 4D). According to this change,
MCs expressed ACTA2 and increased expression of Acta2 mRNA (Fig. 4E,F). A
chemical inhibitor (SB431542) of TGFBR1 blocked the MMT induced by TGF-  (Fig.
4D-F). Peritoneal MCs also increased the expression of Ctgf and Pdpn mRNAs in
response to TGF-  (Fig. 4F). Different from liver MCs (Li et al., 2013), peritoneal MCs
did not decrease Gpm6a mRNA in response to TGF-  (Fig. 4F). TGF-  treatment
induced the nuclear localization of phosphorylated-SMAD3 in MCs (Fig. 4G). These
results indicate that TGF-  induces MMT in peritoneal MCs.
33
We confirmed that Wt1+ MCs give rise to ACTA2+ myofibroblasts in culture.
After tamoxifen injection into Wt1
CreERT2/+
; R26TG
fl/fl
mice, peritoneal MCs were isolated
by MACS. After 1 week in culture, GPM6A+ MCs expressed either TOMATO or GFP
(Fig. 4H). After TGF-  treatment, the induction of ACTA2 was observed in both GFP+
and GFP- myofibroblasts (Fig. 4I), demonstrating that Wt1+ MCs give rise to ACTA2+
myofibroblasts in culture.

34
Figure 4

Figure 4. Induction of peritoneal MMT by TGF-  in vitro. Peritoneal MCs were isolated
from the mouse body wall using anti-GPM6A antibodies and MACS. (A) QPCR of
GPM6A- (-) and GPM6A+ cells (+) separated from body wall cells by MACS. The
GPM6A+ fraction highly expresses mRNAs for Gpm6a, Cd200, Msln, Pdpn, Upk1b,
Wt1, Krt8, and Krt19. (B) Culture of GPM6A+ peritoneal MCs. MCs form epithelial
35
colonies. Some MCs lose epithelial cell morphology in culture. (C) QPCR of peritoneal
MCs in culture. MCs increase the expression of Acta2 mRNA in accordance with the
morphological changes shown in B. (D) Peritoneal MCs were cultured in the presence or
absence of TGF-  and an inhibitor for TGFBR1 (SB431542). TGF-  induces MMT,
whereas SB431542 blocks TGF- -induced morphological changes. (E)
Immunocytochemistry of cultured peritoneal MCs with antibodies against ACTA2 (red)
and GPM6A (green). TGF-  strongly induces the expression of ACTA2, whereas
SB431542 blocks the induction of ACTA2. (F) QPCR of cultured peritoneal MCs in the
presence of DMSO (-),TGF-  (TG), SB431542 (SB), and both TGF-  and SB431542
(TG+SB). TGF-  induces the expression of Acta2, Ctgf, Gpm6a, and Pdpn mRNAs. (G)
Immunocytochemistry of cultured peritoneal MCs. TGF-  induces the nuclear
localization of phosphorylated-SMAD3 (P-SMAD3, green). (H,I) Lineage tracing of
Wt1+ peritoneal MCs in culture. After tamoxifen injection to Wt1
CreERT2/+
; R26TG
fl/fl

mice, GPM6A+ MCs were isolated by MACS. (H) MCs express either TOMATO (red)
or GFP (green). Arrowheads indicate MCs co-expressing GFP (green) and GPM6A (red).
(I) Upon TGF-  treatment (+TGF- ), both GFP+ (arrowheads) and GFP- MC-derived
myofibroblasts (arrow) express ACTA2. No signals were detected in a negative control
without primary antibodies (Negative). Nuclei were counterstained with DAPI. Scale bar:
20 m (B,D) and 10 m (E,G-I).

36
Induction of MMT by TGF- 
Next, we examined whether TGF-  is responsible for the induction of MMT in peritoneal
fibrosis. After labeling MCs as GFP+ cells in Wt1
CreERT2/+
; R26TG
fl/fl
mice, peritoneal
fibrosis was induced by CG injections combined with treatment using a soluble TGFBR2
(STR) for antagonizing TGF-  signaling or control IgG (Fig. 5A). H&E staining showed
that the STR treatment reduces the development of the connective tissue in the body wall
compared with the control IgG (Fig. 5B). The STR treatment significantly reduced the
area of connective tissue from 63,017 m
2
to 15,657 m
2
(Fig. 5C).
Immunohistochemistry showed that the MC layer disappears from the body wall surface
and ACTA2+ GFP+ myofibroblasts accumulate in the connective tissue in the IgG group
(Fig. 5D). However, the STR group showed that GPM6A+ GFP+ MCs are present on the
surface, and few GFP+ myofibroblasts were observed in the connective tissue (Fig. 5D).
These data indicate that antagonism of TGF-  signaling inhibits MMT and peritoneal
fibrosis.

Conditional deletion of Tgfbr2 gene in MCs
To further delineate whether direct TGF-  signaling in MCs rather than indirect effects of
TGF-  via other cell types is responsible for MMT, we conditionally deleted exon 4 of
the Tgfbr2 gene in Wt1+ MCs. We produced Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
mice and
conditionally deleted the Tgfbr2 gene in Wt1+ MCs by tamoxifen injection (Fig. 5E).
CreERT2 abolishes the expression of Tgfbr2 while inducing GFP expression. To
37
determine the efficiency of the Tgfbr2 gene knockout, we injected tamoxifen twice into
Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
mice and isolated MCs from the body wall. To
efficiently sort MCs by FACS, we first enriched MCs by MACS using an anti-GPM6A
antibody anti-rat IgG microbeads. Then, GPM6A+ MCs were labeled with anti-rat IgG
Alexa Fluor 647 antibodies and were separated into GFP+ GPM6A+ and GFP- GPM6A+
MCs by FACS (Fig. 5F). QPCR showed a 77% reduction of Tgfbr2 mRNA in the GFP+
MCs compared with GFP- MCs, while increased expression of GFP was observed in the
Tgfbr2
fl/fl
conditional knockout mouse (Fig. 5G, fl/fl), validating the selective deletion of
the Tgfbr2 gene in the GFP+ MCs. The deletion of Tgfbr2 gene reduced the expression of
Gpm6a mRNA (Fig. 5G). The expression of Tgfb1, Tgfb2, and Ctgf mRNAs was not
changed between GFP- and GFP+ MCs (Fig. 5G). After 10 CG injections, the area of
connective tissue was significantly decreased in Tgfbr2
fl/fl
mice compared with Tgfbr2
fl/+

mice (Fig. 5H,I). GFP+ ACTA2+ myofibroblasts were observed in the connective tissue,
and only 2.1±1.7% of GPM6A+ MCs were found on the body wall surface in the control
Tgfbr2
fl/+
mice (Fig. 5J). However, Tgfbr2
fl/fl
mice showed few GFP+ ACTA2+
myofibroblasts in the connective tissue, and 20.4±18.7% of the body wall surface was
covered with GPM6A+ MCs (Fig. 5J). We confirmed down-regulation of nuclear
phosphorylated-SMAD2/3 in GFP+ MCs in Tgfbr2
fl/fl
mice (Fig. 5K, double
arrowheads).

38
Figure 5

Figure 5. Essential role of TGF-  signaling in peritoneal MMT in fibrosis. TGF-
 signaling was inhibited using a soluble form of TGFBR2 (STR) (A-D) or conditional
deletion of the Tgfbr2 gene in peritoneal MCs (E-J). (A) Experimental design. After
39
labeling peritoneal MCs as GFP+ cells by tamoxifen injection in Wt1
CreERT2/+
; R26TG
fl/fl

mice, peritoneal fibrosis was induced by CG injections with either STR or IgG injection 5
times. (B) H&E staining of the body wall treated with an IgG control or STR in CG-
induced fibrosis. (C) Quantification of the area of the connective tissue treated with an
IgG control or STR in B (n=3). ** P < 0.01. (D) Immunohistochemistry of the body wall
using anti-GFP antibodies (green) and ACTA2 or GPM6A antibodies (red). Arrowheads
and arrows indicate MCs and myofibroblasts, respectively. The STR treatment inhibits
MMT on the body wall. (E) Experimental design of the conditional deletion of the Tgfbr2
gene. The tamoxifen-induced CreERT2 in peritoneal MCs activates the expression of
GFP while deleting exon 4 of the Tgfbr2 gene. (F) FACS of MCs isolated from the fl/+ or
fl/fl mouse. After isolation of MCs using the anti-GPM6A antibody and MACS, the cells
(MACS+) were further incubated with anti-rat IgG Alexa Fluor 647 to efficiently detect
GPM6A antibody+ MCs. GFP+ GPM6A+ and GFP- GPM6A+ MCs were sorted by
FACS from the fl/+ or fl/fl Tgfbr2 knockout mouse. MCs (Cre- MACS+) isolated from
Wt1
+/+
; R26TG
fl/fl
; Tgfbr2
fl/+
or Tgfbr2
fl/fl
mice and Cre+ MACS- fractions were used as
negative controls. (G) QPCR of the GFP+ GPM6A+ (+) and GFP- GPM6A+ (-) MCs.
GFP+ GPM6A+ MCs decreased Tgfbr2 mRNA expression, while increasing Gfp mRNA
in the homozygous (fl/fl) MCs. (H) H&E staining of the fibrotic body wall induced by
CG from the Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/+
(fl/+) or Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
(fl/fl) mouse. (I) Quantification of the area of connective tissue in Tgfbr2
fl/+
and Tgfbr2
fl/fl

mice treated with 10 CG injections in H (n=3). * P < 0.05. (J,K) Immunohistochemistry
of the body wall from Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/+
(fl/+) and Wt1
CreERT2/+
;
40
R26TG
fl/fl
; Tgfbr2
fl/fl
(fl/fl) mice treated with 10 CG injections using anti-GFP antibodies
(green) and ACTA2, GPM6A, or phosphorylated-SMAD2/3 antibodies (P-SMAD2/3,
red). Arrowheads and arrows indicate MCs and myofibroblasts, respectively. (J) The MC
layer remains on the body wall after the conditional deletion of Tgfbr2 in MCs
(arrowheads). (K) Double arrowheads indicate GFP+ MCs without P-SMAD2/3
expression. Nuclei were counterstained with DAPI. Bars, 100 m (B,H) and 10 m (D,J).

41
Role of Tgfbr2 Gene in Regeneration
We isolated MCs from R26TG
fl/fl
; Tgfbr2
fl/fl
mice and analyzed the effects of the
deletion of Tgfbr2 gene. After treatment with Adenovirus carrying Cre, MCs started to
express GFP and lost Tgfbr2 mRNA expression (Fig. 6A,B). Interestingly, TGF- 1
treatment strongly down-regulated the Tgfbr2 gene expression in MCs treated with the
control Adenovirus carrying LacZ (Fig. 6B), suggesting a negative feedback regulation of
TGF-  signaling. Upon treatment with TGF- 1, MCs infected with Adenovirus carrying
control LacZ induced nuclear localization of phosphorylated-SMAD3 and the expression
of Acta2 and Ctgf mRNAs (Fig. 6A,B). In contrast, after Cre expression, GFP+ MCs did
not show phosphorylated-SMAD3 expression in their nuclei by TGF- 1 and increased
expression of Acta2 and Ctgf mRNAs (Fig. 6A,B). These data indicate that the canonical
TGF-  signaling induces the myofibroblastic conversion of peritoneal MCs in vitro.
We also investigated the effects of Tgfbr2 gene deletion in MCs during regeneration
of the peritoneum from CG-induced peritoneal fibrosis (Fig. 6C). One month after the
last CG injection, the connective tissue became thinner in both Tgfbr2
fl/+
and Tgfbr2
fl/fl

mice (Fig. 6D,E) compared to before regeneration (Fig. 5H,I). Immunohistochemistry
showed regeneration of GPM6A+ MCs on the body wall surface in both genotypes (Fig.
6F). A few ACTA2+ myofibroblasts were still observed in the Tgfbr2
fl/+
but not in the
Tgfbr2
fl/fl
peritoneum (Fig. 6G). GFP was observed 6.8±2.4% and 2.6±1.3% of MCs in
the Tgfbr2
fl/+
and Tgfbr2
fl/fl
body walls, respectively.

42
Figure 6

Figure 6. Deletion of Tgfbr2 gene in peritoneal MCs. (A,B) Deletion of Tgfbr2 gene in
cultured peritoneal MCs. GPM6A+ MCs were isolated from five R26TG
fl/fl
; Tgfbr2
fl/fl

mouse body walls. Cultured MCs were infected with an Adenovirus vector (Ad) carrying
LacZ or Cre (MOI 50) and were treated with TGF- 1 for 3 hours (A) or 12 hours (B). (A)
Immunocytochemistry of MCs with GFP (green) and phosphorylated-SMAD3 (P-
SMAD3, red). Arrowheads indicate P-SMAD3 in MCs treated with Ad-LacZ and TGF-
1. Nuclei were counterstained with DAPI. (B) QPCR of MCs infected with Ad-LacZ or
43
Cre followed by treatment with TGF- 1. ** P < 0.01. (C-G) Lineage tracing of MCs
during recovery from fibrosis. (C) Experimental design of the conditional deletion of the
Tgfbr2 gene. The tamoxifen-induced CreERT2 in peritoneal MCs activates the
expression of GFP while deleting the Tgfbr2 gene. Two weeks after tamoxifen injection,
peritoneal fibrosis was induced by 10 CG injections, and the mice were allowed to
recover for 4 weeks by discontinuing the CG treatment. (D) H&E staining of the body
wall 4 weeks after the last CG injection in the Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/+
(fl/+) or
Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
(fl/fl) mouse. (E) Quantification of the area of
connective tissue 4 weeks after the last CG injection in D. ns; statistically not significant
(n=3). (F,G) Immunohistochemistry of the body wall 4 weeks after the last CG injection
in Tgfbr2
fl/+
(fl/+) and Tgfbr2
fl/fl
(fl/fl) mice using anti-GFP antibodies (green) and
GPM6A or ACTA2 (red). Arrowheads and arrows indicate MCs and myofibroblasts,
respectively. Bars, 10 m (A,F,G) and 50 m (D).

44
CG Causes Liver Injury
CG has been used to induce peritoneal fibrosis in the body wall by causing injury to
peritoneal MCs (Kushiyama et al., 2011; Yokoi et al., 2012). We examined whether 10
CG injections into the peritoneal cavity also causes injury to MCs in the liver as a
visceral organ. Using the same Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/+
or Tgfbr2
fl/fl
mouse and
CG treatment, we compared the phenotypes of liver and peritoneal MCs (Fig. 7A). By
tamoxifen injection, 14.5±6.8% of GPM6A+ MCs began to express GFP in the normal
liver, and no GFP expression was observed except in MCs (Fig. 7B). MCs did not
express ACTA2 in the normal liver (Fig. 7B). Sirius red staining revealed that normal
mouse liver has thin collagen lamella beneath MCs and its thickness is around 2.4±0.9
m (Fig. 7C, D). After CG injections, the thickness of the collagen lamella was
increased to 10.2±1.5 m beneath the liver surface in Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
+/+

control mice (Fig. 7C, D). The increase of collagen by CG treatment was reduced to
4.8±0.8 m in Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
knockout mice (Fig. 7C, D).
Immunohistochemistry showed that CG treatment induced accumulation of ACTA2+
myofibroblasts only beneath the liver surface in control mice and 11.9±9.2% of ACTA2+
myofibroblasts co-expressed GFP (Fig. 7E). Compared to control or Tgfbr2 heterozygous
mice, Tgfbr2 knockout mice showed less accumulation of ACTA2+ myofibroblasts
beneath MCs and a few GFP+ ACTA2+ myofibroblasts were observed (Fig. 7E). The
cell lineage tracing demonstrates that CG induces MMT in both liver and peritoneal MCs
and TGF-  signaling in MCs is responsible to MMT. Different from the peritoneum,
45
MCs were negative for ACTA2 on the injured liver surface and GPM6A+ MCs remained
on the liver surface (Fig. 7E,F). ACTA2+ GFP+ myofibroblasts lost the expression of
GPM6A, a marker of MCs, and were observed up to 55 m in depth from the surface,
and no GFP+ cells were found beyond this distance (Fig. 7E,F). One month after the last
CG injection, the thickness of the collagen fiber was reduced to 4 to 5.5 m in all
genotypes (Fig. 7C,D). Immunohistochemistry showed the presence of GFP+ MCs on the
liver surface and without ACTA2 expression after regeneration (Fig. 7H).

