DISS. ETH Nr. 18687
Identification of molecular mechanisms
controlling lymphatic vessel formation by use of
the embryonic stem cell-derived embryoid body
assay
A dissertation submitted to
ETH ZURICH
for the degree of
Doctor of Sciences
presented by
Daniela Marino
Laurea Magistrale, Faculty of Biotechnology, University of Milan, Italy
Born November 17, 1981
Citizen of Italy
Accepted on the recommendation of
Prof. Dr. Michael Detmar
Prof. Dr. Cornelia Halin-Winter
2009
"Fatti non foste a viver come bruti,
ma per seguir virtute e canoscenza"
“You were not born to live like animals
but to pursue virtue and possess knowledge”
(Dante Alighieri, Divina Commedia,
Inferno canto XXVI, 116-120)
Table of contents
1
SUMMARY
1.1
1.2
2
7
Summary
Riassunto
7
10
INTRODUCTION
13
2.1
The lymphatic vascular system: Embryonic development and role in health and disease
2.1.1
2.1.2
2.1.3
2.1.4
2.1.5
2.1.6
2.1.7
2.2
The lymphatic system history
Concepts of lymphatic vascular system development
Mechanisms of mammalian lymphatic vascular development
Lymphatic vascular development in Xenopus Laevis
Molecular markers of lymphatic endothelium
Molecular mediators of adult lymphangiogenesis
Involvement of lymphatic vessels in disease
13
16
17
18
28
31
33
35
Mouse embryonic stem cell-derived embryoid bodies
2.2.1
2.2.2
42
EB culture methods
44
Vasculogenesis, blood and lymphatic vascular development and (anti-) lymphangiogenesis research
45
3
AIM AND OUTLINE OF THIS DISSERTATION
48
4
IDENTIFICATION OF MOLECULAR MECHANISMS CONTROLLING LYMPHATIC
VESSEL FORMATION
49
4.1
A role for all-trans-retinoic acid in the early steps of lymphatic vasculature development 49
4.1.1
4.1.2
4.1.3
4.1.4
4.1.5
4.2
49
50
52
58
73
Activation of the epidermal growth factor receptor promotes lymphangiogenesis in vitro
and in vivo
78
4.2.1
4.2.2
4.2.3
4.2.4
4.2.5
5
6
7
8
Abstract
Introduction
Materials and methods
Results
Discussion
Abstract
Introduction
Materials and methods
Results
Discussion
CONCLUSIONS AND OUTLOOK
ACKNOWLEDGMENTS
REFERENCES
CURRICULUM VITAE
78
79
80
86
98
103
106
108
124
1 SUMMARY
1.1
Summary
The lymphatic vascular system, the second vascular system present in vertebrates, has emerged
as a fundamental player in physiological, as well as pathological, processes. It maintains tissue
fluid balance, contributes to immune cell trafficking and absorbs lipids in the intestine. Recent
scientific findings have defined its role in lymphedema, obesity, inflammation, tumor metastasis
and hypertension. Considerable progress has been made in understanding the major mechanisms
controlling the formation of the lymphatic vascular system during development as well as
lymphatic vessel remodeling (lymphangiogenesis) in the adult organism. Nevertheless, further
studies are needed to develop therapeutic strategies for the effective control of lymphatic vessel
growth or regression in pathological conditions.
During embryogenesis, the lymphatic endothelial cells arise from the anterior cardinal veins of
the embryo and form lymphatic vessels in a stepwise manner that includes acquisition of
lymphatic competence and commitment, budding and sprouting, lymph sac formation and
vessel maturation. Analyses of genetically modified animal models have characterized the later
phases of lymphatic development; however, the processes that regulate the earliest steps remain
unknown. The first part of the present work aimed at unravelling the molecular mechanisms
controlling the earliest steps of lymphatic competence and commitment. Using mouse
embryonic stem cell-derived embryoid bodies (EBs), an in vitro model that largely mimics
mouse embryogenesis, we identified all-trans-retinoic acid (RA) as a novel regulator of the
expression of the lymphatic vessel endothelial receptor (LYVE)-1 (lymphatic competence) and
the transcription factor Prox1 (lymphatic commitment) by venous endothelial cells. The studies
also revealed a synergistic effect of cAMP on the induction of LYVE-1/Prox1 expression by
7
RA. The use of antagonist molecules and immunohistochemical stainings indicated that retinoic
acid receptor (RAR)-α and cAMP-dependent protein kinase (Pk)-A are probable mediators of
the RA/cAMP activity in the lymphatic differentiation processes. Moreover, in vivo studies
revealed that exposure of mouse and Xenopus embryos to RA upregulated LYVE-1 and Prox1
expression in the endothelial cells of the cardinal veins and the developing primary lymph sacs.
In contrast, in utero exposure of mouse embryos to the RAR-α antagonist, Ro 41-5253,
decreased LYVE-1 and Prox1 expression in the same vascular regions. Taken together, these
findings indicate that RA regulates the earliest steps of lymphatic vasculature development.
Interestingly, intraperitoneal injection of RA in adult mice resulted in increased dermal
lymphatic vessel growth; thus, investigating the role of RA in adult lymphangiogenesis might
lead to the development of novel therapeutic strategies for e.g. lymphedema.
The second part of this thesis aimed at using the EB model to identify novel lymphangiogenic
pathways. To this end, EBs were treated with small organic molecules with well-defined
pathway specificities and the formation of lymphatic vessels was evaluated. The screening
identified the small molecule GW2974 (N4-(1-benzyl-1H-indazol-5-yl)-N6,N6-dimethylpyrido[3,4-d]pyrimidine-4,6-diamine), which targets the tyrosine kinase associated with
epidermal growth factor receptor (EGFR/ErbB2), as an inhibitor of lymphatic vessel formation
in the EBs. Conversely, EBs treated with EGF, an activating ligand of EGFR/ErbB2, displayed
increased number of lymphatic vessel networks as compared to control EBs. We found that
human dermal lymphatic endothelial cells express low levels of EGFR; GW2974, as well as
EGFR/ErbB2 blocking antibodies, inhibited in vitro migration and tube formation of these cells
whereas EGF significantly induced those processes. In vivo, mice subcutaneously implanted
with matrigel plugs containing EGF displayed increased lymphatic vessel size and density in the
plug-associated skin. Moreover, analysis of skin from mice expressing amphiregulin (another
EGFR ligand) under control of the keratin 14 promoter revealed more and larger dermal
lymphatic vessels. These findings suggest that EGFR signalling has a role in lymphatic vessel
8
formation and that targeting EGFR might represent a novel strategy to interfere with
lymphangiogenesis in pathological conditions, such as tumor metastasis.
Collectively, our results identify an important unexpected role for retinoic acid in lymphatic
vascular system development and a novel function for EGFR signalling in lymphangiogenesis.
9
1.2
Riassunto
Il sistema linfatico, il secondo sistema vascolare presente nei vertebrati, si è rivelato essenziale
sia in processi fisiologici che patologici. Esso, infatti, mantiene l’equilibrio dei fluidi tissutali,
contribuisce al trasporto delle cellule immunitarie e assorbe i lipidi nell’intestino. Recenti
ricerche scientifiche, inoltre, hanno riconosciuto il ruolo di questo sistema vascolare in patologie
quali linfedema, obesità, infiammazione, metastasi tumorale ed ipertensione. Nonostante sia
stato raggiunto un progresso straordinario nella comprensione dei meccanismi principali che
controllano la formazione del sistema linfatico durante lo sviluppo embrionale così come la
crescita dei vasi linfatici nell’organismo adulto (linfangiogenesi), ulteriori studi sono necessari
per sviluppare strategie terapeutiche che possano efficacemente controllare la crescita o la
regressione dei vasi linfatici in situazioni patologiche.