Distinct Origins of Peritoneal and Liver MCs
In contrast to liver MCs, peritoneal MCs begin to express ACTA2 upon CG treatment,
and the MC layer largely disappears in peritoneal fibrosis (Fig. 2C and D). During
regeneration, the surviving peritoneal MCs might be the source of regeneration of the
peritoneal mesothelium. In addition to this possibility, visceral MCs might traverse the
peritoneal cavity, integrate into the parietal mesothelium, and participate in regeneration
of the peritoneal MC layer during regeneration. In fact, floating MCs were isolated from
the peritoneal cavity in patients who underwent peritoneal dialysis (Yanez-Mo et al.,
2003). To test whether liver MCs traverse the peritoneal cavity and form a peritoneal MC
layer in CG-induced fibrosis, we analyzed the developmental lineages of liver and
peritoneal MCs. We previously reported that liver MCs are derived from Mesp1+
mesoderm (Asahina et al., 2009; Lua et al., 2014). MESP1 is a transcription factor that is
transiently expressed in nascent mesoderm during gastrulation and contributes to a part of
the mesoderm (Saga et al., 1999). Using Mesp1
Cre/+
; R26TG
fl/fl
mice, we evaluated
46
whether both liver and peritoneal MCs are derived from the same mesodermal origin
(Fig. 7I). In the normal Mesp1
Cre/+
; R26TG
fl/fl
liver, 48.8±11.7% of MCs express GFP
(Fig. 7J, arrowheads). By contrast, no GFP+ MCs were observed in the body wall (Fig.
7J, arrowheads). Hepatic stellate cells in the liver and fibroblasts in the peritoneum
expressed GFP (Fig. 7J, arrows). The cell lineage tracing indicates that the origin of
peritoneal MCs is distinct from that of liver MCs.
Next, we analyzed whether liver MCs contribute to the regeneration of the
peritoneal MC layer. We induced peritoneal fibrosis using 10 CG injections into the
Mesp1
Cre/+
; R26TG
fl/fl
mice and analyzed the body wall 4 weeks after the last CG
injection (Fig. 7I). After recovery from CG-induced injury, GFP expression was observed
in fibroblasts in the connective tissue (Fig. 7K, arrows), but was rare in regenerated MCs
in the body wall (Fig. 7K, arrowheads), demonstrating a negligible contribution of liver
MCs to the regeneration of the peritoneal mesothelium during the recovery from
peritoneal fibrosis.

47
Figure 7

Figure 7. Lineage tracing of liver and peritoneal MCs in CG-induced injury. (A-H)
Lineage tracing of MCs in the liver using Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
mice. (A)
Experimental design. After two tamoxifen injections into Wt1
CreERT2/+
; R26TG
fl/fl
;
48
Tgfbr2
fl/fl
mice, MCs begin to express GFP and lose Tgfbr2 mRNA expression. Two
weeks after tamoxifen injection, the mice were treated with CG by 10 intraperitoneal
injections. Four weeks after the last CG injection, the liver tissues were analyzed to
examine the recovery. (B) Immunohistochemistry of GFP (green) and GPM6A or
ACTA2 (red) in the normal liver. GFP expression is observed in GPM6A+ MCs
(arrowhead) but not fibroblasts (arrow) beneath MCs. Liver MCs do not express ACTA2
(arrowheads). (C) Sirius red staining of the liver after 10 CG injections (CG) or 4 weeks
after the last CG injection (CG+4W). Sirius Red staining shows thin collagen lamella
beneath the liver surface. (D) Quantification of the length of the collagen lamella in C
(n=3). * P < 0.05, ** P < 0.01. (E-G) After labeling MCs as GFP+ cells in Wt1
CreERT2/+
;
R26TG
fl/+
; Tgfbr2
fl/+
or Tgfbr2
fl/fl
mice, liver injury was induced by 10 CG injections. (E)
GFP+ ACTA2+ myofibroblasts (arrows) are observed beneath MCs (arrowheads) in the
Tgfbr2
fl/+
(fl/) mice. MCs do not express ACTA2 (arrowheads). Tgfbr2
fl/fl
(fl/fl) liver
shows less accumulation of ACTA2+ myofibroblasts. (F) MCs remain on the liver
surface and express GPM6A upon liver injury (arrowheads). Arrows indicate GFP+ MC-
derived myofibroblasts beneath MCs. (G) Phosphorylated-SMAD2/3 (P-SMAD2/3, red)
is localized in the nuclei of MCs (arrowheads), myofibroblasts (arrows), and hepatocytes
(large round nuclei) by CG treatment in the Tgfbr2
fl/+
liver. Tgfbr2
fl/fl
liver shows down-
regulation of P-SMAD2/3 in GFP+ cells (double arrowheads), indicating the loss of
TGF-  signaling in GFP+ cells. (H) Four weeks after the last CG injection, the liver
tissues were analyzed by immunohistochemistry for GFP (green) and ACTA2 or GPM6A
(red) in Tgfbr2
fl/+
or Tgfbr2
fl/fl
mice. GFP+ MCs are negative for ACTA2 (arrowheads)
49
and positive for GPM6A (arrowheads). An arrow indicates GFP+ cells beneath MCs in
regenerating liver. (I-K) Mesoderm lineage tracing in the body wall and liver. (I)
Mesodermal lineage was traced using Mesp1
Cre/+
; R26TG
fl/fl
mice (n=4). (J) Normal liver
and body wall tissues from the Mesp1
Cre/+
; R26TG
fl/fl
mouse were immunostained with
GFP (green) and PDPN (red). Liver MCs, but not peritoneal MCs, express GFP
(arrowheads). Arrows indicate GFP+ hepatic stellate cells in the liver and fibroblasts in
the body wall. (K) Peritoneal fibrosis was induced by 10 CG injections into Mesp1
Cre/+
;
R26TG
fl/fl
mice, and the mice were allowed to recover for 4 weeks by discontinuing the
CG treatment. The body wall tissues were immunostained with GFP (green) and PDPN
or ACTA2 (red) 4 weeks after 10 CG injections. Note that no GFP expression is evident
in regenerated peritoneal MCs (arrowheads). Arrows indicate fibroblasts expressing GFP
in the connective tissue. Nuclei were counterstained with DAPI. Bars, 10 m (B,E-H,J,K)
and 20 m (C).

50
2.4 Discussion
The transition of jCs to myofibroblasts has been suggested during peritoneal fibrosis.
However, no lineage tracing study using mouse genetics was conducted to demonstrate
this hypothesis. With the use of conditional cell lineage tracing, this study demonstrated
for the first time that peritoneal MCs give rise to myofibroblasts in a mouse peritoneal
fibrosis model. Based on the labeling efficiency of MCs in the Wt1
CreERT2/+
; R26TG
fl/fl

mouse, we estimated that 16.8% of myofibroblasts are derived from peritoneal MCs.
What are other sources of myofibroblasts during peritoneal fibrosis? Fibroblasts that are
positive for VIM exist between MCs and skeletal muscles in the peritoneum and are
likely to be another source of myofibroblasts (Fig. 8). Although we attempted to trace
lineages of fibroblasts using Col1a1
CreERT2
mice (Jackson laboratory, #016241), we failed
to label fibroblasts by tamoxifen injections (data not shown). More CreERT2 mouse lines
will be necessary to precisely estimate the contribution of fibroblasts to myofibroblasts
during peritoneal fibrosis. Nevertheless, the present study demonstrates that MCs
partially contribute to myofibroblasts in peritoneal and liver fibrosis (Fig. 8).
Using peritoneal fibrosis induced by the injection dialysis solution or TGF-  in the
same Wt1
CreERT2
or Col1a1
GFP
mice, Chen et al. reported that the contribution of MCs to
myofibroblasts is rare in the liver (Chen et al., 2014). By treating Wt1
CreERT2
mice with
tamoxifen 10 times through oral gavage, the authors labeled 83% of MCs and 24% of
fibroblasts beneath MCs in the liver and concluded that fibroblasts beneath MCs are the
major source of myofibroblasts in the liver. In addition, they reported that MCs are
negative for GFP in the Col1a1
GFP
mouse liver (Chen et al., 2014). Chen et al. did not
51
analyze the body wall in these models. In our experimental condition, we labeled 12.5
and 14.5% of MCs in the body wall and liver, respectively, in the Wt1
CreERT2
mice, and
we did not observe any labeling in fibroblasts in these tissues. Different from their report,
we found that the normal Col1a1
GFP
mouse expresses GFP in both MCs and fibroblasts in
the normal body wall and liver. These differences may cause different outcomes between
the present study and their study.
TGF-  has been shown to be a major inducer of MMT in peritoneal fibrosis (Patel
et al., 2010; Strippoli et al., 2010; Duan et al., 2014). Similar to previous reports, we
found that TGF-  induces MMT in peritoneal MCs (Fig. 8). Suppression of TGF- 
signaling with STR significantly reduced the number of MC-derived myofibroblasts and
the area of the connective tissue induced by CG injections. Furthermore, the MC layer
was protected by STR treatment in the CG model. TGF-  is known to induce fibrosis in
organs by activating myofibroblasts, whereas it suppresses inflammation and
carcinogenesis in epithelial cells (Lamouille et al., 2014). In fact, the overexpression of a
STR plasmid enhanced inflammation and failed to attenuate peritoneal fibrosis induced
by TGF-  (Motomura et al., 2005). Canonical TGF-  signaling is mediated by SMAD2/3
(Lamouille et al., 2014). In MCs, SMAD3, but not SMAD2, mediates MMT (Patel et al.,
2010; Duan et al., 2014). In addition to the canonical pathway, the TGF- /mTOR
pathway is also involved in MMT (Patel et al., 2010; Lamouille et al., 2014). In TGF- -
induced MMT, SNAIL1 has been shown to down-regulate CDH1 expression and thereby
induce MMT, similar to the mechanism of cancer metastasis (Strippoli et al., 2010; Zhou
52
et al., 2013; Lamouille et al., 2014). However, we could not detect CDH1 expression in
either peritoneal or liver MCs by immunohistochemistry in mice (Li et al., 2013). We
assume that SNAIL1-mediated down-regulation of CDH1 may not be involved in MMT
in mice.
Using Wt1
CreERT2/+
; R26TG
fl/fl
; Tgfbr2
fl/fl
mice, we conditionally deleted Tgfbr2 in
Wt1+ MCs. We visualized the migration and differentiation of Tgfbr2-null MCs as GFP+
cells in the body wall. Compared with Tgfbr2
fl/+
heterozygous mice, Tgfbr2
fl/fl

homozygous knockout mice treated with CG showed few GFP+ myofibroblasts in the
connective tissue, demonstrating that direct TGF-  signaling in MCs is essential for
MMT. Although CreERT2-mediated recombination occurred only in 12.5% of MCs in
our condition using the Wt1
CreERT2
mouse, the conditional Tgfbr2 knockout mouse
showed the presence of mesothelium on the body wall and a reduction in the connective
tissue area induced by CG injections, indicating that protecting the intact MC layer is
beneficial in preventing peritoneal fibrosis.
We identified the expression of GPM6A, CD200, PDPN, UPK1B, KRT8,
KRT19, VIM, and COL I in both peritoneal and liver MCs. GPM6A is a four-
transmembrane domain-protein and regulates neurite growth (Alfonso et al., 2005).
Expression of Gpm6a gene is down-regulated in neurons by stress. Gpm6a-deficient mice
develop normally, but show abnormal behavior by stress (El-Kordi et al., 2013).
However, nothing is known about the function of GPM6A in MCs. We previously
reported that liver MCs down-regulate mRNA and protein expression of GPM6A by
TGF-  treatment (Li et al., 2013). Intriguingly, the present study showed that peritoneal
53
MCs down-regulate GPM6A protein expression and up-regulate Gpm6a mRNA
expression by TGF-  treatment, implying that the phenotype of peritoneal MCs is not
identical to that of liver MCs. In fact, we demonstrated a distinct origin of peritoneal and
liver MCs. At early embryogenesis, the lateral plate mesoderm is divided into the somatic
and splanchnic mesoderm that gives rise to the parietal and visceral mesothelium,
respectively (Thomason et al., 2012). MESP1 is a basic helix-loop-helix transcription
factor that is transiently expressed in the nascent mesoderm at the onset of gastrulation,
and Mesp1+ mesoderm contributes to a part of the lateral plate mesoderm (Saga et al.,
1999). In the normal liver, Mesp1+ mesoderm gives rise to MCs and hepatic stellate cells
that reside in the space of Disse (Asahina et al., 2009; Lua et al., 2014). In liver fibrosis,
both cells give rise to myofibroblasts (Lua et al., 2014). The present study showed that
Mesp1+ mesoderm does not contribute to parietal MCs in mice, demonstrating distinct
origins of MCs in the liver and body wall (Fig. 8).
Peritoneal and liver MCs change phenotypes differently in CG-induced peritoneal
fibrosis. Upon CG injection, peritoneal MCs on the body wall begin to express ACTA2
and differentiate into myofibroblasts. The MC layer largely disappears after repeated CG
injections. However, liver MCs continue to cover the liver surface after CG injections
and do not express ACTA2. Liver MCs give rise to ACTA2+ myofibroblasts that migrate
beneath the MC layer, while the MC layer remains on the liver surface (Fig. 8).
Myofibroblasts derived from peritoneal MCs but not liver MCs express PDPN in the
connective tissue. TGF-  suppresses Gpm6a mRNA in the liver (Li et al., 2013), but not
in peritoneal MCs. Our data indicate that peritoneal MCs and liver MCs are inherently
54
different. Shelton et al. reported that visceral MCs express autotaxin and have more
migration capacity compared with parietal MCs (Shelton et al., 2013). A recent study
using organ transplantation revealed that visceral MCs in the developing lung are derived
from the lung but not from other organs (Winters et al., 2014). The intrinsic difference
between parietal and visceral MCs may underlie the differential responses of liver and
peritoneal MCs to injury caused by CG.
The peritoneal fibrosis induced by CG injections is reversible (Nishioka et al.,
2008). In our model, the MC layer of the body wall is regenerated in 4 weeks after CG
injections. Interestingly, regenerated MCs are negative for GFP in Mesp1
Cre/+
; R26TG
fl/fl

mice, refuting the possible contribution of liver MCs or body wall fibroblasts to the
regenerated peritoneal MCs. After reversal from peritoneal fibrosis in Wt1
CreERT2/+
;
R26TG
fl/fl
mice, we found that 17.6% of peritoneal MCs express GFP, which is similar to
the original labeling efficiency of peritoneal MCs (12.5%) in this model, implying that
surviving MCs are likely to be the source of regenerating MCs on the body wall (Fig. 8).
Another possible source is that MC-derived myofibroblasts revert to MCs during
recovery from peritoneal fibrosis. However, the Wt1
CreERT2/+
; R26TG
fl/fl
mouse model
cannot delineate whether GFP+ MCs on the regenerated peritoneum are derived from
survived MCs or reversal of MC-derived myofibroblasts and this hypothesis remains to
be determined.
In summary, the present study demonstrated that MCs partially contribute to
myofibroblasts in peritoneal fibrosis. Conditional deletion of Tgfbr2 in Wt1+ MCs
suppressed MMT and reduced peritoneal fibrosis. The protection of MCs will be a
55
therapeutic target for the prevention and suppression of peritoneal fibrosis.

56
Figure 8

Figure 8. A proposed model of CG-induced fibrosis. In the body wall, a single layer of
MCs covers the body wall and basal lamina is formed between MCs and fibroblasts. On
the other hand, on the liver surface, no basal lamina is formed between MCs and
fibroblasts. Mesp1+ mesoderm gives rise to liver MCs, but not peritoneal MCs. Upon
injury caused by CG injections to the peritoneal cavity, both peritoneal and liver MCs
undergo MMT and differentiate into myofibroblasts expressing ACTA2. In the body
wall, the MC layer and basal lamina largely disappear upon injury. On the other hand,
57
liver MCs remain on the liver surface, while migrating inward and differentiating into
myofibroblasts in CG-induced injury. Conditional cell lineage tracing demonstrated that
17% of ACTA2+ myofibroblasts are derived from MCs in peritoneal fibrosis in the body
wall. Thus, both MCs and fibroblasts are likely to be the source of myofibroblasts in
peritoneal fibrosis. TGF-  induces MMT of MCs in both peritoneal and liver MCs. When
CG treatment is discontinued, the MC layer regenerates on the body wall. Liver MCs do
not traverse the body cavity and contribute to regeneration of peritoneal MC layer.

58
Chapter 3
Isolation of hepatic stellate cells, portal fibroblasts, and mesothelial
cells from mouse livers and quantification of their contribution to
myofibroblasts in liver fibrosis
3.1 Introduction
Previous chapter has described the contribution of MCs in liver fibrosis. This chapter will
focus on portal fibroblasts (PFs), another liver mesenchymal cell that gives rises to
myofibroblasts in liver injury. As its name implies, PFs are fibroblasts found adjacent to
the bile duct and portal vein in the portal triad and uniquely express COL15A1, elastin
(ELN), ectonucleoside triphosphate diphosphohydrolase-2
(ENTPD2/NTPDase2/CD39L1), and THY1 in rat livers (Dranoff et al., 2002; Uchio et
al., 2002; Wells et al., 2004; Li et al., 2007; Lemoinne et al., 2015). It is believed that
their primary function is to maintain the integrity of bile duct, portal vein, and hepatic
artery in the portal triad. Morphologically, PFs show a spindle shape and are
distinguished from HSCs that reside in the space of Disse in the sinusoid. Unlike the
HSCs, PFs do not store VitA lipids in the portal triad.
In Chapter 1, we briefly mentioned that myofibroblasts synthesize extracellular
matrix proteins and participates in liver fibrosis. Depending on the etiology,
myofibroblasts appear at different locations in fibrotic livers. When hepatocytes are
injured caused by virus infection or alcohol intake, damaged hepatocytes release
inflammatory cytokines and soluble factors that activate HSCs near the damaged
hepatocytes. These ACTA2+ activated HSCs/myofibroblasts appeared around the central
vein with accumulated extracellular matrix proteins (Friedman, 2010). In contrast,
59
obstruction of the bile duct leads to accumulation of bile acid in the liver, causes damage,
and develops biliary fibrosis around the bile duct (Hirschfield et al., 2010). In biliary
fibrosis, PFs have been suggested to be the major source of myofibroblast near the bile
duct (Iwaisako et al., 2014). However, conflict results were reported, in which HSCs are
the major source of myofibroblasts even in biliary fibrosis (Mederacke et al., 2013).
Other than the HSCs and PFs, myofibroblasts appears on the liver surface are derived
from MCs and this fibrosis pattern is defined as capsular fibrosis (Lua et al., 2015). Other
liver mesenchymal cells might contribute to liver fibrosis as well. Second-layer cells
(SLCs) in the central vein and capsular fibroblasts (CFs) beneath the mesothelium were
characterized based on their morphology and location in the liver (Bhunchet and Wake,
1992). However, due to the lack of availability of markers and isolation methods, little is
known about the contribution of each cell type to liver fibrosis.
The present study aimed to develop a new isolation method of PFs from mouse
livers to study their conversion to myofibroblasts comparing to that of HSCs and MCs.
We separated PFs, HSCs, and MCs from Col1a1
GFP
transgenic mouse livers with FACS
by the combined expression of VitA lipid autofluorescence, GFP and GPM6A. We also
quantified the contribution of these cells to myofibroblasts in liver fibrosis induced by
bile duct ligation (BDL), 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC) diet feeding,
or carbon tetrachloride (CCl
4
) injections.

60
3.2 Materials
Mice. Col1a1
GFP
and Upk1b
RFP
mice were described in Chapter 2. Fibrosis was induced
by 0.1% DDC diet feeding for 1 month, subcutaneous injection of CCl
4
(1 ml/kg body
weight; n=3) with mineral oil in a 1:3 dilution every third day for a total of 12 injections
or BDL for 3 weeks (Preisegger et al., 1999; Li et al., 2013; Lua et al., 2014). All animal
experiments were performed in accordance with the NIH guidelines under the protocol
approved by the IACUC at the University of Southern California.

Immunohistochemistry. Immunohistochemistry method was described in Chapter 2. The
primary antibodies and additional treatment are listed in Table 1.The primary antibodies
were detected with secondary antibodies conjugated with AlexaFluor dyes. Nuclei were
counterstained with DAPI. Signals were captured with 90i microscope (Nikon). The
paraffin-embedded specimens for normal human livers (n=2) and alcohol-induced
fibrosis (n=2) at the Harbor-UCLA Medical Center or biliary atresia (n=3) at Children’s
Hospital Los Angeles were used for immunohistochemistry under study protocols
approved by the institutional review boards (HS-11-00476, CCI-10-00148). The primary
antibodies were detected with SuperPicure HRP Polymer (Life Technologies).

Cell isolation. Nonparenchymal cells (NPCs) were isolated by the NPC Core in
University of Southern California. Mouse livers were perfused through the superior vena
cava by 0.5% pronase (Roche) and 0.044% collagenase (Sigma). After agitation of the
digested tissue with 10 g/ml DNase, the cells were placed on the top of four OptiPrep
61
gradients (1.034, 1.043, 1.058, 1.085) in Beckman ultracentrifuge tubes and were
centrifuged in the SW-41Ti rotor at 20,000 rpm for 15 minutes. The 1.058 fraction was
used as NPCs.

FACS. NPCs were incubated with the anti-GPM6A antibody (MBL) at 1,500-fold
dilution for 30 minutes. After washing, the primary antibody was detected with anti-rat
IgG AlexaFluor647 (Life Technologies). After excluding propidium iodide (PI)+ dead
cells, PI- live cells were analyzed with a krypton laser and a 424nm filter to detect VitA
autofluorescence with FACS Aria I (BD Bioscience) in the USC Flow Cytometry Core
(Li et al., 2013; Lua et al., 2014). The VitA- fraction was further separated based on the
signal intensities for GPM6A and GFP. MCs were isolated from the liver surface as
previously described in Chapter 2. After whole liver digestion with 1 mg/ml pronase, the
cells were incubated with the anti-GPM6A antibody. The primary antibody was detected
with anti-rat IgG AlexaFluor568 antibodies and VitA-GPM6A+ MCs were sorted by
FACS.

Cell culture and immunocytochemistry. After FACS, the cells (2 x 10
4
cells) were plated
on collagen-coated 24-well plates in DMEM containing 10% FBS. The cells were treated
with 10 ng/ml TGF- 1 (Sigma) or 100 pg/ml platelet-derived growth factor (PDGF)-BB
(eBioscience). For immunocytochemistry, cells were cultured on a glass cover and fixed
with 4% paraformaldehyde. After blocking, the sections were incubated with primary
62
antibodies. The primary antibodies are listed in Table 1. The primary antibodies were
detected with secondary antibodies conjugated with AlexaFluor568.

QPCR. QPCR method was the same as described in Chapter 2. Primer sequences are
listed in Table 2.

Microarray analysis. After FACS sorting, total RNA was extracted, and the microarray
probes were synthesized using the Ovation RNA amplification system (Nugen) as
previously described (Asahina et al., 2009). GEO accession number for microarray data:
GSE66788.

Statistical analysis. Statistical significance was assessed by ANOVA followed by post-
hoc Tukey HSD test among multiple samples or Student’s t-test between two samples. A
P value of less than 0.05 was considered statistically significant.

63
3.3 Results
Broad Expression of GFP in the Col1a1
GFP
Mouse Liver
Immunohistochemistry showed that the normal Col1a1
GFP
liver expresses GFP in some
DES+ HSCs in the sinusoid, as well as in the PFs adjacent to the bile duct and SMCs in
the hepatic artery and portal vein (Fig. 9A). GFP expression was also observed in
possible DES+ SLCs around the central vein as well as in DES+ CFs beneath MCs (Fig.
9B,C). GFP expression was observed in MCs expressing GPM6A which is an MC
marker (Fig. 9D). No GPM6A expression was observed in the portal area (Fig. 9E).
PDPN was expressed in MCs, bile duct, and LYVE1+ lymphatic vessels adjacent to the
portal vein (Fig. 9F,G). THY1 expression was evident in PFs, lymphatic vessels, and
some MCs (Fig. 9H,I).

64
Figure 9

Figure 9. Expression of GFP in HSCs, PFs, SMCs, SLCs, CFs, and MCs in Col1a1
GFP

mouse livers. Expression of GFP and different cell markers was examined by
immunohistochemistry in the Col1a1
GFP
(A-F,H) or wild-type (G,I) mouse livers. (A)
GFP expression is observed in DES+ HSCs (arrows), PFs (arrowheads) adjacent to bile
duct (bd), and SMCs (arrowheads) in the hepatic artery (ha) and portal vein (pv).
Asterisks indicate lymphatic vessels (lv). (B) GFP expression is observed in DES+ SLCs
(arrowheads) in the central vein (cv). (C) MCs (double arrowheads) and CFs (double
arrows) express GFP. Arrows indicate GFP+ HSCs. (D) Both GPM6A+ MCs (double
arrowheads) and GPM6A- CFs (double arrows) express GFP. (E) No GPM6A expression
65
in the portal triad. (F) PDPN+ MCs express GFP (double arrowheads). (G) Bile duct and
lymphatic vessels (asterisks) are positive for PDPN. (H) THY1+ PFs express GFP
(arrowheads). (I) Expression of THY1 in lymphatic vessels (asterisks). Some MCs
express THY1 (double arrowheads). Scale bar: 10 m.

66
Separating HSCs with FACS
HSCs store VitA as retinyl ester (Geerts, 2001). VitA shows autofluorescence upon
exposure to UV light. Therefore, VitA+ HSCs may be purified based on the
autofluorescence of VitA storing in their cytoplasm. Using FACS, we attempted to isolate
HSCs and PFs based on the autofluorescence of VitA in HSCs and GFP expression.
NPCs prepared from the Col1a1
GFP
mouse showed VitA+ (52.3±14.1%) and VitA-GFP+
(6.7±3.0%) populations (Fig. 10A). To characterize these populations, we surveyed gene
expression using microarray and found that VitA+ HSCs highly express Reelin (Reln)
(Table 3). We validated the high expression of Reln mRNA in VitA+ HSCs by QPCR
(Fig. 10B). The VitA+ HSCs also expressed Des and Gfap, general markers for HSCs
(Fig. 10B). Immunohistochemistry showed a specific expression of RELN in DES+
HSCs in the sinusoid (Fig. 10C). In the portal triad, lymphatic vessels were also positive
for RELN (Fig. 10C,D). In the Col1a1
GFP
livers, RELN expression was observed in
GFP+ HSCs, but not in MCs, CFs, and SLCs (Fig. 10E). These data indicate that RELN
is a specific marker for HSCs and lymphatic vessels in the liver.

67
Figure 10

Figure 10. Separation of VitA+ HSCs and VitA-GFP+ population from Col1a1
GFP
livers.
(A) VitA+ and VitA-GFP+ populations were sorted from Col1a1
GFP
NPCs by FACS
(n=3). Wild-type NPCs were used as negative controls. (B) QPCR of NPCs before FACS
(All), VitA+ HSCs (V+), and VitA-GFP+ (V-G+) population separated from the
Col1a1
GFP
liver. *P<0.05, **P<0.01. (C-G) Immunohistochemistry of the wild-type
(C,D) and Col1a1
GFP
(E-G) mouse livers. (C) RELN is expressed in HSCs (arrows) and
lymphatic vessels (asterisks), but not in PFs (arrowheads) around the bile duct (bd) and
68
portal vein (pv). (D) RELN is expressed in LYVE1+ lymphatic vessels (asterisks). (E)
No RELN expression in MCs (double arrowheads), CFs (double arrows), and SLCs (an
arrowhead) adjacent to the central vein (cv). (F,G) GFP+ PFs (arrowheads) express ELN
and ENTPD2. Scale bar: 10 m.

69
Table 3. Summary of microarray analysis.

70
The Presence of PFs and MCs in the VitA-GFP+ Population
Microarray analysis and QPCR showed that the VitA-GFP+ population highly expressed
PF markers including Eln and Entpd2 (Table 3, Fig. 10B). We confirmed expression of
ELN and ENTPD2 in the GFP+ PFs of the Col1a1
GFP
livers (Fig. 10F,G).
We also found that the VitA-GFP+ population expresses MC markers including
Gpm6a, Msln, Pdpn, and Upk1b (Table 3, Fig. 10B). Iwaisako et al. reported the
separation of a similar VitA-GFP+ population from the Col1a1
GFP
mouse and identified
MSLN as a PF marker (Iwaisako et al., 2014). Given that both MCs and PFs express GFP
(Fig. 9A,C), we assumed that both MCs and PFs are enriched in the VitA-GFP+
population according to FACS. To test our assumption, we reevaluated the expression of
MC markers in mouse livers. We isolated VitA-GPM6A+ MCs by FACS (Fig. 11A).
Purified MCs highly expressed Gpm6a mRNA (Fig. 11B). Microarray analysis of VitA-
GPM6A+ MCs revealed high expression of Col1a1, Gpm6a, Krt19, Msln, Pdpn, and
Upk1b mRNAs (Table 3). KRT19 expression in MCs and bile duct was confirmed by
immunohistochemistry (Fig. 11C). MSLN expression was exclusively observed in MCs
but not in PFs (Fig. 11D). Using Upk1b
RFP/+
mice, we confirmed exclusive expression of
RFP in MCs (Fig. 11E,F). Based on these data, we concluded that PFs and MCs are
enriched in the VitA-GFP+ population.
In normal human livers, MSLN was exclusively expressed in MCs and no MSLN
expression was observed in the portal area (Fig. 11G). Fibrotic livers caused by biliary
atresia or chronic alcohol abuse showed the accumulation of ACTA2+ myofibroblasts in
71
the portal area and these myofibroblasts were negative for MSLN (Fig. 11H,I). We
concluded that MSLN is a marker for MCs in both mouse and human livers.

72
Figure 11

Figure 11. Expression of MC markers in mouse and human livers. (A) Isolation of MCs
from mouse liver with FACS using anti-GPM6A antibodies. VitA-GPM6A+ MCs were
sorted for QPCR and microarray analysis (n=3). (B) QPCR of Gpm6a mRNA in liver
cells before FACS and VitA-GPM6A+ MCs. **P<0.01. (C-F) Immunohistochemistry of
wild-type (C,D,F) and Upk1b
RFP/+
(E) livers. (C) MCs (double arrowheads) and bile duct
(bd) express KRT19. (D) MSLN is expressed in the MCs (double arrowheads) but not in
the PFs. Asterisks indicate lymphatic vessels. (E) Upk1b
RFP/+
liver shows RFP expression
in the MCs (double arrowheads). (F) No RFP expression in the control Upk1b
+/+
liver.
73
(G-I) Immunohistochemistry of human liver specimens. (G) MSLN is exclusively
expressed in MCs (arrowheads) in normal livers. (H) Biliary atresia specimens show
ACTA2 expression in the portal area. Myofibroblasts are negative for MSLN. (I)
ACTA2+ myofibroblasts in alcohol-induced fibrosis are negative for MSLN. Scale bar:
10 m.

74
Enrichment of PFs by FACS
To enrich PFs from the heterogeneous VitA-GFP+ population, we subtracted GPM6A+
MCs and enriched PFs as VitA-GFP+GPM6A- cells. PI- live cells were separated into
VitA+ HSCs (P1, 6.3±2.0%, n=8) and VitA- population, and then the VitA- population
was further separated into P2 (52.2±12.3%), P3 (1.1±0.3%), and P4 (0.3±0.2%)
populations based on the signal intensities for GPM6A and GFP (Fig. 12A). QPCR
confirmed enrichment of HSCs in the VitA+ P1 population that expresses Des, Gfap,
Lrat, and Reln mRNAs but not markers for PFs (Eln, Entpd2) and MCs (Gpm6a, Msln,
Upk1b) (Fig. 12B). The P2 population expressed markers for hepatocytes (Alb) and
cholangiocytes (Epcam, Krt19), without enriching other cell markers (Fig. 12B). The P3
population expressed Des, Eln, and Entpd2 but not HSC markers or MC markers (Fig.
12B), suggesting the enrichment of PFs. In addition, high Acta2 mRNA expression in the
P3 population suggests that SMCs are also fractionated in this population. The P4
population expressed markers for MCs including Krt19 (Fig. 12B).

75
Figure 12

Figure 12. Separation of HSCs, PFs, and MCs from the Col1a1
GFP
livers by FACS. (A)
NPCs prepared from wild-type or Col1a1
GFP
livers were stained with PI and anti-GPM6A
antibodies. PI- cells were analyzed for autofluorescence of VitA and VitA+ HSCs were
sorted (P1). The VitA- fraction was further analyzed for the expression of GPM6A and
GFP into P2-P4 factions. Wild-type or Col1a1
GFP
NPCs incubated with the isotype IgG
were used as negative controls. (B) QPCR of cells before FACS (All) and P1-P4 fractions
obtained by FACS. *P<0.05, **P<0.01.

76
Myofibroblastic Conversion of HSCs, PFs, and MCs
After sorting P1-HSCs, P3-PFs, and P4-MCs from normal Col1a1
GFP
livers by FACS, we
analyzed the phenotypic changes of these cells in culture. After 3 days in culture, HSCs
showed autofluorescence of VitA and dendritic processes (Fig. 13A). PFs showed
fibroblastic morphology, without VitA autofluorescence. MCs exhibited a round shape
and did not show VitA autofluorescence. Immunocytochemistry showed the expression
of DES and RELN in the HSCs, but not in the MCs (Fig. 13B). The PFs expressed DES,
ELN, and ENTPD2 (Fig. 13B). Only the MCs expressed GPM6A and MSLN (Fig. 13B).
ELN was weakly expressed in MCs (Fig. 13B). We confirmed that ENTPD2+ PFs are
negative for MSLN in cultured NPCs (Fig. 13C).
QPCR showed that these cells expressed Tgfbr1 and Tgfbr2 mRNAs (Fig. 13D).
MCs did not express Pdgfra and Pdgfrb mRNAs (Fig. 13D). After treatment with TGF-
1, HSCs, PFs, and MCs increased the expression of Acta2 and Col1a1 mRNAs (Fig.
13E). Interestingly, TGF- 1 strongly suppressed Cyclin D1 (Ccnd1) mRNA in the PFs
but not in the HSCs and MCs (Fig. 13E). PDGF-BB induced Ccnd1 mRNA only in the
HSCs (Fig. 13E). TGF- 1 induced the nuclear localization of P-SMAD3, a downstream
effector of TGF-  signaling, in HSCs, PFs, and MCs (Fig. 13F). These cells
differentiated into ACTA2+ myofibroblasts by TGF- 1 (Fig. 13F). These data indicate
that even though HSCs, PFs, and MCs have the differentiation potential to
myofibroblasts, their proliferation is differently regulated by TGF- 1 and PDGF-BB.

77
Figure 13

Figure 13. Differential responses of HSCs, PFs, and MCs against TGF- 1 and PDGF-
BB. (A) Culture of HSCs, PFs, and MCs purified from Col1a1
GFP
livers by FACS. (B)
Immunocytochemistry of HSCs, PFs, and MCs on day-3 in culture. (C)
Immunocytochemistry of NPCs for ENTPD2 and MSLN 24 hours after plating. (D)
QPCR of the HSCs, PFs, and MCs. *P<0.05; **P<0.01. (E) QPCR of HSCs, PFs, and
MCs treated with either TGF- 1 or PDGF-BB. (F) Immunocytochemistry of P-SMAD3
78
or ACTA2 in cultured HSCs, PFs, and MCs treated with TGF- 1 for 3 hours (P-SMAD3)
or for 3 days (ACTA2). Scale bar: 10 m.