In particolare, durante lo sviluppo embrionale, le cellule endoteliali linfatiche originano dalle
vene cardinali anteriori dell’embrione e formano i vasi linfatici in un processo a stadi che
include l’acquisizione della competenza e della specificazione linfatica, migrazione, formazione
dei sacchi linfatici e maturazione dei vasi. L’analisi di modelli animali geneticamente modificati
ha contribuito a caratterizzare le fasi finali dello sviluppo linfatico, d’altrocanto però, i processi
che regolano le fasi iniziali rimangono, a tutt’oggi, sconosciuti. La prima parte del presente
lavoro ha avuto come scopo l’identificazione dei processi molecolari che controllano
l’acquisizione della competenza e della specificazione linfatica. Mediante l’uso di corpi
embrionici ottenuti da cellule staminali embrionali di topo (EBs), modello in vitro, questo, che
ampiamente mima l’embriogenesi di topo in vivo, abbiamo scoperto come l’acido retinoico
(RA) regoli l’espressione del gene LYVE-1 (competenza linfatica) e del gene Prox1
(specificazione linfatica) nelle cellule endoteliali venose. Questi studi in vitro, hanno inoltre
evidenziato un effetto sinergistico di cAMP nell’ attivazione dell’espressione di LYVE-1/Prox1
10
da parte di RA. L’utilizzo di molecole antagoniste e di colorazioni immunoistochimiche hanno
suggerito che il recettore per l’acido retinoico (RAR)-α e la proteina chinasi dipendente da
cAMP (PkA) potrebbero essere i mediatori dell’attività di RA/cAMP nei processi di
differenziamento linfatico. Inoltre, studi in vivo, hanno rivelato che l’esposizione all’RA di
embrioni di topo e di rana (Xenopus laevis) ha indotto una maggiore espressione di LYVE-1 e
Prox1 nelle cellule endoteliali delle vene cardinali e dei nascenti sacchi linfatici. Al contrario,
l’esposizione in utero all’antagonista del RAR-α, Ro 41-5253, ha diminuito l’espressione di
LYVE-1 e Prox1 nelle stesse regioni vascolari. Riassumendo, i nostri risultati indicano come
RA regoli le fasi iniziali dello sviluppo del sistema vascolare linfatico. Inoltre, è interessante
notare come un’ iniezione intraperitoneale di RA in topi adulti, abbia accresciuto la formazione
dei vasi linfatici (risultati non mostrati); quindi, ulteriori studi sul ruolo di RA nella
linfangiogenesi adulta potrebbe contribuire allo sviluppo di nuove strategie terapeutiche per
poter, per esempio, curare il linfedema.
La seconda parte di questa tesi, ha avuto lo scopo di identificare nuovi pathway linfangiogenici,
mediate l’utilizzo dei modelli EB. Per questo, gli EBs sono stati prima trattati con piccole
molecole organiche aventi specificità per definiti pathway molecolari e poi analizzati in termini
di crescita di vasi linfatici. Lo screening, ha rivelato come la molecola GW2974 (N4-(1-benzyl1H-indazol-5-yl)-N6,N6-dimethyl-pyrido[3,4-d]pyrimidine-4,6-diamine), che ha come target la
tirosin chinasi associata al recettore del fattore di crescita epidermale (EGFR-ErbB2), agisca da
inibitore della formazione di vasi linfatici negli EBs. Al contrario, EBs trattati con EGF, un
ligando che attiva EGFR-ErbB2, hanno presentato un numero maggiore di reti linfatiche rispetto
agli EB non trattati. Inoltre, abbiamo scoperto che le cellule endoteliali linfatiche umane
esprimono bassi livelli di EGFR. GW2974, così come anticorpi che bloccano l’EGFR-ErbB2,
hanno inibito la migrazione e la formazione di vasi in vitro di queste cellule, mentre EGF le ha
indotte in modo significativo. In vivo, topi a cui è stato impiantato matrigel contenente EGF,
hanno mostrato un incremento di misura e densità dei vasi linfatici nella pelle in contatto con il
11
gel. Inoltre, l’analisi di campioni di pelle prelevati da topi transgenici che overesprimono
l’anfiregulina (un altro ligando per EGFR-ErbB2) sotto il controllo del promotore del gene per
la keratina 14, ha rivelato più e più grandi vasi linfatici dermali. Queste scoperte suggeriscono
come l’EGFR-ErbB2 abbia un ruolo nella formazione dei vasi linfatici e come il targeting
dell’EGFR-ErbB2 possa rappresentare una nuova strategia per interferire con la linfangiogensi
in condizioni patologiche quali, ad esempio, la metastasi tumorale. Riassumento, i nostri
risultati identificano un importante inaspettato ruolo dell’acido retinoico nello sviluppo dei vasi
linfatici durante l’embriogenesi e una nuova funzione per l’EGFR-ErbB2 nella linfangiogenesi.
12
2 INTRODUCTION
2.1
The lymphatic vascular system: Embryonic development
and role in health and disease
Apart from the cardiovascular system, vertebrates also possess a lymphatic system that consists
of lymphatic vessels and lymphoid organs (lymph nodes, spleen, thymus, tonsils, Peyer’s
patches and lymphoid tissues associated with the respiratory systems). The organization and
function of the two vascular systems are distinct. The cardiovascular system is a closed
circulatory system with a central pump, the heart that distributes blood cells, oxygen, nutrients,
and hormones to all tissues. In contrast, the lymphatic vasculature is a one-way system that
removes fluid, cells, macromolecules, microbes and other substances from interstitial spaces
without a central driving force.
The lymphatic vascular system consists of blind-ended capillaries that collect and transport
extravasated protein-rich fluid that originates from blood serum, the lymph, from tissues to
precollector lymphatic vessels (Sacchi et al. 1997) then to collecting lymphatic vessels that
converge into lymphatic trunks to finally return the lymph to the venous circulation via the
thoracic duct through the connection with jugular and subclavian veins (Casley-Smith 1980). In
addition, in the intestine, lacteal lymphatic vessels, which are found in the villi, absorb and
transport vitamins and dietary fat released in form of lipid particles called chylomicrons (Jurisic
and Detmar 2009). Furthermore, the lymphatic system carries antigens and antigen-presenting
cells from the interstitium to the lymph nodes where they are presented to B and T cells (Jurisic
and Detmar 2009). Recently, interest in basic lymphatic research has been enhanced by
evidence demonstrating that the lymphatic system is involved in a number of human
13
pathological conditions, such as lymphedema, inflammation, cancer metastasis (Cueni and
Detmar 2008) and hypertension (Machnik et al. 2009). The lymphatic vasculature is present in
the skin and in several internal organs, with the exception of the epidermis, nails, hairs,
cartilage, cornea, brain, bone marrow and retina. Lymphatic capillaries are lined by a single
layer of overlapping lymphatic endothelial cells (LECs) and lack a continuous basement
membrane (Figure 2.1.1) as well as pericyte or smooth muscle cell coverage. LECs of the
capillaries are oak leaf shaped, inter-connected by button-like junctions and linked to the
extracellular matrix by anchoring elastic fibers (Leak and Burke 1968; Gerli et al. 1991; Baluk
et al. 2007) (Figure 2.1.1). Under high interstitial pressure conditions, the junctions open via
anchoring fibrillin filaments stretching, thus allowing fluid, macromolecules and cells to move
into the vessel lumen. Overlapping endothelial cell-cell contacts, or primary valves, prevent
fluid from escaping the capillaries (Trzewik et al. 2001; Schmid-Schonbein 2003). Lymphatic
capillaries then merge into pre-collector vessels (still presenting anchoring filaments in some
areas) that, together with downstream collecting lymphatic vessels, present zipper-like
junctions, a basement membrane and smooth muscle cells (SMCs) (Scavelli et al. 2004). The
contractile activity of the SMCs and of the skeletal muscles, together with the respiratory
movement and the presence of intraluminal valves, promote lymph flow (Von der Weid and
Zawieja 2004).
14
Figure 2.1.1 Structure of lymphatic vessels (Karpanen and Alitalo 2008). In contrast to blood vessels,
lymphatic vessels have thin walls and a relatively wide lumen. The endothelial cells of lymphatic capillaries
(green) partly overlap, forming valve-like openings, which allow easy access for fluid, macromolecules, and cells
into the vessel lumen. Lymphatic capillaries lack smooth muscle cells (SMCs; red) and have no basement
membrane. Anchoring filaments connect lymphatic capillary endothelial cells to the surrounding extracellular
matrix. The lymph drains from the lymphatic capillaries to precollecting and collecting lymphatic vessels, which
are finally emptied into veins in the jugular region. The precollecting and collecting lymphatic vessels have a
basement membrane, are surrounded by vascular smooth muscle cells and contain valves that prevent backflow of
the lymph. On its way, the lymph is filtered through a series of lymph nodes. In contrast, the endothelial cells of
blood vessels have a distinct basement membrane and are surrounded by pericytes/vSMCs, which form one or
multiple layers increasing in thickness with vessel size.
15
2.1.1
The lymphatic system history
Scientific advances in the 17th century were the result of a new concept of science that coupled
experimentation and innovation. However, in the case of human anatomy, physicians and
scientists were limited to observation alone. Most of the descriptions and theories of human
anatomy were established by the Greek physician Galen in the second century. Galen developed
theories on the physiology of human blood circulation, but the role of the lymphatic system was
not discussed despite the fact that earlier the Greek anatomist Erasistratus (ca. 304-ca.250
B.C.E.) had described the presence of vessels containing “white blood” in human cadavers
(Lord 1968; Vegetti 1995). Finally, on 23rd July 1622, the Italian physician Gaspare Aselli
(1581-1626), while studying the digestive system of a dog, noted that some glands and
capillaries contained a cloudy fluid. In his book “De lactibus sive lacteis venis” ((Asellius 1627)
the first anatomical report in color, Figure 2.1.2), Aselli incorrectly concluded that these glands
were associated with the blood circulation and that the content was transported by “milky veins”
to the liver (at that time, still considered the center of the venous system).