79
Expression of HSC, PF and MC Markers in BDL-Induced Biliary Fibrosis
Next, we analyzed phenotypic changes of HSCs, PFs, and MCs in biliary fibrosis induced
by BDL for 3 weeks in Col1a1
GFP
mice. BDL surgery is known to induce myofibroblasts
around the portal vein. Expression of ENTPD2 was restricted in GFP+ myofibroblasts
around the bile duct but not in HSCs in the sinusoid in biliary fibrosis (Fig. 14A).
Although expression of ACTA2 was observed in PFs adjacent to the bile duct, not all PFs
became positive for ACTA2 (Fig. 14B). The expression of ACTA2 was also observed in
some GFP+ HSCs in the injured parenchyma (Fig. 14C). GFP+ MCs expressed KRT19
but not EPCAM on the liver surface (Fig. 14D,E). KRT19+ or EPCAM+ biliary
epithelial cells were negative for GFP in the large bile duct (Fig. 14F,G). Unexpectedly, a
few biliary epithelial cells showed the GFP expression in the small bile duct in biliary
fibrosis (Fig. 14H,I). Deposition of ELN was observed around the bile duct and beneath
the MCs (Fig. 14J). RELN was expressed in HSCs in the sinusoid (Fig. 14K). Similar to
the normal livers, MSLN was expressed in MCs but not in PFs around the portal vein
after BDL (Fig. 14L).

80
Figure 14

Figure 14. Expression of markers in biliary fibrosis induced by BDL. Col1a1
GFP
(A-K)
and wild-type (L) mouse livers were analyzed by immunohistochemistry. (A) ENTPD2 is
expressed in GFP+ myofibroblasts around the bile duct (bd, arrowheads). (B) ACTA2 is
expressed in GFP+ PFs (arrowheads) adjacent to the bile duct. Not all GFP+ PFs express
ACTA2. (C) ACTA2 expression in GFP+ HSCs (arrows). (D,E) GFP+ MCs express
KRT19 but not EPCAM (double arrowheads). (F-I) A few KRT19+ or EPCAM+ biliary
epithelial cells in the small bile duct express GFP (double arrows). (J) The deposition of
ELN is observed around the bile duct (arrowheads) and beneath the GFP+ MCs (double
arrowheads). (K) RELN is expressed in GFP+ HSCs in the sinusoid (arrows). (L) MSLN
81
is exclusively expressed in PDPN+ MCs (double arrowheads) but not in PFs around the
portal vein (pv). Scale bar: 10 m.

82
Expression of HSC, PF and MC Markers in DDC Diet-Induced Biliary Fibrosis
We also characterized the expression of markers in DDC diet-induced biliary fibrosis.
DDC diet induced expansion of GFP+ cells in the portal triad and some PFs adjacent to
the bile duct expressed ACTA2 (Fig. 15A). ENTPD2 was weakly expressed in PFs (Fig.
15B). Similar to the BDL model, GFP expression was observed in biliary epithelial cells
in small bile ducts (Fig. 15C-E). GPM6A and MSLN were exclusively expressed in MCs
(Fig. 15F-H).

83
Figure 15

Figure 15. Expression of markers in biliary fibrosis induced by the DDC diet. Col1a1
GFP

mice were fed DDC diet for 1 month and the livers were analyzed by
immunohistochemistry for GFP (green) and ACTA2 (A, red), ENTPD2 (B), KRT19
(C,D), EPCAM (E), GPM6A (F), or MSLN (G,H). (A) ACTA2 is expressed in GFP+
myofibroblasts (arrowheads) near the portal vein (pv). Not all GFP+ myofibroblasts
express ACTA2. (B) ENTPD2 is expressed in GFP+ myofibroblasts around the expanded
bile duct (bd, arrowheads). No ENTPD2 expression in GFP+ HSCs in the sinusoid
(arrows). (C) KRT19 expression in the bile duct. (D,E) A few KRT19+ or EPCAM+
biliary epithelial cells express GFP in the small bile duct (double arrows). (F) GPM6A+
MCs express GFP (double arrowheads). An arrow indicates HSCs. (G,H) MSLN is
84
exclusively expressed in GFP+ MCs (double arrowheads). No MSLN expression in
GFP+ HSCs (arrow) and myofibroblasts around the bile duct. Scale bar: 10 m.

85
Expression of HSC, PF and MC Markers in CCl
4
-Induced Liver Fibrosis
Next, we induced liver fibrosis by injections of CCl
4
, by which HSCs in the sinusoid
differentiate into myofibroblasts around the central vein. Different from the BDL and
DDC models, the CCl
4
model does not induce biliary fibrosis. After CCl
4
injections,
accumulation of ACTA2+ myofibroblasts was observed around the central vein and
beneath the mesothelium (Fig. 16A,B). ENTPD2 and ELN were expressed in PFs that are
negative for ACTA2 (Fig. 16C,D). The expression of RELN was observed in GFP+DES+
HSCs and its expression was not evident in MCs and myofibroblasts near the liver
surface (Fig. 16E,F). Differing from the BDL and DDC models, CCl
4
did not induce GFP
expression in biliary epithelial cells (Fig. 16G,H). MCs expressed GPM6A, PDPN,
CD200, and MSLN (Fig. 16I-L). No MSLN expression was observed in the portal triad.

86
Figure 16

Figure 16. Expression of markers in CCl
4
-induced liver fibrosis. Immunohistochemistry
of markers in the livers from Col1a1
GFP
(A,E,G,H,K) or wild-type (B-D,F,I,J,L) mice
treated with CCl
4
. (A) ACTA2+ myofibroblasts near the central vein (cv) express GFP
(arrows). (B) Double arrows indicate ACTA2+ myofibroblasts beneath MCs (double
arrowheads). DES+ PFs adjacent to the bile duct (bd) are negative for ACTA2
(arrowheads). ha; hepatic artery, lv; lymphatic vessel, pv; portal vein. (C,D) ACTA2- PFs
express ENTPD2 and ELN (arrowheads). (E,F) RELN is expressed in DES+ HSCs
87
(arrows), but not in MCs (double arrowhead) and DES+ mesenchymal cells beneath MCs
(double arrows). (G,H) No GFP expression in biliary epithelial cells (arrowheads). (I,J)
MCs express GPM6A, PDPN, and CD200 (double arrowheads). PDPN is also expressed
in bile duct and lymphatic vessels (asterisk). Endothelial cells in the portal vein and
lymphatic vessels express CD200. (K,L) MCs express MSLN, PDPN, and GFP (double
arrowheads). No MSLN expression is observed in the portal triad. Scale bar: 10 m.

88
Isolation of HSCs, PFs, and MCs from Fibrotic Livers
To characterize HSCs, PFs, and MCs in fibrosis induced by BDL, DDC diet, or CCl
4

injections, NPCs prepared from Col1a1
GFP
livers were analyzed by FACS (Fig. 17 A-D).
We sorted VitA+ HSCs (P1, 11.6±4.3%, n=4), VitA- GFP+GPM6A- PFs (P3, 1.7±0.3%),
and VitA-GFP+GPM6A+ MCs (P4, 0.3±0.1%) from the BDL model (Fig. 17B). We also
observed P5-GFP
Dim
cells between P2 and P3 after BDL (Fig. 17B). QPCR revealed that
P5 population expresses Epcam and Krt19 mRNAs (Fig. 17E), indicating the enrichment
of GFP+ biliary epithelial cells in P5. In contrast, the P3-PFs showed less expression of
Epcam and Krt19 and did not express Reln, Gpm6a, and Msln (Fig. 17E). P3-PFs
increased expression of Acta2 after BDL (Fig. 17E). Although PFs expressed more
Col1a1 mRNA than HSCs, PFs slightly decreased Col1a1 by BDL (Fig. 17E). PFs
decreased the expression of Entpd2. After BDL, P1-HSCs decreased the expression of
Reln, while increasing Acta2 (Fig. 17E). P4-MCs did not increase the expression of Acta2
and Col1a1 by BDL (Fig. 17E). MCs kept expressing Gpm6a, Msln, and Timp1 mRNAs.
We also analyzed the gene expression of these cells by microarray between the sham and
BDL samples and confirmed the similar expression patterns (Table 3). These results
suggest that BDL moderately induces activation of both HSCs and PFs in mouse livers.
We also isolated P1-P5 populations from fibrotic Col1a1
GFP
livers induced by
DDC or CCl
4
(Fig. 17C,D). Similar to the BDL model, P1-HSCs increased the expression
of Col1a1 and Timp1 in the DDC model (Fig. 17E). HSCs reduced the expression of Reln
(Fig. 17E). P3-PFs decreased the expression of Acta2 mRNA compared to the BDL
model, implying that the DDC model does not fully induce myofibroblastic conversion of
89
PFs. P4-MCs kept expressing Gpm6a, Msln, Krt19, and Timp1. Similar to the BDL
model, DDC diet induced GFP expression in P5-biliary epithelial cells (Fig. 17C). In the
CCl
4
model, GFP+ biliary epithelial cells were few in P5 (Fig. 17D). As expected, CCl
4

induced the activation markers in P1-HSCs (Fig. 17E). P3-PFs did not up-regulate the
expression of Acta2 and Col1a1 by CCl
4
.
Based on the FACS data, we estimated the relative contribution of HSCs, PFs,
and MCs to GFP+ myofibroblasts in these models. Our method did not allow detecting
GFP- PFs. In addition, liver injury caused by different etiology changed the number of
blood cells in the NPC fraction. Thus, we quantified the ratio of GFP+ HSCs, PFs, and
MCs against VitA+ HSCs in the NPC fraction (Fig. 17A-D). As expected, CCl
4
treatment
increased the percentage of GFP+ HSCs in VitA+ HSCs (74.5±4.0%) compared to the
control (Fig. 17F). Interestingly, BDL and DDC models also increased the percentage of
GFP+ HSCs (52.7±14.2% in BDL and 49.3±9.7% in DDC), indicating activation of
HSCs in biliary fibrosis (Fig. 17F). In the control liver, the ratio of GFP+ PFs against
VitA+ HSCs was 7.5±1.8% and this ratio was increased to 13.9±3.8% by BDL (Fig.
17G). These results suggest that PFs partly contribute to GFP+ myofibroblasts in biliary
fibrosis induced by BDL. According to the increase of GFP+ HSCs in the CCl
4
model,
the percentage of GFP+ P3-PFs decreased (Fig. 17F,G). DDC-induced fibrosis did not
increase the percentage of GFP+ P3-PFs (Fig. 17G). GFP+ P4-MCs occupied only 0.9-
2.9% all groups (Fig. 17H).

90
Figure 17

Figure 17. Isolation of HSCs, PFs, and MCs from Col1a1
GFP
fibrotic livers. (A-D) NPCs
were prepared from control (A, n=6) or injured Col1a1
GFP
livers induced by BDL (B,
n=6), DDC (C, n=3), and CCl
4
(D, n=4). PI- cells were analyzed for autofluorescence of
VitA and P1 VitA+ HSCs were sorted. The VitA- fraction was further analyzed for the
expression of GPM6A and GFP. P1-P5 factions were sorted for further analysis. (E)
QPCR of P1-P5 fractions. (F-H) Percentages of GFP+ HSCs (F), PFs (G), and MCs (H)
in VitA+ HSCs in different models. *P<0.05, **P<0.01.
91
3.4 Discussion
During liver fibrosis, myofibroblasts actively synthesize collagen and participate in liver
fibrosis. Depending on the etiology, genesis of ACTA2+ myofibroblasts varies, e.g.,
myofibroblasts appear around the central vein in the CCl
4
model, whereas around the bile
duct in biliary fibrosis induced by BDL and DDC models. Although HSCs seem to be the
major source of myofibroblasts in liver fibrosis (Mederacke et al., 2013), PFs are
suggested to be another source in biliary fibrosis (Dranoff et al., 2002; Uchio et al., 2002;
Wells et al., 2004; Li et al., 2007; Lemoinne et al., 2015). In addition, MCs were shown
to differentiate into myofibroblasts (Li et al., 2013). To understand the contribution of
different cell types to myofibroblasts and their roles, we isolated HSCs, PFs, and MCs
from normal or injured livers induced by different insults by FACS. We used the
Col1a1
GFP
transgenic mice to detect activated HSCs, PFs, and MCs. In the present study,
we separated GFP+ cells in Col1a1
GFP
livers and estimated the contribution of HSCs,
PFs, and MCs to GFP+ myofibroblasts in different models.
The Col1a1
GFP
transgenic mice have been used to detect and isolate activated
GFP+ HSCs (Yata et al., 2003; D'Ambrosio et al., 2011; Kisseleva et al., 2012; Yang et
al., 2012; Iwaisako et al., 2014). We found that the normal Col1a1
GFP
liver broadly
expresses the GFP in some HSCs, PFs, and MCs. Based on the VitA storage and
expression of GFP and GPM6A, we separated VitA+ HSCs, VitA-GFP+GPM6A- PFs,
and VitA-GFP+GPM6A+ MCs by FACS. We found that HSCs and lymphatic vessels
express RELN, which was previously reported in rat liver (Kobold et al., 2002; Samama
and Boehm, 2005). No RELN was observed in PFs and MCs, indicating that RELN is a
92
good marker to distinguish HSCs and lymphatic vessels (Fig. 18). PFs expressed ELN
and ENTPD2 in the portal triad (Fig. 18). Li et al. reported that rat PFs express ELN but
not DES (Li et al., 2007). In contrast, we found the expression of DES in mouse PFs.
ELN was expressed in PFs and MCs in biliary fibrosis. ENTPD2 was originally identified
as a PF marker in rat livers, and its expression was shown to decrease in PFs and increase
in HSCs in culture activation (Li et al., 2007). Our data showed that Entpd2 mRNA is
down-regulated in PFs in fibrosis induced by different etiology. Differing from rat livers,
Entpd2 mRNA was not induced in activated mouse HSCs.
After BDL, we observed expansion of GFP+ PFs in the portal triad of the
Col1a1
GFP
mouse. Although ENTPD2+GFP+ PF-derived myofibroblasts were observed
around the portal vein, co-expression of ACTA2, a general myofibroblast marker, was
restricted near the bile duct, implying that activation state of PFs is heterogeneous. After
sorting PFs from the control and BDL models, P3-PFs increased Acta2 mRNA
moderately. GFP+ PFs in the normal liver expressed high Col1a1 mRNA compared to
HSCs. However, GFP+ PFs isolated from the BDL model did not further increase Col1a1
mRNA. Thus, our data indicate that PFs actively express Col1a1 mRNA in the normal
liver and they expand in biliary fibrosis with limited conversion to ACTA2+
myofibroblasts. Based on the FACS data, we estimated that the ratio of GFP+ PFs against
VitA+ HSCs is increased from 8% in the control to 14% in the BDL model. GFP+ HSCs
were increased from 11% to 53% by BDL. Compared to the BDL model, DDC diet did
not increase Acta2 mRNA in P3-PFs. The presence of GFP+ PFs was low in the DDC
model compared to the BDL model. These data indicate that although PFs contribute to
93
GFP+ myofibroblasts in the portal area in biliary fibrosis caused by BDL or DDC diet,
the contribution of HSCs is still dominant (Fig. 18). In agreement with our results,
Mederacke et al. reported that HSCs, but not PFs, are a major source of myofibroblasts in
biliary fibrosis using Lrat
Cre
mice (Mederacke et al., 2013). As expected, CCl
4
treatment
extensively increased the ratio of GFP+ HSCs in VitA+ HSCs to 75% and the ratio of
GFP+ PFs was only 2%. Our quantification data suggest that HSCs are the major source
of GFP+ myofibroblasts in liver fibrosis induced by CCl
4
injections. PFs partly contribute
to myofibroblasts in the BDL model (Fig. 18).
Our conclusion contradicts the results from the recent paper in which the authors
separated GFP+VitA- cells as PFs from the same Col1a1
GFP
mouse model and identified
MSLN as a PF marker (Iwaisako et al., 2014). Based on the increased number of
GFP+VitA- cells in the BDL model, they estimated that PFs are the major source of
myofibroblasts. In the present study, we found that BDL or DDC diet induces de novo
GFP expression in the small bile duct of the Col1a1
GFP
mouse. Thus, the presence of
GFP+ biliary epithelial cells in GFP+VitA- population leads to overestimation of the
contribution of GFP+ PFs in biliary fibrosis in their study. Furthermore, our data indicate
that the GFP+VitA- population also contains MCs. Immunohistochemistry showed that
MSLN is expressed in MCs, but not in PFs in mouse and human livers. During the
preparation of NPCs from mouse livers, perfused livers were further digested with
collagenase, and MCs appeared to be collected in the NPC fraction. In fact, FACS
revealed the presence of GPM6A+ MCs in the NPC fraction. Although we attempted to
94
trace Msln+ cells using Msln
CreERT2-IRES-lacZ
;R26TG
fl/fl
mice (Rinkevich et al., 2012), they
did not specifically label MCs in the liver with tamoxifen treatment (data not shown).
In culture, PFs showed fibroblastic morphology without storing VitA lipids, and
these characteristics differed from those in HSCs and MCs. After TGF- 1 treatment, the
PFs differentiated into myofibroblasts expressing ACTA2, similar to the activation of
HSCs. Interestingly, TGF- 1 suppressed Ccnd1 mRNA in PFs but not in HSCs, similar
to the previous report of rat livers (Wells et al., 2004). These data indicate that HSCs and
PFs have the potential to differentiate into myofibroblasts and that their proliferation is
differently regulated by TGF-  and PDGF-BB.
We previously reported that MCs migrate inward from the liver surface and give
rise to HSCs in the BDL model or to myofibroblasts in CCl
4
-induced fibrosis (Li et al.,
2013). MCs contribute to 2% of myofibroblasts in the CCl
4
model near the liver surface
(Li et al., 2015). Upon differentiation to HSCs or myofibroblasts, MCs lose the
expression of GPM6A (Li et al., 2013). In the present study, we isolated MCs based on
the expression of GPM6A from livers and this method was unable to isolate MC-derived
HSCs from the BDL model. This might be a reason isolated MCs do not show increased
expression of Acta2 and Col1a1 mRNAs in injured livers. As we have previously
reported (Li et al., 2013), cultured MCs induce ACTA2+ myofibroblasts by TGF- 1.
Differing from HSCs and PFs, MCs do not express Pdgfra and Pdgfrb mRNAs and do
not respond to PDGF-BB, indicating unique feature of MCs. Microarray analysis
revealed that MCs highly express Upk1b and Upk3b (Table 3). UPK complex is
composed of UPK1B/UPIII and UPK1A/UPII pairs and functions as a permeability
95
barrier of the urothelium (Rudat et al., 2014). Upk1b
RFP/+
heterozygous mice showed
specific RFP expression only in MCs in the liver. Although we produced Upk1b
RFP/RFP

knockout mice, they were born without noticeable abnormalities in the liver or bladder
(data not shown). Similarly, Upk3b knockout mice do not show any abnormalities in the
mesothelium (Rudat et al., 2014), indicating that UPK is not essential for the
development of MCs.
According to immunohistochemistry of the Col1a1
GFP
livers, CFs, SLCs, and
SMCs are also expected to appear in the VitA-GFP+GPM6A- population when using
FACS. Differing from PFs, ENTPD2 expression was not observed in the Glisson’s
capsule on the liver surface, suggesting that there may be a phenotypic difference
between PFs and CFs. The markers we identified will be useful for creating specific Cre
lines for further study on the cell lineages of different mesenchymal cells in liver fibrosis.

96
Figure 18

Figure 18. Summary of the contribution of HSCs, PFs, and MCs in liver fibrosis. HSCs
show autofluorescence of VitA with dendritic processes and express RELN. PFs acquire
fibroblast like morphology and express ENTPD2. MCs on the liver surface appear as
epithelial sheet and express GPM6A. HSCs, PFs, and MCs differentiate into ACTA2+
myofibroblasts upon liver injury. FACS analysis revealed that HSCs are the dominant
source of myofibroblasts in the BDL, DDC or CCl
4
-injured liver injury. MCs and PFs
partly contributed to myofibroblasts in capsular fibrosis beneath the liver surface and
biliary fibrosis around the portal vein.

97
Chapter 4
No transdifferentiation of mesenchymal cells to epithelial cells in
liver fibrosis
4.1 Introduction
As described in Chapter 2 and 3, our data indicate that different types of mesenchymal
cells including MCs, PFs, and HSCs give rise to myofibroblasts in liver injury. Among
these mesenchymal cells, HSCs are the major source of myofibroblasts in liver fibrosis
irrespective of different etiology. In normal adult liver, quiescent HSCs are characterized
by storage of VitA lipids and the presence of substantial dendrite-like processes along the
sinusoid. When liver injury occurs, HSCs express ACTA2, lose VitA lipids, and acquire
a myofibroblastic phenotype. In normal condition, HSCs express not only mesenchymal
cell markers such as DES, but also neural cell markers, including GFAP, nestin, and p75
neurotrophin receptor (p75NTR) (Geerts, 2001). Based on the intermediate phenotype of
mesenchymal and neural cells, the neural crest was suggested to be the origin of HSCs.
Nevertheless, a cell lineage tracing using Wnt1
Cre
and Rosa26 reporter mice refuted this
hypothesis (Cassiman et al., 2006). Using MesP1
Cre
and Rosa26lacZ
flox
mice, Asahina et
al. demonstrated that MesP1+ mesoderm gives rise to MCs, which differentiate into
HSCs and PFs during liver development (Asahina et al., 2009). They did not observe the
contribution of MesP1+ mesoderm to liver epithelial cells such as hepatocytes and
cholangiocytes, which arise from definitive endoderm. However, several in vivo and in
vitro studies reported that HSCs differentiate into epithelia cell lineage, including
hepatocytes, cholangiocytes, and progenitor cell types known as oval cells, thereby acting
98
as stem cells in the liver (Yang et al., 2008; Michelotti et al., 2013). Oval cells are
epithelial cells with ovoid nuclei and minimal cytoplasm that express cholangiocyte
markers such as cytokeratin 19 (KRT19), SOX9, EPCAM, and osteopontin (Okabe et al.,
2009; Carpentier et al., 2011; Espanol-Suner et al., 2012). A morphological study
suggested that oval cells arise from the Canals of Hering, a connection between
hepatocytes and cholangiocytes near the portal triad, and give rise to both cell types
during regeneration in a manner comparable to the differentiation pathway of embryonic
hepatoblasts (Paku et al., 2001).
To test whether HSCs give rise to epithelial cells in adult liver, we determined the
hepatic mesenchymal cell lineages using MesP1
Cre
and Rosa26mTmG
flox
mice. Genetic
cell lineage tracing revealed that the MesP1+ mesoderm gives rise to MCs, PFs and
HSCs, but not to hepatocytes or cholangiocytes, in the adult liver. Upon CCl
4
or BDL
surgery-mediated liver injury, mesodermal mesenchymal cells, including MCs, PFs and
HSCs differentiate into myofibroblasts but not into hepatocytes or cholangiocytes.
Furthermore, differentiation of the mesodermal mesenchymal cells into oval cells was not
observed. These results indicate that HSCs are not sufficiently multipotent to produce
hepatocytes, cholangiocytes, or oval cells via mesenchymal-epithelial transition in vivo.

99
4.2 Materials
Mice. MesP1
Cre
, Rosa26mTmG
flox
(R26TG
fl
) mice were described previously in Chapter
2. Fibrosis was induced injection of CCl
4
30 times (Li et al., 2013). To induce biliary
fibrosis, the mice were subjected to BDL for 2 weeks (n=3) (Lua et al., 2016). For oval
cell induction, 5 mice were fed 0.1% DDC chow and were sacrificed 3 mice at 4 weeks
(Preisegger et al., 1999). The remaining 2 mice were additionally fed normal chow for 4
weeks. MesP1
+/+
; R26TG
fl/fl
mice were used for negative controls. The mice were
handled in accordance with protocols approved by the Institutional Animal Care and Use
Committee of the University of Southern California.

Immunohistochemistry and Quantification. Immunohistochemistry method was
described in Chapter 2. The antibodies used in immunostaining are listed in Table 1. To
quantify GFP expression, at least 3 sections were immunostained for each mouse. Digital
images were captured using a 40x objective (at least 15 images per animal) and more
than 1,000 cells were counted for each antigen.

Isolation and culture HSCs. Isolation method was described previously in Chapter 3. In
brief, a pure HSC fraction was collected from the medium/1.035/1.043 interfaces and was
either subjected to FACS or cultured in DMEM containing 10% FBS.

FACS. HSCs were subjected to FACS using a FACS Aria sorter (BD Bioscience) in the
USC Flow Cytometry Core. GFP was detected by an argon laser and a 530 nm filter.
100
Autofluorescence of VitA was analyzed with a krypton laser and a 424 nm filter. Cells
were sorted based on the intensities of GFP and VitA autofluorescence.

Immunocytochemistry. Cultured HSCs were fixed with 4% paraformaldehyde in PBS for
10 min at 4 °C. After bleaching TOMATO fluorescence, the cells were blocked with 5%
serum for 30 min and incubated with primary antibodies for 1 h at room temperature. The
primary antibodies were detected with secondary antibodies conjugated to fluorescent
dyes. The antibodies used in immunostaining are listed in Table 1. The signals were
captured with a fluorescent microscope (Axio Observer; Carl Zeiss) equipped with a
digital camera (AxioCam; Carl Zeiss).

QPCR. QPCR method was described in Chapter 2. Primer sequences are listed in Table
2.

Statistical analysis. Statistical significance was assessed by ANOVA followed by post-
hoc Tukey HSD test among multiple samples or Student’s t-test between two samples. A
P value of less than 0.05 was considered statistically significant.

101
4.3 Results
MesP1
+
Mesoderm Gives Rise to HSCs, PFs, SMCs, and MCs in the Adult Liver
MESP1 is a basic helix-loop-helix transcription factor transiently expressed in early
mesoderm during mouse gastrulation (Saga et al., 1999). The MesP1
Cre
mouse has been
used for tracing a mesodermal lineage in the developing heart.

We previously
demonstrated that liver mesenchymal cells, including HSCs, fibroblasts around the vein,
and SMCs in the portal vein, are derived from MesP1+ mesoderm in embryonic livers
using the MesP1
Cre
and Rosa26lacZ
flox
mice (Asahina et al., 2009; Asahina et al., 2011).
However, it remained to be determined whether these mesodermal mesenchymal cells are
the source of myofibroblasts in liver fibrosis. Furthermore, recent studies raised the
possibility that HSCs undergo mesenchymal-epithelial transition (MET) and give rise to
hepatocytes and oval cells in injured livers (Yang et al., 2008; Michelotti et al., 2013).
Thus, the present study was undertaken to trace cell lineages of the MesP1+ mesoderm-
derived mesenchymal cells in the adult liver using MesP1
Cre
and R26TG
fl
reporter mice.
Upon recombination by Cre, the R26TG
fl
mouse switches from expression of membrane-
tagged TOMATO to GFP (Fig. 19A) (Muzumdar et al., 2007). The membrane-bound
GFP enabled us to observe cell morphology and unequivocally identify GFP-expressing
cells in the tissues. In the normal MesP1
Cre/+
; R26TG
fl/fl
adult liver, TOMATO was
expressed on the membrane of hepatocytes that could be distinguished by their large
round nuclei (Fig. 19B). GFP was expressed in cells in the sinusoid but not in
hepatocytes (Fig. 19B). After quenching TOMATO fluorescence with methanol, we
characterized GFP expressing cells via immunohistochemistry. As shown in Fig. 19C,
102
many DES+ HSCs expressed GFP in the sinusoid (80.9±3.1%, 1,656 DES+ HSCs
examined, n=3 mice). MCs, which act as progenitor cells for HSCs (Asahina et al., 2011;
Li et al., 2013), expressed GFP on the liver surface (Fig. 19C). PDPN is a mucin-type
transmembrane glycoprotein and is expressed in MCs, the lumen of the bile duct, and
lymphatic vessel in the mouse liver (Li et al., 2013). We confirmed the specific
expression of GFP in PDPN+ MCs (Fig. 19D). GFP expression was also observed in
DES+ fibroblasts in the connective tissue around the portal vein and bile duct (Fig. 19C).
Deposition of ELN, a marker for PFs (Li et al., 2007), was observed around the GFP+
fibroblasts (Fig. 19E), indicating mesodermal origin of PFs. GFP was also expressed in
ACTA2+ SMCs of the portal vein and hepatic artery (Fig. 19C,F). In addition,
endothelial cells in the portal vein and hepatic artery expressed GFP (Fig. 19C,F). As
expected, no GFP expression was observed in CDH1+ HNF4+ hepatocytes and CDH1+
CK19+ cholangiocytes in the liver (Fig. 19G-I). Cell lineage tracing demonstrated that
MesP1+ mesoderm gives rise to MCs and mesenchymal cells, including HSCs, PFs, and
SMCs, but not epithelial cells, such as hepatocytes and cholangiocytes, in the adult
mouse liver.

103
Figure 19

Figure 19. Contribution of MesP1+ mesoderm to HSCs, PFs, SMCs, and MCs, but not
hepatocytes and cholangiocytes, in the adult liver. (A) A cell lineage analysis using the
MesP1
Cre
and R26TG
fl
mice. (B) Expression of TOMATO and GFP in the MesP1
Cre/+
;
R26TG
fl/fl
adult liver. Membrane-tagged TOMATO and GFP are observed in hepatocytes
104
(hc) and HSCs (hsc) in the sinusoid, respectively. No GFP expression in hepatocytes
(2,000 hepatocytes examined; n=3 mice). (C-I) Immunohistochemistry of the adult
MesP1
Cre/+
; R26TG
fl/fl
mouse liver for GFP with DES (C), PDPN (D), ELN (E), ACTA2
(F), CDH1 (G), HNF4 (H), or CK19 (I). Arrows indicate GFP+ HSCs in the sinusoid.
Arrowheads and double arrows indicate DES+ ELN+ PFs adjacent to the bile duct (bd)
and GFP+ MCs on the liver surface, respectively. Double arrowheads indicate DES+
ACTA2+ SMCs in the portal vein (pv) and hepatic artery (ha). GFP+ endothelial cells in
the portal vein and hepatic artery are indicated by asterisks. No GFP expression in CDH+
HNF4+ hepatocytes and CDH+ CK19+ cholangiocytes (2,049 CDH1+, 1,836 HNF4+
hepatocytes and 1,036 CDH1+, 1,093 CK19+ cholangiocytes examined; n=3). Nuclei
were counterstained with DAPI. Scale bar: 10 m.

105
Rare Contribution of the MesP1+ Mesoderm to Kupffer Cells
PDPN+ bile duct and lymphatic endothelial cells were negative for GFP (Fig. 20A).
Expression of GFP and CD31, a marker for endothelial cells, or CD45, a leukocyte
marker, demonstrated only minimal overlap in the sinusoid (Fig. 20B,C). A few F4/80+
Kupffer cells were positive for GFP (Fig. 20D). The control MesP1
+/+
; R26TG
fl/fl
liver
did not show GFP expression (Fig. 20E,F). Cell lineage tracing demonstrated that few
Kupffer cells are derived from MesP1+ mesoderm in the adult mouse liver.

106
Figure 20

Figure 20. Few Kupffer cells express GFP in MesP1
Cre/+
; R26TG
fl/fl
mouse liver. (A-D)
Immunohistochemistry of the adult MesP1
Cre/+
; R26TG
fl/fl
mouse liver mouse liver for
GFP with PDPN (A), CD31/PECAM1 (B), CD45 (C), or F4/80 (D). (A) No GFP
expression in PDPN+ bile duct (bd) and lymphatic endothelial cells (lv). ha, hepatic
artery. (B) Asterisks indicate CD31+ sinusoidal endothelial cells (sec), which are located
in close proximity to GFP+ HSCs (hsc, arrows). No GFP expression in hepatocytes (hc).
(C) No GFP expression in CD45 leukocytes in the sinusoid. (D) A rare GFP+ F4/80+
Kupffer cell. (E) Immunostaining of GFP and DES in the MesP1
+/+
; R26TG
fl/fl
mouse
liver. No GFP expression. (F) Negative control without primary antibodies. Nuclei were
counterstained with DAPI. Scale bar: 10 m.

107
Mesodermal Origin of HSCs
To validate specific expression of GFP in HSCs, we isolated HSCs from the normal
MesP1
Cre/+
or MesP1
+/+
; R26TG
fl/fl
adult livers via collagenase perfusion and subsequent
discontinuous gradient centrifugation. In culture, primary HSCs isolated from the
MesP1
Cre/+
; R26TG
fl/fl
mouse showed mutually exclusive expression of TOMATO and
GFP, which indicated no de novo Cre activation in HSCs (Fig. 21A). The control HSCs
isolated from the MesP1
+/+
; R26TG
fl/fl
liver did not show GFP expression (Fig. 21A).
After quenching TOMATO fluorescence, the HSCs were immunostained with antibodies
against ACTA2 or DES. As shown in Fig. 21B, both GFP+ and GFP- HSCs expressed
ACTA2 or DES at day 7 in culture. A negative control without first antibodies did not
show signal (Fig. 21C) and HSCs from MesP1
+/+
; R26TG
fl/fl
liver did not show GFP
expression (Fig. 21D). These results demonstrate that HSCs are mesodermal in origin and
are capable of differentiating into ACTA2+ myofibroblasts.

108
Figure 21

Figure 21. In vitro activation of mesodermal HSCs. HSCs were isolated from the normal
MesP1
Cre/+
or MesP1
+/+
; R26TG
fl/fl
adult livers and were cultured for 6 (A) and 7 days (B-
D). (A) Expression of TOMATO and GFP in HSCs. Asterisks indicate GFP+ HSCs
isolated from the MesP1
Cre/+
; R26TG
fl/fl
adult liver (Cre+). No GFP expression in HSCs
from the MesP1
+/+
; R26TG
fl/fl
liver (Cre-). (B) Immunostaining of GFP and ACTA2 or
DES in HSCs isolated from the MesP1
Cre/+
; R26TG
fl/fl
adult liver (Cre+). Asterisks
indicate GFP+ HSCs expressing ACTA2 or DES. Arrows indicate GFP- HSCs
expressing ACTA2 or DES. (C) A negative control without primary antibodies. (D)
109
Immunostaining of GFP and DES. No GFP expression in HSCs isolated from the
MesP1
+/+
; R26TG
fl/fl
liver (Cre-). Nuclei were counterstained with DAPI.