Figure 2.1.2 De lactibus sive lacteis venis. In 1622, the Italian physician Gaspare Aselli, while studying the
digestive tract of a dog (A), noted glands and capillaries containing a milky fluid (white lines). He reported his
findings in the De lactibus sive lacteis venis (1627, B): the first anatomical report in color (C).
16
Aselli’s observation inspired other scientists, like the French Jean Pecquet (1622-1674) who, in
a dog, found a lumbar reservoir draining the lacteals, now known as the cysterna chyli. The
Danish physician Thomas Bartholin (1616-1680) was the first to describe the lymphatic system
in humans (Eales 1974); in 1652 he published the “De Lacteis Thoracsis” where he described
the anatomy of the thoracic duct and recognised the lacteal veins as a second vascular system
within the body that he named the lymphatic system. Years later William Hunter (1718-1713)
and Alexander Monro (1733-1817) described the absorbent nature of the lymphatic vessels
draining fluid from the tissue and bringing it back to the blood circulation (Eales 1974).
Although the description of the anatomy of the lymphatic vascular system was progressing
rapidly, its embryonic origin remained unclear for a long time until Florence R. Sabin (19021904) and F. Lewis (1905) postulated the “centrifugal theory” proposing that LECs derive from
the embryonic venous endothelium (Sabin 1902). An alternative “centripetal theory” postulated
by Huntington and Mc Clure in 1910 suggested that LECs arise from mesenchymal progenitor
cells, a lymphangioblast (Huntington and McClure 1910). These two opposing theories have
been the center of a century-old debate that nowadays seems to have been partially resolved.
2.1.2
Concepts of lymphatic vascular system development
The centrifugal theory proposes that in the developing embryo endothelial cells bud out from
the anterior cardinal veins, proliferate, migrate and form primary jugular lymph sacs (Oliver
2004). From the sacs, the endothelial cells sprout and form vessel networks throughout the
body. Several recent studies, using genetically engineered mouse models (Wigle et al. 2002),
lineage tracing experiments (Srinivasan et al. 2007) and in vivo imaging studies in zebrafish
(Yaniv et al. 2006) have demonstrated that lymphatic endothelial cells of venous origin are the
main source for the developing lymphatic vasculature. However, a dual origin from both
embryonic veins and from a mesenchymal ancestor has also been proposed. Studies in chicken
17
embryos, suggest that the superficial parts of the jugular lymph sacs, as well as dermal
lymphatics, are derived from local lymphangioblasts (Wilting et al. 2006). In Xenopus tadpoles,
mesodermal precursor cells contribute to lymphatic vessel formation (Ny et al. 2005).
Furthermore, scattered mesenchymal cells, co-expressing lymphatic and leukocyte markers
were detected in mouse embryos in the jugular regions (Buttler et al. 2006), and cells coexpressing lymphatic and macrophage markers have been suggested to integrate into lymphatic
vessels (Buttler et al. 2008).
2.1.3
2.1.3.1
Mechanisms of mammalian lymphatic vascular development
Lymphatic competence
In the mouse embryo, the lymphatic vascular system begins to form after the establishment of
the intra-embryonic blood vasculature (Figure 2.1.3) In particular, at embryonic day (ED) 9,
endothelial cells lining the anterior cardinal veins start to express lymphatic endothelial
hyaluronan receptor-1 (LYVE-1) indicating lymphatic competence, the first step of lymphatic
differentiation. It has been proposed that the acquisition of competence, prepares the endothelial
cells to receive further lymphatic specification signals (Wigle et al. 2002). LYVE-1, an
homologue of the blood vascular endothelium-specific hyaluronan receptor CD44 (Banerji et al.
1999), is a member of the link protein superfamily and was identified as a cell surface protein of
lymphatic endothelial cells (Banerji et al. 1999; Jackson et al. 2001; Prevo et al. 2001). The
molecular mechanisms that control LYVE-1 expression have not been identified; however, a
recent study has shown that the chicken ovalbumin upstream promoter-transcription factor
(COUP-TF)-II regulates its expression in human cultured LECs (Lee et al. 2009). The function
of LYVE-1 is still unclear; LYVE-1 knockout mice do not display any lymphatic vascular
18
defects (Gale et al. 2007). However, mice deficient for the cell surface retention binding
protein-1 (CRSBP-1), a gene that is identical to LYVE-1, exhibit identifiable morphological and
functional alterations of lymphatic capillary vessels in certain tissues, such as the intestine,
marked by the constitutively increased interstitial-lymphatic flow and lack of typical irregularlyshaped lumens (Huang et al. 2006). In the adult, LYVE-1 expression is maintained by the LECs
and it is one of the most used markers for lymphatic vessels. Its expression is also found in a
subset of macrophages (Maruyama et al. 2005; Schledzewski et al. 2006) and in liver sinusoids
(Mouta Carreira et al. 2001; Nonaka et al. 2007).
2.1.3.2
Lymphatic commitment
At ED 9.5, polarized expression of the transcription factor prospero-related homeobox (Prox)-1
is detected on one side of the anterior cardinal veins, thus conferring lymphatic commitment to
the LYVE-1+, lymphatic-competent endothelial cells (Figure 2.1.3). Prox1 is the most specific
lineage marker for lymphatic endothelium (Oliver and Detmar 2002) and it controls the initial
steps of lymphatic development. Inactivation of Prox1 in mice results in complete absence of
lymphatic vascular structures (Wigle et al. 2002) and in utero death at around ED 14.5. In the
absence of Prox1, endothelial cells that have budded from the cardinal veins continue to express
blood vascular endothelial cell (BECs) lineage-specific genes. Conversely, ectopic expression of
Prox1 in human BECs in vitro induces expression of lymphatic vascular signature genes (Hong
et al. 2002; Petrova et al. 2002; Wigle et al. 2002), such as VEGFR-3 (Petrova et al. 2002),
integrin α9 (Mishima et al. 2007) and FGFR-3 (Shin et al. 2006). siRNA knock down of Prox1
in LECs downregulates VEGFR-3, podoplanin and FGFR-3 and upregulates the BEC signature
genes neuropilin-1, ICAM (inter-cellular adhesion molecule 1) and MCP1 (monocyte
chemotactic protein-1) (Lee et al. 2009). At present the signals that induce Prox1 expression in a
restricted subpopulation of the embryonic cardinal veins are not well understood. Recently, it
19
has been shown that vessels in mice deficient of SOX18, a transcription factor that belongs to
the SRY-relates HMG domain family, do not undergo definitive lymphatic differentiation.
Furthermore, SOX18 is expressed by the endothelial cells lining the dorsolateral sector of the
cardinal veins as early as ED 9, before Prox1 expression has begun. SOX18 expression remains
in the budding and jugular lymph sac forming lymphatic progenitor cells but it is not detectable
during lymphatic vascular system maturation or in the adult lymphatic vessels. These findings
suggest that SOX18 is not required for maintenance of LECs once they are fully differentiated.
Studies on the Prox1 promoter show that SOX18 directly activates this lymphatic master gene
(Francois et al. 2008). In addition, COUP-TF II physically interacts with Prox1 to form a stable
complex in various cell types including LECs (Lee et al. 2009). COUP-TF II is thought to
function as a coregulator of Prox1 and to be required, along with Prox1, to maintain the LEC
phenotype. Interestingly, COUP-TF II expression has been found on CD31+/Prox1+/LYVE-1+
endothelial cells of embryonic cardinal veins and on dermal adult lymphatic vessels in vitro and
in vivo (Lee et al. 2009). Mice lacking COUP-TF II die in utero around ED 10 and present head
and heart defects, hemorrhages, edema, enlarged blood vessels and malformed cardinal veins; it
still remains to be elucidated whether the presence of COUP-TF II is essential for lymphatic
differentiation.
2.1.3.3
Budding, migration and proliferation of LECs
Starting from ED 10.5, lymphatic committed embryonic endothelial cells response to vascular
endothelial growth factor (VEGF)-C, a ligand for VEGFR-3 and start to bud out from the
cardinal veins, proliferate, migrate and form the primary jugular lymph sacs (Tammela et al.
2005; Karpanen and Alitalo 2008) (Figure 2.1.3). During embryonic development, VEGFR-3 is
expressed by all endothelial cells and it is important for blood vessel remodeling. Indeed,
VEGFR-3 null mice die in utero at ED 9.5 due to severe cardiovascular defects (Dumont et al.