110
Activation of Mesodermal HSCs in Culture
We noted that there were GFP- HSCs in the MesP1
Cre/+
; R26TG
fl/fl
liver. To quantify
GFP+ HSCs in the MesP1
Cre/+
; R26TG
fl/fl
liver, we analyzed HSCs via FACS on the basis
of GFP expression and storage of VitA lipids. Combining the detection of VitA
autofluorescence and GFP using 350 (FL5) and 530 nm (FL1) filters, we separated the
primary HSCs isolated from the MesP1
Cre/+
; R26TG
fl/fl
(Cre+)

liver into VitA+GFP- or
VitA+GFP+ fractions (Fig. 22A). FACS analysis revealed that approximately 70±9.4%
(n=4) of VitA+ HSCs express GFP (Fig. 22A). The control MesP1
+/+
; R26TG
fl/fl
(Cre-)
liver did not show GFP expression (Fig. 22A). We cultured HSCs isolated from the
MesP1
Cre/+
; R26TG
fl/fl
liver for 13 days and performed another FACS analysis. As shown
in Fig. 22B, the proportion of GFP+ HSCs remains the same (71%) in culture. These
results indicate that at least 70% of HSCs derived from the MesP1+ mesoderm in the
adult liver.
In general, the Cre-loxP system cannot achieve 100% recombination in mouse
tissues due to insufficient amounts of Cre expression, DNA recombination, and partial
inactivity of the promoter at the Rosa26 gene locus. Nevertheless, our results suggest that
30% of HSCs arise from origin(s) other than the MesP1+ mesoderm. To determine
whether the GFP+ and GFP- HSCs have different characteristics, we sorted the
VitA+GFP+ and VitA+GFP- HSCs from the primary (day 0) or culture activated HSCs
(day 13) via FACS and compared their gene expression by QPCR (Fig. 22C,D). After
isolation of HSCs by collagenase perfusion and discontinuous centrifugation,
approximately 95% of the cells exhibited autofluorescence of VitA. The primary HSCs
111
before FACS expressed HSC markers such as Des and Col1a1 (Fig. 22C, lane 1). QPCR
detected inevitable contamination of hepatocytes (Alb), endothelial cells (Cd31), and
Kupffer cells (Cd68) in the primary HSCs before FACS (Fig. 22D, lane 1). After sorting
HSCs into VitA+GFP+ and VitA+GFP- fractions, the expression of these markers
became undetectable (Fig. 22D, lanes 2,3), which confirmed the successful purification
of HSCs. These genes were also undetectable in the day-13 HSCs (Fig. 22D, lane 4),
suggesting that contaminating non-HSCs do not survive in culture.
The sorted VitA+GFP+ and VitA+GFP- HSCs from primary HSCs showed
similar levels of Des, Col1a1, Timp1, Hgf, and Tgfb1 expression (Fig. 22C, lanes 2 and
3). Upon activation in culture, primary HSCs showed increased expression of activation
markers (Acta2, Col1a1, and Timp1) when not subjected to FACS (Fig. 22B, lanes 1 and
4), while both VitA+GFP+ and VitA+GFP- HSCs from day-13 HSCs showed similar
increases in these genes (lanes 5 and 6). This result suggests that the both GFP+ and
GFP- HSCs have a similar phenotype in the normal liver and upon activation in vitro.

112
Figure 22

Figure 22. Similar phenotypes of GFP+ and GFP- HSCs in the MesP1
Cre/+
; R26TG
fl/fl

liver.(A) HSCs were isolated from the MesP1
Cre/+
; R26TG
fl/fl
(Cre+) or MesP1
+/+
;
R26TG
fl/fl
(Cre-) livers and were subjected to FACS. HSCs were analyzed by storage of
VitA (FL5) and expression of GFP (FL1). (B) HSCs were isolated from the Cre+ and
Cre- mice, cultured for 13 days, and subjected to FACS analysis. (C,D) HSCs were
113
isolated from the MesP1
Cre/+
; R26TG
fl/fl
liver and the primary (day 0) and cultured HSCs
(day 13) were subjected to QPCR. In addition, VitA+GFP+ and VitA+GFP- HSCs sorted
from the primary or day-13 HSCs were also subjected to QPCR for measurement for
HSC markers (C) and other liver cell types (D). The results are expressed as relative
expression compared against primary HSCs (day 0). The values were normalized against
Gapdh. Each value is the mean ± standard deviation of the triplicate measurements.

114
No Contribution of HSCs and PFs to Hepatocytes and Cholangiocytes in Liver
Fibrosis Given that the MesP1
Cre/+
; R26TG
fl/fl
mouse labels 70% of liver mesenchymal
cells in the adult liver, we tested whether the MesP1+ mesoderm-derived mesenchymal
cells undergo MET and differentiate into hepatocytes and cholangiocytes in liver fibrosis.
Following 30 CCl
4
injections, mouse livers developed fibrosis around the central vein.
GFP expression was observed in mesenchymal cells in fibrotic septa or sinusoids and did
not overlap with TOMATO+ hepatocytes (Fig. 23A). Immunohistochemistry confirmed
that ACTA2+ DES+ myofibroblasts co-express GFP (Fig. 23B,C, 80.0±3.6% in
ACTA2+ myofibroblasts; n=3); however, no GFP expression was observed in HNF4+
hepatocytes and CK19+ cholangiocytes (Fig. 23D,E).
We also analyzed the differentiation potential of the mesenchymal cells in biliary
fibrosis. Two weeks after BDL, no GFP expression was observed in TOMATO+
cholangiocytes and hepatocytes (Fig. 23F). Immunohistochemistry showed that ACTA2+
myofibroblasts express GFP adjacent to the dilated bile duct (Fig. 23G, 79.3±4.0% in
2,865 ACTA2+ myofibroblasts examined; n=3). GFP expression was observed in DES+
myofibroblasts and activated HSCs in the sinusoid (Fig. 23H) but not in HNF4+
hepatocytes and CK19+ cholangiocytes (Fig. 23I,J). These data demonstrate that
mesodermal mesenchymal cells, including HSCs and PFs, comprise a major source of
myofibroblasts and do not undergo a MET during fibrogenesis to yield hepatocytes and
cholangiocytes.

115
Figure 23

Figure 23. No contribution of mesodermal mesenchymal cells to hepatocytes and
cholangiocytes in liver fibrosis. Lineage of mesodermal mesenchymal cells was traced
using the MesP1
Cre/+
; R26TG
fl/fl
mouse in liver fibrosis induced by CCl
4
injections 30
116
times (A-E) or by BDL for 2 weeks (F-J). (A,F) Expression of TOMATO and GFP in the
liver. No GFP expression in TOMATO+ hepatocytes (hc) and cholangiocytes (bd). An
arrow indicates GFP+ HSCs in the sinusoid. (B-E) Immunohistochemistry of the CCl
4
-
induced fibrotic livers with antibodies against GFP and ACTA2 (B), DES (C), HNF4 (D),
or CK19 (E). GFP is expressed in ACTA2+ DES+ myofibroblasts (arrowheads) and
DES+ activated HSCs in the sinusoid (arrows) but not in HNF4+ hepatocytes and CK19+
cholangiocytes (1,645 hepatocytes and 1,322 cholangiocytes examined; n=3 mice). (G-J)
Immunohistochemistry of the BDL-induced fibrotic livers with antibodies against GFP
and ACTA2 (G), DES (H), HNF4 (I), or CK19 (J). GFP is expressed in ACTA2+ DES+
myofibroblasts (arrowheads) and DES+ activated HSCs in the sinusoid (arrows) but not
in HNF4+ hepatocytes and CK19+ cholangiocytes (1,852 hepatocytes and
2,560cholangiocytes examined; n=3). Nuclei were counterstained with DAPI. Scale bar:
10 m.

117
No Contribution of HSCs and PFs to Oval Cells
Next, we challenged the notion that HSCs are progenitor cells for oval cells and give rise
to hepatocytes and cholangiocytes via MET (Yang et al., 2008; Michelotti et al., 2013).
After inducing oval cells with a DDC diet for 4 weeks, GFP expression was observed in
myofibroblasts adjacent to GFP- TOMATO+ oval cells (Fig. 24A, 81.7±2.5% in
ACTA2+ myofibroblasts; n=3). The immunohistochemistry of A6 antigen (Engelhardt et
al., 1990) or EPCAM, markers for oval cells, confirmed no GFP expression in oval cells
(Fig. 24B,C). We noted rare GFP+ cells embedded in oval cells, but these cells did not
co-express EPCAM (Fig. 24C, double arrowheads). Oval cells expressing PROM1
(CD133)(Rountree et al., 2007) did not express GFP (Fig. 24D). No GFP expression was
observed in CK19+ cholangiocytes/oval cells and CDH1+ HNF4+ hepatocytes (Fig. 24
E-G). GFP+ myofibroblasts were associated with GFP- oval cells and expressed ACTA2
or THY1 (Fig. 24H,I). The cell lineage tracing refutes the notion that HSCs undergo
MET and differentiate into oval cells, cholangiocytes, and hepatocytes.

118
Figure 24

Figure 24. No contribution of mesodermal mesenchymal cells to oval cells, hepatocytes
and cholangiocytes in injured liver. Lineage of mesodermal mesenchymal cells was
traced using the MesP1
Cre/+
; R26TG
fl/fl
mouse in injured liver induced by DDC diet for 4
119
weeks. (A) Expression of TOMATO and GFP in the liver. No GFP expression in
TOMATO+ oval cells (oc). Arrowheads indicate mesenchymal cells. bd, bile duct; ha,
hepatic artery. (B-I) Immunohistochemistry of the injured liver with antibodies against
GFP and A6 (B), EPCAM (C), PROM1 (D), CK19 (E), CDH1 (F), HNF4 (G), ACTA2
(H) or THY1 (I). (B-D) No GFP expression in oval cells expressing A6 antigen, EPCAM,
and PROM1 (1,899 EPCAM+ oval cells examined; n=3 mice). Arrowheads indicate
GFP+ myofibroblasts associated with GFP- oval cells. Asterisks indicate EPCAM+ oval
cells. Double arrows show rare GFP+ cells embedded in oval cells. pv, portal vein. (E-G)
No GFP expression in CK19+ cholangiocytes/oval cells and CDH1+ hepatocytes
(hc)/oval cells, and HNF4+ hepatocytes (2,102 CK19+ cells, 2,749 CDH1+ cells and
2,120 HNF4+ hepatocytes examined; n=3). (H,I) Myofibroblasts associated with GFP-
oval cells co-express GFP with ACTA2 or THY1. lv, lymphatic vessel. Nuclei were
counterstained with DAPI. Scale bar: 10 m.

120
No Contribution of Mesodermal Mesenchymal Cells to Epithelial Cells in the
Regenerating Liver
Following the induction of oval cells, we halted the DDC diet, fed the mice normal chow
for 4 weeks, and determined whether GFP+ mesenchymal cells give rise to hepatocytes
or cholangiocytes during recovery from injury. Termination of the DDC diet reduced the
number of ACTA2+ myofibroblasts and oval cells expressing EPCAM (Fig. 25A-C).
Although mesenchymal cells continued to express GFP, neither EPCAM+ CK19+
cholangiocytes nor CDH1+ HNF4+ hepatocytes were positive for GFP (Fig. 25C-F).
Collectively, these data demonstrate that mesodermal mesenchymal cells, including
HSCs and PFs, do not contribute to oval cells, hepatocytes, and cholangiocytes in mouse
liver injury.

121
Figure 25

Figure 25. No contribution of mesodermal mesenchymal cells to hepatocytes and
cholangiocytes in the regenerating liver. Lineage of mesodermal mesenchymal cells was
traced using the MesP1
Cre/+
; R26TG
fl/fl
mouse. After induction of oval cells by DDC diet
for 4 weeks, we changed the diet to the normal chow and induced regeneration for 4
weeks. (A) Expression of TOMATO and GFP in the liver. No GFP expression in
TOMATO+ hepatocytes (hc). bd, bile duct; pv, portal vein. (B-F) Immunohistochemistry
of the regenerating liver with antibodies against GFP and ACTA2 (B), EPCAM (C),
CK19 (D), CDH1 (E), or HNF4 (F). (B) No ACTA2 expression in GFP+ HSCs (hsc) in
the sinusoid. ha, hepatic artery. (C) No GFP expression in EPCAM+ cholangiocytes.
122
Arrowheads indicate GFP+ mesenchymal cells adjacent to EPCAM+ GFP cholangiocytes.
(D-F) No GFP expression in CK19+ CDH1+ cholangiocytes and CDH1+ HNF4+
hepatocytes (2,072 HNF4+ hepatocytes and 1,002 CK19+ cholangiocytes examined;
n=2). Nuclei were counterstained with DAPI. Scale bar: 10 m.

123
4.4 Discussion
The present study demonstrates that HSCs derive from the MesP1+ mesoderm in the
adult liver. Our data demonstrate that this mesoderm population contributes to HSCs but
not to endodermal cells, such as hepatocytes and cholangiocytes, in the adult liver (Fig.
26). Given that MesP1 is transiently expressed during gastrulation, and not expressed in
the developing and adult liver, our data indicate that no differentiation of HSCs to
endodermal epithelial cells occurs during or after liver development.
We previously found that MCs migrate inward from the liver surface and give rise
to HSCs, fibroblasts around the vein, and SMCs during liver development (Asahina et al.,
2011). In contrast, MCs did not differentiate into liver epithelial cells. In agreement with
this notion, the present study found that fibroblasts around the portal vein and SMCs in
the portal vein are derived from the MesP1+ mesoderm similar to HSCs (Fig. 26). We
defined these fibroblasts around the portal vein as PFs based on expression of ELN, a
marker for rat PFs, and DES. In the BDL model using the MesP1
Cre
; R26TG
fl
mouse,
GFP+ myofibroblasts accumulated around the bile duct in the portal area and no GFP+
cholangiocytes and hepatocytes were found, indicating that PFs are the primary source of
myofibroblasts and that no MET occurs during biliary fibrosis. Similar to the BDL
model, CCl
4
-induced liver fibrosis did not provoke MET from MesP1+ mesoderm-
derived HSCs to hepatocytes and cholangiocytes in the MesP1
Cre
; R26TG
fl
mouse (Fig
26). In conclusion, our study did not find evidence of mesodermal mesenchymal cells
undergoing MET to become hepatocytes or cholangiocytes during liver fibrosis.
124
Contrary to our conclusion, Yang et al. reported that HSCs give rise to oval cells
using the GFAP
Cre
and Rosa26YFP reporter mice (Yang et al., 2008). These authors
traced HSCs based on the expression of GFAP, which is a marker for HSCs in rats and
humans. However, in mice, both HSCs and cholangiocytes express GFAP. Given that
cholangiocytes are likely to be the origin of oval cells, the GFAP
Cre
mouse would label
cholangiocyte-derived oval cells in addition to HSCs. Thus, the GFAP
Cre
mouse is not
appropriate for tracing HSC lineages. The same group also reported differentiation of
myofibroblasts to hepatocytes and cholangiocytes in the BDL model (Michelotti et al.,
2013). After BDL, they labeled activated HSCs by tamoxifen using the ACTA
CreERT2
and
Rosa26YFP
flox
mouse and subsequently found YFP+ liver epithelial cells. In contrast to
their findings, our study did not show such transdifferentiation from HSCs to liver
epithelial cells in the BDL model in the MesP1
Cre
; R26TG
fl
mice. One possible reason for
this discrepancy is that the BDL surgery on the ACTA
CreERT2
mouse may activate de novo
expression of Acta2 mRNA in epithelial cells, thereby inducing YFP expression upon
tamoxifen injection in Acta2-expressing epithelial cells independent of HSC
transdifferentiation. In fact, the same group reported induction of ACTA2 expression in
hepatocytes and cholangiocytes (Sicklick et al., 2006; Yang et al., 2008). Similar to our
conclusion, no MET from HSCs to liver epithelial cells was observed in CCl
4
-induced
fibrosis using Collagen1a2
Cre
and Vimentin
CreERT2
mice (Scholten et al., 2010; Troeger et
al., 2012). In addition, Mederacke et al. generated a HSC-specific lecithin-retinol
acyltransferase (Lrat) Cre mouse line, which efficiently marks 99% of HSCs in the mouse
liver (Mederacke et al., 2013). Using this mouse, they reached the same conclusion that
125
HSCs are the major source of myofibroblasts and do not give rise to oval cells,
hepatocytes, and cholangiocytes. An important addition of our study using the MesP1
Cre

mouse is that we demonstrated at least 70% of HSCs are derived from MesP1+
mesoderm and they do not differentiate into endodermal epithelial cells throughout liver
development, growth, injury, and regeneration. Therefore, we conclude that mesodermal
liver mesenchymal cells are the primary source of myofibroblasts and do not undergo
MET in vivo to yield oval cells or other liver epithelial cell types.
We found that 70% of HSCs express GFP in the MesP1
Cre
; R26TG
fl
mouse liver.
It is unclear why not all HSCs express GFP in this model, but inefficiency of DNA
recombination by Cre or silencing of the CAG promoter in the Rosa26 locus may have
attributed to the generation of GFP- HSCs in the liver. Another possible explanation is
that the GFP- HSCs originated from other sources than the MesP1+ mesoderm. Although
we cannot identify the origin of the GFP- HSCs in this model, we did not observe
differences between GFP+ and GFP- HSCs in the MesP1
Cre
; R26TG
fl
mouse in
morphology or gene expression in either the quiescent or activated state. MesP1
expression was restricted to the portion of the mesoderm ingressed through the primitive
streak in early embryos (Saga et al., 1999). Therefore, GFP- HSCs might have originated
from the MesP1- mesoderm adjacent to the MesP1+ mesoderm area.
HSCs have been known to exhibit multipotency in culture (Kordes et al., 2007;
Conigliaro et al., 2013). However, we have not been able to detect the transdifferentiation
of HSCs in embryonic or adult livers. As demonstrated by direct cell reprogramming with
defined factors in vitro (Aoi et al., 2008), in vitro culture environments might change
126
gene expression and chromatin state of HSCs and negate the negative regulation that
inhibits the multi-differentiation potential of HSCs in vivo. Although HSCs can
differentiate into hepatocytes in vitro, our study indicates that the in vivo environments
are not permissive for HSCs to alter their programmed lineage to differentiate into other
germ layers. Our data indicate that the fate of liver mesenchymal cells is tightly regulated
in vivo.
Myofibroblasts synthesize proinflammatory cytokines and extracellular matrices
and participate in liver fibrosis. Our studies determined that liver mesenchymal cells are
the major source of myofibroblasts. Specific targeting of HSCs, PFs, or MCs will be an
important issue for the suppression of fibrosis and its progression to cirrhosis.