20
1998; Hamada et al. 2000) while VEGF-C null mice do not develop a lymphatic vasculature and
die in utero at ED 17-19 (Karkkainen et al. 2004). VEGF-C is expressed by mesenchymal cells
in the jugular area of developing embryos. In its absence, the committed Prox1+ endothelial
cells arise normally in the cardinal veins but fail to sprout and migrate. VEGFR-3 has another
known ligand, VEGF-D; studies in VEGF-D deficient mice have demonstrated that this protein
is not essential for lymphatic vasculature development (Karkkainen et al. 2004; Baldwin et al.
2005). Recently, integrins have been shown to be involved in lymphatic vascular system
development; mice deficient for integrin α9 β1 die soon after birth because of chylothorax
(Huang et al. 2000). Integrin α9 β1 is a receptor for VEGF-C and D (Vlahakis et al. 2005) and
its stimulation, by extracellular matrix (ECM) components, induces tyrosine phosphorylation of
VEGFR-3 (Wang et al. 2001). Since VEGF-C and -D null mice, unlike to the VEGFR-3 null,
have no blood vascular phenotype it is thought that integrins contribute to VEGFR-3 signalling
in blood vessel remodeling (Haiko et al. 2008). Furthermore, VEGFR-3 interacts with
neuropilin-2 (NRP2) (Karpanen et al. 2006) and mice lacking NRP2, a receptor for class III
semaphorins, have abnormal lymphatic capillaries, while large vessel develop normally (Yuan
et al. 2002).
2.1.3.4
Sprouting from the lymphatic plexus
Starting from ED 11.5, the LECs that formed the jugular lymph sac proliferate and sprout to
expand the primary lymphatic plexuses (Figure 2.1.3). Recently, adrenomedullin has been
shown to play a role in this process. Adrenomedullin is a multifunctional vasodilator peptide
that transduces its effects through the calcitonin receptor-like receptor (calcrl) when the receptor
is associated with a receptor activity-modifying protein (RAMP)-2. Adrenomedullin-, calcrl-, or
RAMP2-null mice die during mid-gestation after developing interstitial lymphedema. A calcrl
conditional knockout in endothelial cells resulted in abnormal jugular lymphatic vessels due to
21
impaired lymphatic endothelial cell proliferation (Fritz-Six et al. 2008). In addition, the
transcription factor NFATc1 (nuclear factor of activated T-cells-1), which is critical for lineage
selection in T-cell differentiation, cardiac valve morphogenesis and osteoclastogenesis, has been
shown to also have a role in lymphatic development and patterning. NFAT is colocalized with
the lymphatic markers Prox1, VEGFR-3 and podoplanin on cardinal vein endothelial cells as
LECs are committed. In both NFATc1 null and in utero treated embryos with the calcineurin,
(which is required for NFAT activation) inhibitor cyclosporine-A, Prox1, VEGFR-3 and
podoplanin positive endothelial cells budded from the cardinal veins at E11.5, but poorly
sprouted off the jugular lymph sacs to form the lymphatic plexus. These results, thus, suggest a
role for calcineurin-NFAT signalling in embryonic lymphatic patterning (Kulkarni et al. 2009).
2.1.3.5
Separation of blood and lymphatic vessels
Later in development (ED13.5-14.5), the first lymphatic vessels completely separate from the
blood vasculature (Figure 2.1.3). The tyrosine kinase, Syk, and the adaptor protein, SLP76, that
are almost exclusively expressed by hematopoietic cells are fundamental for this separation
process. Mice lacking Syk/Slp76 die perinatally due to haemorrhaging (Abtahian et al. 2003;
Sebzda et al. 2006). Recently, phospolipase gamma (PLCγ)-2, which is a downstream target of
Syk/SLP76, has been shown to play an essential role in initiating and maintaining the separation
of the blood and lymphatic vasculature (Ichise et al. 2009). Podoplanin, a small transmembrane
mucin-like protein, is highly expressed in lymphatic endothelial cells starting at ED 11.
Podoplanin null mice die at birth because of abnormal lung development and display impaired
lymphatic transport, dilation of lymphatic vessels and congenital lymphedema (Schacht et al.
2003). The precise molecular function of podoplanin has not been yet identified. However,
podoplanin is known to be involved in cytoskeletal organization of endothelial and tumor cells
(Schacht et al. 2003; Martin-Villar et al. 2006; Wicki et al. 2006) and to aggregate platelets via
22
interaction with the C-type lectin-like receptor (CLEC)-2 (Kato et al. 2003; Suzuki-Inoue et al.
2007) which signals via Syk and SLP76 (Suzuki-Inoue et al. 2007). Finally, a recent study
reported blood-filled lymphatic vessel phenotype in mice lacking, or with reduced, podoplanin
expression (Fu et al. 2008). Thus, given the phenotypes of Syk/SLP76, PLCγ2 and podoplanin
null mice one could speculate on a possible function for podoplanin in forming thrombi to
prevent anastomoses between blood and lymphatic vessels. In addition, the phenotype of mice
null for sprouty-related ena/VASP homology domain-containing proteins (Spred) 1/2
(Taniguchi et al. 2007), two negative regulators of growth factor and ERK activation, resembles
that of Syk/SLP76 knockouts. Furthermore, studies on angiopoietin-like (Angptl)-4, have
suggested that organ-specific mechanisms are essential after birth to maintain blood and
lymphatic vessels separated. In fact, mice deficient for Angptl-4, display blood filled intestinal
lymphatics (Backhed et al. 2007).
2.1.3.6
Maturation of lymphatic vessel network
Primary lymphatic vessels undergo drastic changes to form a complex and organized network of
lymphatic capillaries and collectors (Figure 2.1.3). Foxc2 is a forkhead transcription factor that
is highly expressed in the developing lymphatic vasculature. Initial lymphatic vascular
development proceeds normally in absence of Foxc2. However, later on, collecting lymphatics
fail to develop valves, and capillaries acquire an excessive smooth muscle cell (SMC) coverage.
A similar phenotype has also been reported for mice lacking the PDZ domain of ephrinB2
(Makinen et al. 2005). Recently, it has been shown that Foxc2 controls lymphatic vessel
maturation through cooperation with NFATc1 (Norrmen et al. 2009). Angiopoietin (Ang)-2 is
also required for the proper development of the lymphatic vasculature. Ang2 -/- mice display
lymphatic vessel disorganization and hyperplasia, impaired recruitment of SMCs into collecting
lymphatic vessels, as well as irregularly patterned lymphatic capillaries (Gale et al. 2002).
23
Importantly, studies on integrins and on elastic microfibril-associated proteins such as emilin-1,
(elastin microfibril interfacer-1) revealed an importance for interactions between the LECs and
the ECM. Mice lacking integrin α9 β1 develop respiratory failure, caused by an accumulation of
large volumes of pleural fluid, and die between 6 and 12 days of age (Huang et al. 2000). In
emilin-1-/- mice lymphatic drainage is impaired due to disorganized lymphatic vessels with
reduced anchoring filaments (Danussi et al. 2008). A similar phenotype to the one observed in
integrin α9 β1 -/- mice, is reported also for Net, a transcription factor member of the ternary
complex factor family which is expressed in sites of vasculogenesis during mouse development
(Ayadi et al. 2001). Moreover, Aspp1, the apoptosis stimulating protein of p53, was recently,
found to be involved in lymphatic vessel assembly as mice lacking Aspp1 have a disorganized
lymphatic plexus and impaired lymphatic drainage in the embryonic skin. Interestingly, the
drainage defects are resolved in the adult animals (Hirashima et al. 2008). Furthermore, targeted
inactivation of neuropilin (Nrp)-2 results in absence or severe reduction of small lymphatic
vessels and capillaries during development (Yuan et al. 2002). Finally, mice generated with
mutations in the phospoinositide 3-kinase (pi3kca) gene display defective development of the
lymphatic vasculature, resulting in perinatal appearance of chylous ascites and reduced number
of lymphatic capillaries (Gupta et al. 2007).