127
Figure 26

Figure 26. A proposed model of MesP1+ mesoderm in the liver. Genetic cell lineage
tracing revealed that the MesP1+ mesoderm gives rise to MCs, PFs and HSCs. Upon
liver injury induced by CCl
4
or BDL, all of these mesodermal mesenchymal cells
differentiate into myofibroblasts, but not into hepatocytes, cholangiocytes, or oval cells.

128
Chapter 5
Lipid mediators in liver fibrosis
5.1 Introduction
In previous chapters, we present evidence that TGF-  induces myofibroblastic
conversion of liver mesenchymal cells, including MCs, HSCs, and PFs. Condition
deletion of Tgfbr2 gene in MCs suppressed MMT and fibrosis. These results support the
concept that TGF-  plays a critical role in the generation of myofibroblasts in organ
fibrosis (Hinz et al., 2012). Although many tools are available to target TGF-  signaling,
it is difficult to regulate TGF-  signaling to specifically suppress fibrosis because of its
diverse roles in HSC activation, inhibition of hepatocyte proliferation, and suppression of
inflammation (Dooley and ten Dijke, 2012). Furthermore, our laboratory recently
reported that conditional deletion of Tgfbr2 gene in MCs did not completely block the
migration and differentiation of MCs to myofibroblasts and HSCs in liver injury (Li et al.,
2016), implying that other factors may also be involved in MMT in liver fibrosis. In the
present study, we found that lysophosphatidic acid (LPA) induces the myofibroblastic
conversion of MCs and HSCs. LPA is a small phospholipids consisting of a glycerol
backbone with fatty acid and phosphate groups (Fig. 27) (Moolenaar and Perrakis, 2011;
Knowlden and Georas, 2014).

129
Figure 27

Figure 27. Chemical structures of major species of LPA. 18:1 form is the most
commonly used in laboratory experiments for activating LPA receptors.

130
LPA acts as an extracellular ligand and bind to its receptor called
lysophosphatidic acid receptors (LPARs). LPARS are G protein-coupled receptors
(GPCRs) and six receptors have been identified: LPAR1 (EDG2), LPAR2 (EDG4),
LPAR3 (EDG7), LPAR4 (P2Y9, GPR23), LPAR5 (GPR92), and LPAR6 (P2Y5)
(Yanagida and Ishii, 2011; Yung et al., 2014). All of LPARs have a seven transmembrane
domain and are associated with heterotrimeric G proteins. After LPA binding, LPARs
induce the dissociation of GTP-bound G  from G  subunits and activate different
downstream signaling pathways (Choi et al., 2010). Among the four G  subfamilies, G 
12/13 can induce actin polymerization via Rho GTPases (Jaffe and Hall, 2005; Kelly et
al., 2007).
Extracellular LPA is synthesized by two major pathways: (1) from the activity of
autotaxin (Atx/Enpp2) on lysophosphatidylcholine (LPC), and (2) from phospholipase
A1 or A2 activity on phosphatidic acid (Fig. 28A,B) (Knowlden and Georas, 2014; Yung
et al., 2014). Although ATX was originally discovered as a tumor cell autocrine motility
factor, it was later identified as a secreted lysophospholipase D that synthesizes LPA
from LPC (Tokumura et al., 2002; Umezu-Goto et al., 2002). Atx-deficient mice are
embryonic lethal with multiple abnormalities (van Meeteren et al., 2006). Atx-
heterozygous mice appear healthy but show half-normal plasma LPA levels, suggesting
that ATX is a major LPA-producing pathway. Both LPA and ATX have short half-lives
in mice and are rapidly cleared from the circulation within minutes by sinusoid
endothelial cells (SECs) (Jansen et al., 2009; Albers et al., 2010). The serum
concentration of LPC is around 200-500 M. In contrast, LPA is present at low
131
concentrations of 0.1-0.3 M in plasma to 2M in serum as an albumin-bound form
(Aoki et al., 2002; Watanabe et al., 2007a). Although serum ATX and LPA levels
increase 3-5-fold in patients with liver fibrosis and correlate significantly with the stage
of fibrosis (Watanabe et al., 2007b; Nakagawa et al., 2011; Pleli et al., 2014), no studies
have examined whether LPA is involved in liver fibrogenesis.

132
Figure 28

A B

Figure 28. LPA synthesis pathways. (A) Extracellular production of plasma LPA by
ATX. (B) Production of LPA by phospholipase A1 and A2. PC, phosphatidic choline; PA,
phosphatidic acid.

133
Hippo signaling negatively regulates coactivators YAP (Yes-associated protein)
and TAZ/Wwtr1 (Transcriptional coactivator with PDZ-domain motif), which play
important roles in tissue homeostasis, organ size, and tumor development (Yu and Guan,
2013; Johnson and Halder, 2014; Varelas, 2014). In mammals, MST1/2 kinases (Stk4/3,
Drosophila Hippo ortholog) activate LATS1/2 kinases (Warts) and induce cytoplasmic
retention of two Yorkie ortholog proteins, YAP and its paralog TAZ by phosphorylation.
Upon inhibition of Hippo signaling, unphosphorylated YAP/TAZ translocate into nuclei
and act as coactivators of TEAD1-4 (Scalloped) for the regulation of target gene
expression, such as Ctgf. YAP overexpression in hepatocytes causes enlargement of the
liver and cancer in mice (Camargo et al., 2007; Dong et al., 2007). In this study, we
found that LPA synergistically enhances TGF- -induced MMT via the nuclear
localization of TAZ/Wwtr1. TAZ, but not YAP, mediated TGF-  and LPA-induced
myofibroblastic conversion of MCs and HSCs. Furthermore, antagonism of LPA
signaling by injecting BrP-LPA, a pan-inhibitor for LPAR1-4 and ENPP2 (Jiang et al.,
2007), resulted in suppression of fibrosis in mouse livers. Based on these data, we
concluded that LPA acts as profibrogenic factors and its signaling pathway through TAZ
is a novel therapeutic target for the suppression of fibrosis.

134
5.2 Materials
Mice. Immortomouse was purchase from Charles River (Jat et al., 1991). Mst1/Mst2
fl/fl

mouse was purchase from the Jackson Laboratory (Lu et al., 2010). Wt1
CreERT2
knock-in
mice and R26TG
fl
mice were described previously in Chapter 2. Tamoxifen injection was
the same as described in Chapter 2. Two weeks after the last injection, mice were treated
with of single injection of CCl
4
(1l/g body weight) and BrP-LPA (1-Bromo-3(S)-
hydroxy-4- (palmitoyloxy)butyl]phosphonate, 2.5-10 g/g body weight, Echelon)
intraperitoneally on the same day as CCl
4
and on the next day. For 3 times CCl
4
injection,
mice were treated with BrP-LPA (7.5 g/g body weight).

Cell lines and cell culture. MC lines (MCLs) was established by using the
Immortomouse that expresses the early region of the SV40 mutant tsA58 under the
control of IFN-  responsive H-2K
b
promoter (Jat et al., 1991). Primary MC and HSC
isolation were described previously in Chapter 2 and 3. Cells were cultured in Ca
2+
depleted medium or serum stravation overnight and treaed with 4 M AM095
(APExBIO), 2.5-10M BrP-LPA (Echelon), 0.2 g/ml Exoenzyme C3 transferase
(Cytoskeleton), 0.5 M Latrunculin A (Santa Cruz), 5-20 M LPA (18: 11-oleoyl-2-
hydroxy-sn-glycero-3-phosphate, Avanti) mixed with 1% essential fatty acid free BSA in
PBS, 10 ng/ml TGF- 1 (Sigma-Aldrich) or 5 M Y-27632 (Calibiochem) in serum free
condition.

135
Cell isolation. MCs and HSCs isolation method were described previously in Chapter 2
and 3. Hepatocytes, SEC and Kupffer cells were isolated by the NPC Core in University
of Southern California (Kreamer et al., 1986; Schrage et al., 2008; Xu et al., 2015).

Adenovirus. MCs and HSCs were isolated from Mst1/Mst2
fl/fl
mouse and treated with an
adenovirus vector carrying LacZ or Cre (Kerafast, multiplicity of infection 50) from day
2 and treated with 10 ng/ml TGF -1 (Sigma) from day 4 for 12 h to measure mRNA
expression.

Cytochemistry and Immunohistochemistry. Methods were described previously in
Chapter 2 and 3. Primary antibodies are listed in Table1.

RNA interference. MCs and HSCs were grown on 24-well plates for 24 hours and
transfected twice with a 48-hour interval using Dharmafect (0.8 l per well) and the ON-
TARGET plus SMART pool directed against mouse YAP and TAZ (60 pmol per well) or
a nonspecific control siRNAs (Dharmacon). At 24 hours after the last transfection, cells
were treated as indicated and RNA was extracted 48 hours later. Knockdown efficiency
was determined by QPCR.

QPCR. Method was described previously in Chapter 2. Primer sequences are listed in
Table 2.

136
Plasma biochemistry. Plasma ALT was determined by a kinetic assay using the ALT
reagent (Raichem) and a temperature-controlled plate reader.

Statistical analysis. Statistical significance was assessed by ANOVA followed by post-
hoc Tukey HSD test among multiple samples or Student’s t-test between two samples. A
P value of less than 0.05 was considered statistically significant.

137
5.3 Results
Cell-cell Contacts Counteract TGF- -induced MMT
The yield of MCs is around 2x10
4
cells from one adult mouse liver, and primary MCs
cannot be passaged more than three times to maintain their epithelial phenotype (Li et al.,
2013). To overcome this limitation, we established MC lines (MCLs) using the
Immortomouse that expresses the early region of the SV40 mutant tsA58 under the H-
2K
b
promoter controlled by IFN-  treatment (Jat et al., 1991). The resulting liver MCLs
grew in DMEM containnig 5% FBS and IFN-  and maintained an epithelial cell
morphology after passaging at least more than 12 times. MCLs were plated at 2x10
3

cells/cm
2
and cultured for 2 days in the absence or presence of TGF- 1 in DMEM
containing 5% FBS. TGF- 1 induced the morphological change of MCLs towards
fibroblastic cells and the expression of Acta2 mRNA (Fig. 29A,B). TGF- 1 suppressed
the expression of Gpm6a mRNA (Fig. 29B). SB431542, a chemical inhibitor for
TGFBR1, suppressed morphological change of MCLs and Acta2 mRNA expression and
increased Gpm6a mRNA expression (Fig. 29A,B). In contrast, MCs plated 10-time more
(2x10
4
cells/cm
2
) formed epithelial colonies and expressed less Acta2 mRNA and high
Gpm6a mRNA compared to those at low density (Fig. 29A,B). Immunostaining results
also confirmed TGF- 1 did not fully induce ACTA2 expression at high cell density and
have higher GMP6A expression compared with low cell density (Fig. 29C), suggesting
cell-cell contact inhibit MMT.
138
E-cadherin is a calicium (Ca
2+
) dependent adhesion molecule predominatly
express in epithelial cells. Ca
2+
facilitates cadherin-cadherin interactions between
epithelial cells in maintaining cell-cell junctions (Kim et al., 2011). Although MCs shown
an intermediate phenotype between epithelial cells and mesenchymal cells, MCs do not
express E-cadherin (Li et al., 2013; Lua et al., 2015). In fact, MCs cultured in Ca
2+

depleted medium did not show phenotypic changes (Fig. 29D). In combination of Ca
2+

depleted medium with TGF- 1treatment did not increase Acta2 mRNA expression
compared with control medium (Fig. 29E), suggesting that Ca
2+
depletion did not disrupt
MCs cell-cell junctions. These results support the idea that the mechanism of MMT is
different from EMT, in terms of calcium dependent cell-cell contact.

139
Figure 29

Figure 29. TGF-   does not fully induce MMT at high cell density. (A) Cell morphology
of MCLs plated at 2x10
3
cells/cm
2
(Low) and 2x10
4
cells/cm
2
(High). Cells were cultured
for 2 days in the absence or presence of TGF- 1 in DMEM containing 5% FBS. (B)
140
QPCR of A. (C) Immunostaining of A. (D) Cell morphology of MCLs plated at middle
cell density (1x10
4
cells/cm
2
) and cultured in calcium free (Ca
2+
) medium with the
absence or presence of TGF- 1. (E) QCPR of D. *, p < 0.05, **, p < 0.01. Scale bar: 10
m.

141
TAZ Regulates MMT
Cell density regulates nuclear localization of YAP and TAZ, which control cell
proliferation, body size, and cancer growth (Yu and Guan, 2013; Johnson and Halder,
2014; Varelas, 2014). We examined the expression of YAP and TAZ in MCLs cultured
in DMEM containing 5% FBS. Expression of TAZ was observed in the cytoplasm of
MCLs at high cell density (Fig. 30A). At low cell density, MCs showed the nuclear
localization of TAZ that was enhanced by TGF-  treatment. At high cell density, TGF-
1 did not induce the nuclear localization of TAZ in MCLs (Fig. 30A). Similar to TAZ
nuclear localization, YAP also localized to nuclei at low cell density, but not at high
density (Fig. 30A). To determine the roles of YAP and TAZ in MMT, we knocked down
these proteins using siRNAs. MCLs were plated at middle cell density (1x10
4
cells/cm
2
)
and treated siRNAs twice with a 48-hour interval. Cells were further cultured for 2 days
in the presence or absence of TGF- 1. siRNAs for Yap specifically suppressed Yap
mRNA (Fig. 30B). Taz siRNAs efficiently downregulated Taz mRNA (Fig. 30B). Taz
siRNAs also weakly suppressed Yap mRNA. Interestingly, Taz siRNAs suppressed TGF-
 -induced induction of Acta2 mRNA (Fig. 30B). Compared to Taz siRNAs, Yap
siRNAs did not efficiently suppress Acta2 mRNA, indicating that TAZ mediates TGF-
 -induced MMT (Fig. 30B). CTGF are members of the CCN family of cysteine-rich
matricellular proteins, and it is known to be up-regulated by TGF- 1 via SMAD3,
STAT3, or YAP/ TAZ (Zhao et al., 2008; Liu et al., 2013). siRNAs for Yap1 or Taz
reduced Ctgf mRNA expression in MCs (Fig. 30B). However, TGF-  -induced Ctgf
142
expression was not suppressed by these siRNAs, implying that TGF- 1 signaling
overwhelms YAP/TAZ-mediated induction of Ctgf mRNA in MCs. We also confirmed
TAZ-mediated MMT in primary liver MCs. Knock-down by Taz, but not Yap, resulted in
down-regulation of Acta2 mRNAs in primary MCs (Fig. 30C).

143
Figure 30

Figure 30. TAZ regulates MMT. (A) The nuclear TAZ and YAP (arrowheads) are
evident at low cell density. TGF-  enhances its nuclear localization. (B) siRNAs for
Taz, but Yap, suppress TGF-  -induced Acta2 mRNA expression in MCLs (C) and in
primary MCs. Sr: scrambled siRNAs. *, p < 0.05, **, p < 0.01.

144
F-actin Formation Enhances MMT
In canonical Hippo signaling, MST1/2 suppress YAP/TAZ via activation of LATS1/2
kinases. In addition, YAP/TAZ are activated independent of MST1/2 (Yu et al., 2012;
Aragona et al., 2013). We isolated MCs from Mst1/Mst2
fl/fl
mouse livers and deleted both
genes using Adenovirus-Cre. The deletion of Mst1/Mst2 in MCs did not block TGF-
 induced Acta2 mRNA expression (Fig. 31A), suggesting MST1/2 are not required for
TAZ-mediated MMT.
TGF- 1 stimulates SMAD and non-SMAD pathways (Lamouille et al., 2014).
TGF- 1 is known to induce Rho GTPase that induces F-actin formation from G-actin via
downstream effectors, such as ROCK and cofilin (Geneste et al., 2002; Clements et al.,
2005). Although Hippo signaling is a major negative regulator of TAZ, cell density,
polarity, and ECMs also affect TAZ nuclear localization via the cytoskeleton without
contribution of MST1/2 Hippo kinases (Yu et al., 2012; Aragona et al., 2013). In fact,
treatment of MC lines with exoenzyme C3 transferase, a Rho inhibitor, inhibit the
morphological change and expression of Acta2 mRNA induced by TGF- 1 (Fig. 31B,C).
Y27632, a chemical ROCK inhibitor, also decreased TGF- 1-induced Acta2 mRNA.
Treatment with latrunculin A (LatA), which prevents F-actin formation, strongly
suppressed Acta2 mRNA in MC lines in the presence or absence of TGF- 1 (Fig.
31B,C), suggesting that F-actin formation enhances MMT.

145
Figure 31

Figure 31. F actin formation, but not MST1/2, is required for MMT. (A) MCs were
isolated from Mst1/Mst2
fl/fl
mouse and infected with Adeno-lacZ or Adeno-Cre along
with TGF-  treatment. (B) Cell morphology of MCLs treated with inhibitors. (C) QPCR
of B. *, p < 0.05, **, p < 0.01.