All known factors involved in lymphatic vascular system development are listed in Table 2.1.1
24
25
Table 2.1.1 Genetic mouse models displaying a lymphatic phenotype
Genes
Function
Model Lethality Lymphatic phenotype
Prox11
Transcription
factor
KO
E14.5
LYVE-12
Hyaluronan
receptor
KO
Normal
No lymphatic vasculature (-/-), adultonset obesity, chylous ascites ( -/+ )
None or only subtle defects: increased
interstitial lymphatic flow, atypical
shape of vessel lumen
VEGF-C
Growth factor
KO
E17-19
No lymphatic vasculature (-/-), delayed
lymphatic vascular development,
lymphedema (-/+)
VEGFR-34
Growth factor
receptor
KO
E10
Cardiovascular failure, defective
remodeling of vascular networks
Syk/SLP-76 5
Tyrosine
kinase
KO
Perinatal
Abnormal blood-lymphatic
connections, chylous ascites, defect in
hematopoietic endothelial progenitors
Spred1/2 6
Neg. regulators
of ERK
Secreted
inhibitor of
lipoprotein
lipase
KO
E12.515.5
Lymphedema, dilated and blood filled
lymphatic vessels
KO
Postnatal lymphatic-venous
< 3 weeks partitioning defect in the small
intestine
3
Angpl4 7
Podoplanin 8a
Membrane
glycoprotein
KO
Perinatal
Foxc2 9
Transcription
factor
KO
E12.5Perinatal
EphrinB2 10
Ligand of
Mutant Perinatal
EphB receptors
Growth factor
receptor
KO
12
Growth factor
KO
Aspp1 13
p-53 binding
KO
Neuropilin-2
11
Angiopoietin-2
Lymphedema, dilatation and
mispatterning of lymphatic vessels,
diminished lymphatic transport. Blood
filled lymphatic vessels 8b
Lymphatic hyperplasia, retrograde
lymph flow ( -/+ ), abnormal patterning
and pericyte investment of lymphatic
vessels, absence of valves, lymphatic
dysfunction (-/-)
Defective remodeling of lymphatic
vascular network, hyperplasia, lack of
valves, chylothorax
Transient absence or severe reduction
Postnatal
of small lymphatic vessels and
(50%)
capillaries during development
Chylous ascites and subcutaneous
Perinatal edema, abnormal patterning of
lymphatic vessels
Subcutaneous edema, disorganized and
non-functional lymphatic vasculature
Normal
in embryo, mispatterned collecting
lymphatic vessels in adult
26
Adrenomedulli Vasoactive
n 14
peptide
KO
E14.5
Emilin-1 15
ECM protein
KO
Normal
Integrinα9 16
Adhesion
receptor
KO
Perinatal
PLCγ2 17
Phospholipase
NFATc1 18
Transcription
KO E12.5
factor
Transcription
KO Perinatal
factor
Transcription
KO E14.5
factor
Phosphoinositi
Postnatal
Mutant
de 3-kinase
(50%)
Net 19
Sox18 20
Pi3kca 21
Mutant Perinatal
Interstitial lymphedema (KO),
abnormal jugular lymphatic vessels
(conditional KO in ECs)
Hyperplastic, enlarged, irregularly
patterned lymphatic vessels, reduction
of anchoring filaments, lymphedema
Chylothorax, lymphedema
Abnormal blood-lymphatic
connections, blood filled lymphatic
vessels
Abnormal formation of jugular lymph
sac
Chylothorax, dilated lymphatic vessels
Edema, chylous ascites, cardiovascular
and hair follicle defects
Chylous ascites, reduction of
lymphatic capillaries
VEGF-C 22
Growth factor
Tg/K14
Hyperplastic lymphatic vessels
VEGF-D 23
Growth factor
Tg/K14
Hyperplastic lymphatic vessels
24
Growth factor
Tg/K14
Enlarged lymphatic vessels
25
Growth factor
Tg/K14
Lymphatic vessel enlargement and
sprouting
HGF 26
Growth factor
Tg
VEGF-A
Angiopoietin-1
Increased number and enlargement of
lymphatic vessels
1
(Wigle et al. 2002; Harvey et al. 2005). 2(Huang et al. 2006; Gale et al. 2007).
(Karkkainen et al. 2004). 4(Dumont et al. 1998). 5(Abtahian et al. 2003; Sebzda et al.
2006). 6(Taniguchi et al. 2007). 7(Backhed et al. 2007). 8a(Schacht et al. 2003). 8b(Fu et al.
2008) 9(Petrova et al. 2004; Seo et al. 2006). 10(Makinen et al. 2005). 11(Yuan et al. 2002).
12
(Gale et al. 2002; Shimoda et al. 2007). 13(Hirashima et al. 2008). 14(Fritz-Six et al. 2008).
15
(Danussi et al. 2008). 16(Huang et al. 2000). 17(Ichise et al. 2009). 18(Kulkarni et al. 2009).
19
(Ayadi et al. 2001). 20(Francois et al. 2008). 21(Gupta et al. 2007). 22(Jeltsch et al. 1997).
23
(Veikkola et al. 2001). 24(Kunstfeld et al. 2004). 25(Tammela et al. 2005). 26(Kajiya et al.
2005).
3
Angptl4, angiopoietin-like protein 4; Aspp, apoptosis stimulating protein of p53; ECM, extracellular
matrix; Foxc, forkhead box C; HGF, hepatocyte growth factor; K14, keratin 14; KO, knockout;
LYVE-1, lymphatic vascular endothelial hyaluronan receptor-1; NRP, neuropilin; PDZ, PSD-95,
DISCS-large, and ZO-1; SLP, Src homology 2-domain containing leukocyte protein; SOX, sex
determining region Y-related high mobility group box; Spred, Sprouty-related Ena/VASP
homology 1 domain-containing protein; Tg, transgenic; VEGF, vascular endothelial growth factor;
VEGFR, vascular endothelial growth factor receptor.
27
2.1.4
Lymphatic vascular development in Xenopus Laevis
Amphibians have a common evolutionary history with mammals that is an estimated 100
million years longer than that between zebrafish and mammals. Being both tetrapods,
amphibians and mammals share extensive synteny at the level of the genomes and have many
similarities in organ development, anatomy, and physiology (Christensen et al. 2008; Raciti et
al. 2008). In the past, embryos and tadpoles of the African clawed frog (Xenopus laevis) have
served as a powerful animal model to study blood vascular development and angiogenesis
(Cleaver and Krieg 1998; Helbling et al. 2000; Levine et al. 2003; Kalin et al. 2007). More
recently, Xenopus embryos were shown to develop also a complex, well-defined lymphatic
vascular system (Ny et al. 2005). Similar to the development of the mammalian lymphatic
vascular system, in Xenopus, LECs transdifferentiate from venous cells BEC and
lymphangioblasts contribute to newly forming lymph vessels that mature to drain fluids from
the peripheral tissues back to the blood circulation. The lymphatic vascular system of the
Xenopus, in contrast to the mammalian one, presents a lymph heart.
2.1.4.1
Rostral lymph sac, lymph heart and caudal lymphatic vessel formation
Prox1 expression is first found in the head from stage 28 on. Later on, cells of the lateral plate
mesoderm co-express Prox1, VEGFR-3 (that is expressed in the whole developing vasculature)
and angioblast markers (e.g mesenchyme-associated serpentine receptor (Msr)). Once the
Prox1+ cells became restricted to the subectodermal regions, Msr expression is lost and the
Prox1+/VEGFR-3+/Msr- population is now defined as lymphangioblasts; these cells will
assemble and form the rostral lymph sac (Ny et al. 2005).
Similar to the mammalian system, at stage 32, a subpopulation of the venous endothelial cells in
the anterior trunk start expressing Prox1 and progressively lose Msr expression; they
28
transdifferentiate into the lymphatic lineage and form the lymph heart. Furthermore, Prox1
expression is detected in the posterior trunk. Prox1+ lymphatic endothelial cells lining the
posterior cardinal veins detach from it and form the ventral caudal lymph vessel. Sometimes
even before the posterior cardinal veins have acquired a lumen, Prox1+ endothelial cell clusters,
the putative lymphangioblast, appear. At stage 33/34, the expression of LYVE-1 is weakly
detected in all vessels. At stage 35, VEGFR-3 expression begins to be restricted to the
lymphatic vessels while LYVE-1 expression is restricted to the lymphatic vessel only from
stage 39. From stages 37/38 onward, endothelial cells of the dorsal longitudinal anastomosing
vessel, a vein draining blood to the heart, were also found to transdifferentiate into Prox1+ cells.
As a result, (by stage 42) a lumenized ventral and dorsal caudal vein are formed. Thus, venous
transdifferentiation and lymphangioblast differentiation contribute to the formation of the caudal
lymph vessels. A schematic representation of the early stages of lymphatic vascular system
development (Prox1 expression) in Xenopus tadpoles is depicted in Figure 2.1.4
Figure 2.1.4 Expression of Prox1 and Msr in early tadpoles (from Ny et al. 2005). (a-d) in situ hybridization
(blue) for Prox1 (left panel, lymphatic vessels) and Msr (righ panel, blood vessels). Prox1 is expressed in the head
where the rostral lymph sac develops (RLS); in the anterior trunk adjacent to the pronephric sinus (PNS), where the
lymph hearth forms (LH); and in the posterior trunk adjacent to the posterior cardinal vein (PVC) where the ventral
29
caudal lymph vessel (VCLV) and dorsal caudal lymph vessel (DCLV) form. Prox1 is also expressed in the eyes (E)
and liver diverticle (L). Later in development additional commitment sites emerge from the dorsal longitudinal
anastomosing vessel (DLAV). Msr is expressed in the dorsal aorta (DA), in the cardinal vein, in the intersomitic
veins (ISV), in the heart (H) and in the aortic arch artery (AAA). Scale bars: a-c, 1 mm; d, 500 µm.
2.1.4.2
Molecular mechanisms of the lymphatic vascular development in Xenopus
Antisense-morpholino knockdown studies of the lymphatic master gene Prox1 revealed that
lack of this transcription factor impairs lymphatic development (and not blood vasculature
development) resembling the Prox1-/- mouse phenotype (Ny et al. 2005). In addition, antisensemorpholino knockdown of the lymphangiogenic factor VEGF-C causes lymphatic vessel defects
similar to the phenotype observed in VEGF-C deficient mice, including impaired LEC sprouting
and migration, and the formation of lymphedema (Ny et al. 2005). Single morpholino antisense
knockdown of VEGF-D did not affect lymphatic commitment, but transiently impaired
lymphatic endothelial cell (LEC) migration. Notably, combined knockdown of VEGF-D with
VEGF-C resulted in more severe migration defects and lymphedema formation than the
corresponding single knockdowns. The stronger phenotype might be possibly due to the fact
that, in the single knockdowns, each factor might have compensated for the loss of the other.
Knockdown of VEGFR-3 or treatment with a VEGFR-3 inhibitor similarly impaired lymph
vessel formation and function and caused pronounced edema. VEGFR-3 silencing by
morpholino knockdown, inhibitor treatment, or VEGF-C/D double knockdown also resulted in
dilation and dysfunction of the lymph heart. These findings document a critical role of VEGFR3 in embryonic lymphatic development and function and reveal a previously unrecognized
modifier (VEGF-D null mice have no lymphatic phenotype (Ny et al. 2008)) role of VEGF-D in
the regulation of embryonic lymphangiogenesis in frog embryos (Ny et al. 2008). Recently,
using a genetic screen in zebrafish, ccbe1 (collagen and calcium-binding EGF domain-1) was
identified as indispensable for embryonic lymphangiogenesis. Ccbe1 acts at the same stage of
30
development as VEGF-C and is required for the budding of the lymphangioblast and angiogenic
sprouting from venous endothelium (Hogan et al. 2009). Furthermore, a novel two-step
screening strategy involving a simple phenotypic read-out (edema formation or larval lethality)
followed by semi-automated in situ hybridization using a chemical library of 1,280 bioactive
compounds identified the adenosine A1 receptor antagonist, 7-chloro-4-hydroxy-2-phenyl-1,8naphthyridine, as an inhibitor of blood vascular and lymphatic development in Xenopus frogs
(Kalin et al. 2009).
2.1.5
Molecular markers of lymphatic endothelium
For many years, the lack of specific lymphatic markers has hampered research in the lymphatic
vascular field. However, the recent establishment of pure blood and lymphatic endothelial cell
populations isolated from human dermal tissues has helped improving our understanding on the
development and function of the lymphatic system (Kriehuber et al. 2001; Makinen and Alitalo
2002; Podgrabinska et al. 2002; Hirakawa et al. 2003). LECs and BECs maintain their lineagespecific differentiation in culture even after several passages and gene expression profiling
experiments have revealed that almost 98% of the investigated genes were expressed at
comparable levels in both cell types (Petrova et al. 2002; Hirakawa et al. 2003). Interestingly,
this approach identified previously unknown lymphatic and blood vascular lineage-specific
markers (see Table 2.1.2)
31
Table 2.1.2 Lymphatic and blood vascular lineage-specific markers
Markers
Function
LV
Prox1
Transcription factor
++
Podoplanin
Transmembrane
glycoprotein
++
LYVE-1
Hyaluronan receptor
++
VEGFR-3
Growth factor receptor
+
Neuropilin-2
Growth factor receptor
Macrophage mannose
Adhesion molecule
receptor-1
CCL21
CC-chemokyne
CCL20
CC-chemokyne
BV References
(Wigle et al. 1999)
(Wetterwald et al. 1996;
Breiteneder-Geleff et al.
1999)
(Banerji et al. 1999)
/(+)1 (Kaipainen et al. 1995)
/(+)2 (Yuan et al. 2002)
(Irjala et al. 2001; Irjala et al.
+
2003)
+
(Gunn et al. 1998)
(Kriehuber et al. 2001;
(++)3 (++)3
Hirakawa et al. 2003)
+
CD44
Anchoring protein of
adherent junctions
Connect cadherins to
cytoskeleton in cell-cell
junction
Adhesion molecule,
VEGFR3 co-receptor
Hyaluronan receptor
VEGF-C
Growth factor
+
VEGFR-1
Growth factor receptor
+
Neuropilin-1
Growth factor receptor
+
Endoglin
TGF-b receptor
++
(Hirakawa et al. 2003)
CD34
IL-8
Adhesion molecule
CXC-chemokine
++
+
N-cadherin
Adhesion molecule
+
ICAM
Adhesion molecule
+
Integrin α5
Adhesion molecule
+
Collagen IV
ECM protein
Versican
Proteoglycan
(Young et al. 1995)
(Petrova et al. 2002)
(Petrova et al. 2002;
Hirakawa et al. 2003)
(Erhard et al. 1996)
(Petrova et al. 2002;
Hirakawa et al. 2003)
(Hirakawa et al. 2003)
(Petrova et al. 2002;
Hirakawa et al. 2003)
(Barsky et al. 1983; Petrova
et al. 2002)
(Petrova et al. 2002;
Hirakawa et al. 2003)
Desmoplakin
Plakoglobin
Integrin α9
Laminin
Collagen XVIII
Basement membrane
molecule
Basement membrane
molecule
+
(Ebata et al. 2001)
+
(Petrova et al. 2002;
Hirakawa et al. 2003)
+
+
/(+)4
/(+)5
++
+
/(+)5
++
/(+)5
++
(Huang et al. 2000; Petrova
et al. 2002)
(Kriehuber et al. 2001)
(Kriehuber et al. 2001;
Hirakawa et al. 2003)
(Hirakawa et al. 2003)
(Hong et al. 2002; Petrova et
al. 2002)
32
BV, blood vessels; CCL, CC chemokine ligand; LV, lymphatic vessel; LYVE-1, lymphatic
vascular endothelial hyaluronan receptor-1; VEGF, vascular endothelial growth factor;
VEGFR, vascular endothelial growth factor receptor. 1VEGFR-3 expression was also found
on same blood capillaries during tumor neovascularisation and in wound granulation tissue
(Valtola et al., 1999; Paavonen et al., 2000). 2Neuropilin-2 is also expressed in veins (Yuan
et al., 2002). 3After activation, both blood vascular and lymphatic endothelial cells strongly
express CCL20 (Kriehuber et al., 2001). 4CD34 expression has also been found on lymphatic
endothelial cells (Sauter et al., 1998; Kriehuber et al., 2001). 5Peripheral lymphatic vessels
sometimes have incomplete basement membrane, large collecting vessels a complete one.
2.1.6
Molecular mediators of adult lymphangiogenesis
The lymphatic vasculature remains quiescent in a physiological adult state. However, during
tissue regeneration, wound healing, tumor growth and inflammation, new lymphatic vessels
develop from preexisting ones. In addition, circulating endothelial progenitor cells (EPCs) and
macrophages have been reported to transdifferentiate into LECs during chronic renal transplant
rejection and corneal lymphangiogenesis (Maruyama et al. 2005; Religa et al. 2005;
Schledzewski et al. 2006). In contrast, bone marrow-derived progenitor cells fail to incorporate
into the endothelium of newly formed lymphatic vessels in mouse tumor xenografts (He et al.
2004) and growing evidence suggests that these cells might only serve as supporting cells that
promote vessel growth (Grunewald et al. 2006; Purhonen et al. 2008).
VEGFR-3 is a potent mediator of adult lymphangiogenesis and VEGF-C and -D are presently
the only known ligands for this receptor (Figure 2.1.5). VEGF-C promotes proliferation,
migration and survival of LECs in vitro (Makinen et al. 2001). VEGF-D also stimulates
lymphangiogenesis in tissue and tumors (Stacker et al. 2001; Veikkola et al. 2001), and
transgenic mice overexpressing VEGF-C or -D in the skin, show hyperplasia of dermal
lymphatic vessels (Jeltsch et al. 1997; Veikkola et al. 2001). In addition, after proteolytic
cleavage, the fully processed, mature forms of VEGF-C and -D can bind to the major known
regulator of angiogenesis, VEGFR-2 (Joukov et al. 1997; Achen et al. 1998). Recently, it has
33
been suggested that VEGFR-2 and -3 might cooperate in terms of LEC migration and
proliferation but not vessel formation (Goldman et al. 2007). However, a mutant VEGF-C,
unable of stimulating VEGFR-2, was shown to be sufficient to activate lymphangiogenesis
(Joukov et al. 1998; Veikkola et al. 2001). Thus, the distinct contribution of VEGFR-3 and 2
towards lymphangiogenesis remain, at present, unclear.