146
LPA Enhances TGF- 1-induced MMT
We usually culture MCLs in DMEM containing 5% FBS. We found that TGF- 1 did not
fully induce MMT in DMEM without FBS, even at low cell density (Fig. 32A,B),
compare to (Fig. 29A,B), implying that soluble factor(s) in the FBS enhance TGF- 1-
induced MMT. A recent study suggests that LPA containing FBS induces the nuclear
localization of YAP/TAZ in HEK293 cells (Yu et al., 2012). Thus, we treated MCLs with
LPA (5-20 M) at medium cell density in DMEM without FBS. LPA treatment of MCLs
for 2 days changed their morphology to myofibroblasts (Fig. 32C). QPCR showed the
upregulation of Acta2 and Ctgf by LPA (Fig. 32D). When MCLs were treated with TGF-
1 and different concentrations of LPA, LPA synergistically induced Acta2 mRNA
expression (Fig. 32E). siRNAs for Taz, but not Yap, abrogated Acta2 mRNA expression
induced by TGF-  , LPA, or both (Fig. 32F). These data indicate that LPA
synergistically enhances TGF- 1-induced MMT via TAZ.

147
Figure 32

Figure 32. LPA enhances TGF-  -induced MMT via TAZ. (A) TGF-  does not fully
induce MMT in the absence of FBS. (B) QPCR of A. (C) LPA stimulates MMT. (D)
QPCR of C. (E) LPA synergistically enhances TGF-  -induced Acta2 mRNA expression
in MCs. (F) siRNAs for Taz suppress Acta2 mRNA expression induced by TGF-   and
LPA. *, p < 0.05, **, p < 0.01.

148
LPA Induces MMT
Little is known expression of LPA receptors (Lpar1-6) and autotaxin (Enpp2) in liver
cells. We separated each cell type from normal mouse liver and analyzed gene expression
by QPCR. MCs expressed Lpar1, Lpar2, and Lpar5 (Fig. 33A). MCs and hepatocytes
expressed Enpp2 mRNA, suggesting that both cells synthesize LPA in the liver (Fig.
33A). To test whether LPA is involved in MMT, we treated MCLs with BrP-LPA that
antagonizes LPAR1-4 and ENPP2 (Jiang et al., 2007) or AM095, a specific LPAR1
inhibitor (Swaney et al., 2011). Both inhibitors suppressed LPA-induced Acta2 mRNA
expression in MCs (Fig. 33B,C), suggesting that LPAR1 is a major receptor in MMT.

149
Figure 33

Figure 33. LPAR expression pattern in liver cells. (A) QPCR of Atx and Lpar1-6 in
mouse liver (L), hepatocytes (H), HSCs (SC), SECs (E), Kupffer Cells (K), and MCs
(MC). (B,C) Inhibition of LPA-induced Acta2 mRNA in MCLs by BrP-LPA or AM095
( M). *, p < 0.05, **, p < 0.01.

150
LPA Activates HSCs through TAZ
Although LPA is known to induce the proliferation and migration of HSCs (Ikeda et al.,
1998; Tangkijvanich et al., 2002), it is unclear whether LPA induces HSC activation in
liver fibrosis. Given that LPA acts as a profibrogenic factor in MCs, we tested its effects
on HSCs. HSCs isolated from mouse livers express mRNAs for Lpar1 and Lpar4 (Fig.
33A). Primary HSCs increased Acta2 mRNA expression from 0.5 M of LPA (Fig. 34A)
that is close to the physiological level in patients with liver fibrosis. Different from MCs,
LPA and TGF-  did not show synergistic effects on Acta2 mRNA expression in HSCs
(Fig. 34B). Treatment of HSCs with BrP-LPA or AM095 blocked LPA-induced
expression of Acta2 mRNA (Fig. 34C), suggesting that LPAR1 is largely responsible for
LPA-induced activation. LPA treatment induced the nuclear localization of TAZ in HSCs
(Fig. 34D). siRNAs for Taz, but not Yap, resulted in down-regulation of Acta2 mRNA in
HSCs (Fig. 34E). These results suggest LPA induces activation of HSCs via LPAR1 and
TAZ similar to MMT.

151
Figure 34

Figure 34. LPA induces HSC activation. (A) LPA induces Acta2 mRNA expression in
HSCs ( M). (B) LPA does not enhance TGF--induced HSC activation. (C) BrP-LPA
or AM095 suppresses LPA-induced Acta2 expression ( M). (D) LPA induces the nuclear
localization of TAZ in HSCs (arrowhead). (E) siRNAs for Taz, but not Yap, suppress
Acta2 mRNA expression in HSCs. *, p < 0.05, **, p < 0.01.

152
Inhibition of LPA Reduces Liver Fibrosis
Next, we examined whether LPA is responsible for the induction of MMT and activation
of HSC in liver fibrosis. After labeling MCs as GFP+ cells in Wt1
CreERT2/+
; R26TG
fl/fl

mice, mice were treated with single injection of CCl
4
combined with BrP-LPA for
antagonizing LPA signaling. Two days after CCl
4
injection, livers treated with BrP-LPA
had a significantly decreased induction of Acta2 mRNA in the liver and plasma ALT
values (Fig. 35A,B). Next, we treated Wt1
CreERT2
; R26TG
fl
mice with CCl
4
3 times and
similarly treated with BrP-LPA. BrP-LPA reduced the differentiation of GFP+ MCs
inward (Fig. 35C), expression of ACTA2 around the central vein (Fig. 35D) and plasma
ALT values (Fig. 35E) in CCl
4
-treated liver, implying LPA signaling is required in liver
fibrosis.

153
Figure 35

Figure 35. Inhibition of LPA signaling in liver fibrosis. (A,B) Mice were injected with
CCl
4
and different doses of BrP-LPA. (A) BrP-LPA reduces Acta2 mRNA expression
and (B) plasma ALT level dose dependently. (C, D) Mice were injected with CCl
4
3
times combined with BrP-LPA. (C) Brp-LPA inhibits differentiation of GFP+ MCs to
ACTA2+ myofibroblasts, (D) expression of ACTA2 around the central veins, (E) and
plasma ALT level. Arrowheads indicate myofibroblasts. *, p < 0.05, **, p < 0.01.
154
5.4 Discussion
Our data indicate for the first time that LPA acts as a profibrogenic factor in MMT and
HSC activation in liver fibrosis. LPA was originally discovered as a mitogen in 1989
(Moolenaar and Perrakis, 2011). It is involved in cancer growth and metastasis, as well as
fibrosis of the lung (Yanagida and Ishii, 2011; Yung et al., 2014). Cancer cells synthesize
LPA, which stimulates cancer cell growth and migration (Mills and Moolenaar, 2003). In
lung fibrosis, LPA stimulates the growth of myofibroblasts (Huang et al., 2013).
Although LPA has been reported to mediate pruritus in cholestatic liver disease (Kremer
et al., 2010), very little are known about the contribution of LPA to liver fibrosis.
LPA binds to six different LPARs; different types of agonists or antagonists have
been developed for anti-cancer or anti-lung fibrosis. For example, a chemical inhibitor of
LPAR1 (BMS-9860202/AM152) has been shown to ameliorate lung fibrosis in mice and
is in phase II trials to evaluate its safety and efficacy in idiopathic pulmonary fibrosis
(ClinicalTrials.gov NCT01766817). Similarly, a LPAR1/3 dual antagonist (SAR100842)
is also in phase II clinical trials in patients with cutaneous system sclerosis
(NCT01651143). Since different types of specific LPA inhibitors have been developed to
treat cancer or lung fibrosis, our finding on the possible roles of LPA in the activation of
MCs and HSCs will be applicable to the design of an anti-fibrosis therapy for patients
with fibrosis and cirrhosis.
Even though LPA signaling seems to be a promising therapeutic target for the
suppression of liver fibrosis, underlying mechanisms of LPA signaling remain to be
clarified in MCs and HSCs by rigorous molecular analyses using mouse genetic models
155
before assessing therapeutic effects of LPA inhibitors in liver disease. For instance,
Lpar1
-/-
mice have been used to analyze peritoneal and lung fibrosis. These mice were
shown to be protected from injury by reducing fibroblast recruitment and proliferation,
respectively (Tager et al., 2008; Sakai et al., 2013). Around 50% of Lpar1
-/-
neonates die
due to abnormal suckling behavior in a C57BL/6 background (Contos et al., 2000). The
survivors show reduced size and craniofacial dysmorphism. Interestingly, Lpar1
-/-
mice
in a BALB/c background exhibit normal suckling behavior, survive nearly 100%, and are
fertile. Therefore, BALB/c Lpar1
-/-
mice will be suitable for further study in liver fibrosis.
In addition, conditional deletion of TAZ in MCs and HSCs will be required to precisely
determine the role of TAZ in MMT and HSC activation. Mice lacking Taz gene exhibit a
high prenatal mortality rate and suffer from polycystic kidney disease and emphysema
(Hossain et al., 2007; Tian et al., 2007). Taz
flox
mice are available and will be useful by
crossing MC or HSC Cre lines (Xin et al., 2013).

We demonstrate LPA synergistically enhances TGF- -induced MMT and we
question how TGF-  and LPA cooperatively regulates MMT. Varelas et al. reported TAZ
directly bind to Smad and control Smad nucleocytoplasmic shuttling in human embryonic
stem cell (Varelas et al., 2008). This might be the mechanism for crosstalk between
LPA/TAZ and TGF- /Smad2/3 pathways in MCs.
In sum, the present study demonstrates for the first time that TGF-  and LPA
induces the myofibroblastic conversion of MCs and HSCs via TAZ (Fig. 36). In addition,
inhibition of LPA signaling ameliorates liver fibrosis, indicating LPA signaling will be a
novel therapeutic target for the suppression of liver fibrosis and cirrhosis.
156
Figure 36

Figure 36. A proposed model of LPA-TAZ signaling. LPA induces the activation of Rho
via LPARs. Rho activation results in F-actin formation, which triggers the nuclear
localization of TAZ, although its mechanism is unknown. TAZ acts as coactivator for
TEAD. TGF-  stimulates SMAD3 and possibly Rho. TAZ/TEAD, and SMAD3 might
cooperatively induce MMT upon stimulation with LPA and TGF- .

157
Chapter 6
Conclusions & Perspectives
6.1 Conclusions
The goal of this study was to develop therapeutic strategy against fibrosis by determining
the source of myofibroblasts and underlying molecular mechanisms. Fibrosis is a wound
healing process in response to repeated chronic injury. Although each organ has different
causative etiology, the fibrotic phenotype shares the common feature of excessive
deposition of extracellular matrix proteins, which is produced from myofibroblasts. Thus,
treatment strategy targeting myofibroblasts activation is an effective way for suppressing
fibrosis. However, as we described in Introduction, myofibroblasts are heterogeneous and
have multiple cell sources. In order to target myofibroblasts, we need to first identify
their origin and fundamental molecular mechanism underlying the myofibroblastic
conversion. In Chapter 2, we identified MCs, an underappreciated cell type, are the
common source of myofibroblasts in liver and peritoneal fibrosis. This study
demonstrated for the first time that the protection of MCs will be a therapeutic target for
suppression of fibrosis. In Chapter 3, we invented a new isolation protocol by FACS and
quantified the contribution of MCs, HSCs and PFs in liver fibrosis. We concluded that
HSCs are the dominant source of myofibroblasts in liver fibrosis caused by different
etiology. At the same time, our data indicate that PFs and MCs partly contribute to
myofibroblasts by uniquely participating in biliary fibrosis and capsular fibrosis,
respectively. This study provided a new tool allowing the liver field to study three
different liver mesenchymal cells at the same time in liver fibrosis. In Chapter 4, we
158
provided clear evidence showing no differentiation of liver mesenchymal cells such as
HSCs into epithelial cells, including stem cells in liver injury and regeneration. Our data
solved a debate about whether HSCs are stem cells for liver epithelial cells and
emphasized the important role of mesenchymal cells in liver fibrosis. Lastly in Chapter 5,
besides to TGF-  signaling, we discovered that LPA acts as a profibrogenic factor and
induces MCs and HSCs differentiation into myofibroblasts through stimulating nuclear
localization of TAZ coactivator. Inhibition of LPA signaling was able to suppress liver
fibrosis in mouse liver injury models. These results provide a novel therapeutic target for
liver fibrosis. Taken together, our studies identified new cell sources of myofibroblast,
established unique tools for studying fibrosis, and purposed a novel signalings for anti-
fibrotic drug development.

159
6.2 Future directions
Although our studies provided breakthrough knowledge in accelerating the development
of anti-fibrotic drugs, further studies will be need to translate our data to clinical
applications. For example, TGF-  has been shown to induce MMT; however, it is unclear
how MCs sense injury signals from the environment. Elucidation of biological and
molecular changes involved in MC activation and fibrogenesis will assist the
development of novel approaches for prevention therapy for fibrosis. In addition to MCs
and HSCs, PFs can convert into myofibroblasts; other cells that are less characterized
might also contribute to fibrosis. Similarly, resident fibroblasts in the peritoneum and
liver capsule on the liver surface are suggested to be another source of myofibroblasts,
yet no lineage tracing tools are available to delineate their contribution. More CreERT2
mouse lines will be necessary to quantify their contribution to myofibroblasts in fibrosis.
By understanding the relative contribution of each cell type to myofibroblasts in fibrosis,
it will help scientist to design strategy targeting each cell type in fibrosis. Lastly, to
support the idea that LPA is a promising target for future clinical development, it is
necessary to understand molecular mechanisms in detail and generate more genetic
mouse models. It will also be interesting to understand the effect of LPA on other liver
cell types including hepatocytes, SECs and Kupffer cells and their roles in liver fibrosis.
We have gained extraordinary insights into the role of mesenchymal cells in organ
fibrosis. Our findings will help the development of new therapies against fibrosis one step
closer to clinical application.
160
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Abstract (if available)

Abstract Organ fibrosis is a worldwide health care problem as it leads to end-stage organ failure in patients. Currently, there is no medical treatment for organ fibrosis, such as in the liver and peritoneum. Myofibroblasts have been suggested to participate in fibrogenesis through synthesizing extracellular matrices and proinflammatory cytokines. Accumulating evidence suggest that there are multiple cellular sources of myofibroblasts. However, it remains elusive how different cell types contribute to myofibroblasts in organ fibrosis. Thus, identification of cell types responsible for fibrogenesis and understanding molecular mechanisms underlying generation of myofibroblasts are essential to determine therapeutic targets for suppression of fibrogenesis. Prolonged injury to the liver often results in liver fibrosis. Similarly, patients who undergo peritoneal dialysis for kidney failure also cause peritoneal fibrosis. In the present study, we focused on the liver and peritoneum as visceral and parietal organs in the peritoneal cavity and identified the cellular sources of myofibroblasts in liver and peritoneal fibrosis. Moreover, we examined molecular mechanisms underlying fibrogenesis in these organs. Mesothelial cells (MCs) form a single epithelial cell layer, called mesothelium, on the surface of the both organs in the peritoneal cavity. We found that MCs give rise to myofibroblasts in both organs. Although MCs partly contributed to myofibroblasts in peritoneal fibrosis, protection of the mesothelium ameliorated fibrosis. In liver fibrosis, hepatic stellate cells (HSCs) reside in the space of Disse and are known to differentiate into myofibroblasts. In addition, portal fibroblasts (PFs) around the bile duct are believed to be the major source of myofibroblasts in biliary fibrosis. However, contribution of PFs to myofibroblasts remains elusive due to insufficient availability of markers and isolation methods. We established a unique protocol to isolate MCs, HSCs, and PFs from normal or injured mouse livers and demonstrated that all three cell types convert into myofibroblasts and transforming growth factor-beta (TGF-beta) stimulates their differentiation. Our data indicate that HSCs are the dominant source of myofibroblasts in liver fibrosis caused by different etiology. Although PFs and MCs partly contributed to myofibroblasts in liver fibrosis, they uniquely participated in biliary fibrosis and capsular fibrosis, respectively. Lastly, we identified lysophosphatidic acid (LPA) as a profibrogenic factor in liver fibrosis. We found that LPA signaling stimulates nuclear localization of TAZ coactivator and induces differentiation of HSCs and MCs toward myofibroblasts. Treatment with an LPA inhibitor reduced liver fibrosis in mice. In conclusion, we demonstrated that MCs act as a source of myofibroblasts in liver and peritoneal fibrosis. In liver fibrosis, HSCs are the major source of myofibroblasts and both MCs and PFs uniquely contribute to fibrosis beneath the liver surface and around the bile duct, respectively. Our data indicate that LPA acts as a profibrogenic factor and its signaling pathway will be a new therapeutic target for suppression of liver fibrosis.

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Asset Metadata

Creator Lua, Ingrid Ahim (author)

Core Title Determination of the source of myofibroblasts and therapeutic strategy against fibrosis

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Genetic, Molecular and Cellular Biology

Publication Date 07/22/2018

Defense Date 06/02/2016

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag fibrosis,hepatic stellate cells,liver,mesothelial cells,OAI-PMH Harvest,peritoneal,portal fibroblasts

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Chuong, Cheng-Ming (committee chair), Asahina, Kinji (committee member), Tsukamoto, Hidekazu (committee member)

Creator Email ilua@usc.edu,ingoody@hotmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c40-274148

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Identifier etd-LuaIngridA-4588.pdf (filename),usctheses-c40-274148 (legacy record id)

Legacy Identifier etd-LuaIngridA-4588.pdf

Dmrecord 274148

Document Type Dissertation

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Rights Lua, Ingrid Ahim

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Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

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