Neuropilins, non-kinase type I transmembrane proteins, are involved in axonal guidance.
Neuropilin-2 expression is also found in veins and lymphatic capillaries. It binds to VEGF-C
(Figure 2.1.5) and this has suggested the possibility that VEGFR-3 signalling could be enhanced
by NRP2 in a similar way as for NRP1 and VEGFR-2 (Soker et al. 2002). Furthermore, the
angiopoietin/Tie signalling pathway is essential for blood vasculature development and recently,
it has been shown that Ang1 also induces lymphangiogenesis in the mouse cornea (Morisada et
al. 2005) and after adenoviral gene transfer in other mouse tissues (Tammela et al. 2005). Ang1
is thought to act either directly on the lymphatic endothelium since its receptor Tie2 is
expressed in LECs in vitro and in vivo (Kriehuber et al. 2001; Petrova et al. 2002) or indirectly
via the VEGF-C-VEGFR-3 pathway, since Ang1 effect could be blocked with soluble VEGFR3 (Tammela et al. 2005b) (Figure 2.1.5). Ang2 is considered to be an antagonist of Tie2, but the
study of Ang2 null mice suggest that Ang2 might act as an agonist of Tie2 in
lymphangiogenesis while being an antagonist in angiogenesis (Gale et al. 2002).
Fibroblast growth factor (FGF)-2 (Chang et al. 2004) and hepatocyte growth factor (HGF) have
been identified as lymphangiogenic factors (Kajiya et al. 2005) promoting in vitro and in vivo
lymphangiogenesis via their receptors FGFR-3, (Shin et al. 2006) and HGFR (Kajiya et al.
2005) (Figure 2.1.5). In contrast to the FGF2 effect, HGF activity was not blocked by inhibiting
VEGFR-3 signalling (Kajiya et al. 2005), so HGF might act directly on the lymphatic
endothelium, while FGF2 could have an indirect effect (Kubo et al. 2002).
Additional lymphangiogenic factors have been proposed, including platelet derived growth
factor (PDGF)-BB (Cao et al. 2004), insulin growth factor (IGF)1-2 (Bjorndahl et al. 2005),
34
growth hormone (GH) (Banziger-Tobler et al. 2008), adrenomedullin (AM) (Jin et al. 2008) and
sphingosine-1-phospate (S1P) (Yoon et al. 2008) (Figure 2.1.5). Conversely, transforming
growth factor (TGF)-β signalling negatively regulates lymphangiogenesis in inflamed tissues as
well as in certain tumor tissues (Oka et al. 2008).
Figure 2.1.5 Receptors expressed by lymphatic endothelium and lymphangiogenic growth factors (from
(Jurisic and Detmar 2009)). Activation of VEGF receptors-2 and -3 (VEGFR-2, VEGFR-3) and neuropilin-2
(Nrp2) by several vascular endothelial growth factors (VEGF-A, VEGF-C, VEGF-D) promotes lymphangiogenesis.
Additional lymphatic growth factors include angiopoietin-1 (ANG-1), hepatocyte growth factor (HGF), fibroblast
growth factor-2 (FGF-2), insulin-like growth factors (IGF-1, IGF-2), platelet-derived growth factor-BB (PDGFBB) and adrenomedullin (AM)
2.1.7
Involvement of lymphatic vessels in disease
2.1.7.1
Lymphedema
Lymphedema is the accumulation of tissue fluid as a direct consequence of a dysfunctional
lymphatic system, where abnormal or damaged lymphatic vessels are unable to drain fluid from
35
the interstitium. Lymph accumulation is characterized by severe swelling of the affected limb,
fibrosis, degeneration of the connective tissue and a high risk of infections (Rockson 2001).
Lymphedemas have been divided into two main groups: primary and secondary.
Primary lymphedema has no identifiable cause and it can appear at birth (congenital), at puberty
(praecox) or in adulthood (tarda). Congenital lymphedema has been associated with a missense
mutation that encodes for a non active VEGFR-3 protein, in Milroy’s disease (Brice et al. 2005).
Lymphedema-distichiasis, that has been associated with mutations in the Foxc2 gene (Fang et
al. 2000; Finegold et al. 2001) is characterized by a double row of eyelashes at birth,
lymphedema at puberty, abnormal lymphatic patterning, agenesis of lymphatic valves, and
abnormal recruitment of SMCs (Petrova et al. 2004). Moreover, mutations of the SOX18 gene
have been described in hypotrichosis-lymphedema-telangiectasia syndrome (Irrthum et al.
2003). Finally, nuchal edema, lymphatic dysplasia and jugular lymph sac distension are
associated with a variety of human syndromes: Edwards syndrome (trisomy of chromosome 18,
(Bekker et al. 2006), Down’s syndrome (trisomy of chromosome 21), Turner’s syndrome (all or
part of one of the X chromosome is absent), Patau’s syndrome (trisomy of chromosome 13),
lethal multiple pterygium syndrome (reviewed in (Chitayat et al. 1989; Christensen et al. 2008)
and Noonan’s syndrome (mutation in the PTPN11, SOS1, KRAS and RAF1 genes or
duplication of chromosome region encompassing gene PTPN11) (Ferreira et al. 2008;
Malaquias et al. 2008). Nuchal edema has been also reported in Fryns syndrome, for which the
disease-causing mutations have not been yet identified (Slavotinek et al. 2005). In most of the
families with lymphedema, however, none of the known mutations occurs, thus several more
genes involved in lymphedema remain unidentified.
Secondary lymphedema develops when the lymphatic vasculature is damaged by infections or
surgery. Filariasis, a parasitic tropical disease, is the main cause of lymphedema (120 million
people affected) (Wynd et al. 2007). The cause of the infection are mosquito-borne parasites
Wuchereria bancrofti and Brugia malayi, which live and reproduce in the lymphatic system;
36
thus the lymphatic vessels are damaged, no lymph fluid is transported and edema appears in the
legs and genitals (Melrose 2002). Current therapies try to target the worms but no drug
(ivermectin or doxycyclin) so far has been able to cure the disease. While the filariasislymphedema is mostly present in tropical and subtropical countries, edema of the arm or of the
axilla after lymph node dissection is the main cause of lymphedema in the industrialized
countries. Lymphedema occurs in 6-30% of mastectomy patients and this rate increases if they
are exposed to radiotherapy. In the future, a combination of VEGF-C therapy and lymph node
transplantation would represent a novel treatment option for individuals with a history of cancer
(Tammela et al. 2007). Importantly, based on its potent lymphangiogenic effect, VEGF-C has
been tested for gene and protein therapy in animals. Adenoviral mediated-VEGF-C and VEGFC156S (a mutant protein that only activates VEGFR-3, (Saaristo et al. 2002a; Saaristo et al.
2002b)) gene therapy has been shown to promote lymphatic vessel generation in the skin
(Karkkainen et al. 2001). Furthermore, regeneration of lymphatic vessels was observed after
injection of VEGF-C in a surgical model of lymphedema in the rabbit (Szuba et al. 2002).
2.1.7.2
Lymphatic vessels and inflammation
Lymphatic vessels participate in inflammation by transporting leukocytes from the site of
inflammation to secondary lymphoid organs. Upon exposure to an inflammatory stimulus,
dendritic cells (DCs) capture antigens in peripheral tissues and migrate to lymph nodes. After
their maturation, DCs express higher levels of the lymphoid chemokine receptor CCR7
(Sallusto et al. 1998) which promotes their migration into lymph nodes, attracted by the
chemokine CCL21 (or SLC, secondary lymphoid tissue chemokine produced by LECs)
(Sallusto et al. 1998; Saeki et al. 1999; Ohl et al. 2004). Once in the lymph node, antigenic
peptide-loaded DCs mediate the presentation of major histocompatibility complex (MHC)peptide complexes to T cells that become activated (Bajenoff et al. 2003). DCs and T cells leave
37
peripheral tissues, enter the lymphatic vessels and migrate towards lymph nodes also via
interaction of CCR7 (Bromley et al. 2005; Debes et al. 2005) with adhesion molecules
expressed by lymphatic endothelium such as CLEVER-1 (common lymphatic endothelial and
vascular receptor-1) (Salmi et al. 2004) and mannose receptor 1 (Irjala et al. 2001). Interaction
of sphingosine-1-phosphate (S1P) with its receptor S1P1 has been implicated in the exit of
lymphocytes from the lymphnodes (Matloubian et al. 2004; Mandala et al. 2002). VEGF-C has
been shown to be upregulated in response to proinflammatory cytokines, presumably through
NF-κB-mediated promoter activation (Ristimaki et al. 1998). Together with VEGF-A, which is
highly expressed by follicular B cells, VEGF-C is a possible mediator of the increased
lymphangiogenesis and DC migration (Angeli et al. 2006). Inflammation-induced
lymphangiogenesis can also be mediated by macrophages (Kerjaschki 2005). They are recruited
into the inflammation site and can transform from naive monocytes into VEGF-C/D-producing
cells (Schoppmann et al. 2002). Human kidney transplant rejection is frequently accompanied
by lymphangiogenesis, and CCL21 produced by the LECs might actively promote the
inflammatory process (Kerjaschki et al. 2004). Furthermore, VEGF-A mediated lymphatic
hyperplasia is observed in UVB-induced skin inflammation in human psoriatic skin lesions
(Kunstfeld et al. 2004; Kajiya et al. 2006). Mice overexpressing VEGF-A in the skin display a
prolonged inflammatory response after induction of delayed-type-hypersensitivity (DTH),
which is associated with LEC proliferation and lymphatic hyperplasia and can be inhibited by
blockade of VEGFR-1 and -2 (Kunstfeld et al. 2004). In chronic airway inflammation induced
by Mycoplasma pulmonis, massive lymphangiogenesis is induced by VEGF-C/-D-producing
inflammatory cells, while blocking VEGFR-3 signalling resulted in bronchial lymphedema
(Baluk et al. 2005). Finally, the role of VEGF-C/D–VEGFR-3 signalling in inflammatory
lymphangiogenesis is also suggested by overexpression of VEGF-C in the joint synovium of
rheumatoid arthritis patients (Paavonen et al. 2002). Together, these studies underline the close
relationship between inflammation, the immune response and lymphangiogenesis.
38
2.1.7.3
Tumor lymphangiogenesis and metastasis
A major prognostic indicator for the progression of human cancers is the detection of metastasis
in regional lymph nodes. Primary cancer cells invade local tissue, enter tumor-associated blood
vessels as a means to target distant organs and enter tumor-associated lymphatic vessels to
target draining (sentinel) distal lymph nodes and distal organs. The lymphatic pathway is the
most common pathway of initial cancer cell dissemination (reviewed by (Stacker et al. 2002a;
Stacker et al. 2002b). Despite this evidence, the molecular mechanisms of lymphatic metastasis
are not completely understood.
Tumor cells can invade pre-existing lymphatic vessels in the surrounding tissues or intratumoral
lymphatic networks (Karpanen et al. 2001; Mandriota et al. 2001; Skobe et al. 2001; Stacker et
al. 2001; Cao et al. 2004; Achen et al. 2005; Cao 2005b; Cao 2005a). In some cases of human
cancers, the presence of intra-tumoral lymphatic vessels was reported to correlate positively
with lymph node metastasis and poor prognosis (Beasley et al. 2002; Dadras et al. 2005; Kyzas
et al. 2005).
Importantly, tumor cells and macrophages secrete growth factors, such as VEGF-C and -D that
can facilitate the transmigration of malignant cells through the lymphatic endothelium.
(Schoppmann et al. 2002), VEGF-C stimulates the growth of new tumor-associated lymphatic
vessels, and in this process nearby LECs send filopodia toward VEGF-C producing tumor cells,
and then form tumor-directed vessel sprouts, where the vessel lumen opens up and allows
facilitated access of tumor cells (He et al. 2005). Moreover, intraluminal VEGF-C also promotes
the dilation of collecting lymphatic vessels, endothelial proliferation in the vessel wall and
further enhances the delivery of tumor cells to sentinel lymph nodes, probably by increasing the
lymph flow rate (He et al. 2005; Hoshida et al. 2006).
39
Direct experimental evidences that increased levels of VEGF-C or VEGF-D promote active
tumor lymphangiogenesis and lymphatic tumor spread to regional lymph nodes have been
provided by studies in animal tumor models (Karpanen et al. 2001; Mandriota et al. 2001;
Skobe et al. 2001; Stacker et al. 2001). These effects are suppressed by specific inhibition of the
VEGFR-3 signalling pathway by either VEGFR-3 blocking antibody, VEGF-C/D trap
(VEGFR-3 extracellular domain fused with immunoglobulin Fc portion) or VEGF-C targeting
siRNA. This inhibition prevents cancer metastasis to lymph nodes and distant organs (Karpanen
et al. 2001; He et al. 2002; Lin et al. 2005; Roberts et al. 2006), thus underlining the association
of VEGF-C or VEGF-D expression and metastasis. Combination of siRNA therapy targeting
both VEGF-C and VEGF-A inhibits lymph node and lung metastasis in a mouse
immunocompetent mammary cancer model (Shibata et al. 2008). Moreover, because VEGF-C
can also activate VEGFR-2 and because VEGF-A promotes lymphangiogenesis, also blocking
VEGFR-2 results in moderate suppression of tumor lymphangiogenesis, as well as
angiogenesis, and a combination of anti-VEGFR-2 and -3 antibodies more potently decreases
lymph node and lung metastasis (Roberts et al. 2006). Recently, it has been shown that
endothelial nitric oxide synthase (eNOS) mediates VEGF-C-induced lymphangiogenesis in
mice; this finding explains the correlation between eNOS and lymphatic metastasis seen in a
number of human tumors and could open the door for potential therapies targeting NO
signalling (Lahdenranta et al. 2009). Moreover, neuropilin-2 expression has been detected on
tumor activated, but not on quiescent, lymphatic vessels (Aslakson and Miller 1992; Caunt et al.
2008). A blocking antibody targeting neuropilin-2 inhibited tumor lymphangiogenesis and
reduced lymphatic metastasis, leaving quiescent mouse lymphatics unaffected (Caunt et al.
2008).
As said, migration to regional lymph nodes represents the first step of tumor metastasis in many
cancers and is an important prognostic indicator for progression of the disease. Interestingly, in
40
animal tumor models it has been shown that primary tumors can prepare a favourable
environment for their future metastatic site even prior to their arrival, partly by producing
lymphangiogenic factors such as VEGF-A (Hirakawa et al. 2005) and VEGF-C (Hirakawa et al.
2007) that promote lymphatic vessel growth in the sentinel lymph nodes. Tumor-infiltrating
macrophages could also contribute to these processes by secreting lymphangiogenic factors
(Schoppmann et al. 2002). Clinicopathological studies of human cancers have shown a direct
correlation between expression of VEGF-C or VEGF-D by tumor cells and lymphatic invasion,
lymph node and distant metastasis, and poor patient survival (Stacker et al. 2002b; Pepper et al.
2003; Achen et al. 2005; Tobler and Detmar 2006). The lymphatic endothelium may also
actively participate in metastasis formation by secreting chemokines such as CCL21, whose
receptor is expressed on some malignant cells (Zlotnik 2004; Shields et al. 2007). Collectively,
these findings suggest that inhibition of lymphangiogenesis at the site of the primary tumor as
well as in the draining lymph node might be of interest for preventing tumor metastasis.
At last, it is important to shortly mention that a role for the lymphatic vascular system in obesity
has been, recently suggested. In fact, fat is accumulated in the edematous tissues of
lymphedema patients and mice lacking one allele of the Prox1 gene develop adult-onset obesity
due to abnormal lymph leakage (Harvey et al. 2005). In addition, a role for lymphatic capillaries
and VEGF-C signalling has also been proposed in hypertension (Machnik et al. 2009). Further
studies will definitely characterize these two processes in the near future.
41
2.2
Mouse embryonic stem cell-derived embryoid bodies
Murine embryonic stem (ES) cells are pluripotent cells capable of differentiating into all
somatic and germ cell types. They are derived from the inner cell mass (ICM) of mouse
blastocysts. When cultured in the presence of anti-differentiation agents such as leukemia
inhibitory factor (LIF) and embryonic fibroblast feeder-cell layers (PMEFs), these cells
maintain their pluripotency and proliferate (Evans and Kaufman 1981) (Figure 2.2.1)
Figure 2.2.1 Mouse embryonic stem cell isolation and culture
(modified from (Wobus and Boheler 2005)). Mouse embryonic
stem (ES) cells are obtained by isolation of the inner cell mass
(ICM) from mouse blastocysts. Once isolated the ES cells can be
cultured in vitro for several passages.
Removal of LIF and PMEFs causes the ES cells to spontaneously differentiate, thus
recapitulating early embryogenesis. When ES cells are cultured in suspension without antidifferentiation agents, they form three-dimensional aggregates called embryoid bodies (EB).
Within 2-4 days in suspension culture, endoderm forms from the inner cell mass, thereby giving
rise to simple EBs (Leahy et al. 1999). On approximately day 4, differentiation of columnar
epithelium with a basal lamina and formation of a central cavity occurs. At this stage, the EBs
are called cystic embryoid bodies (Evans and Kaufman 1981). Upon continued in vitro culture,
EBs give rise to all three germ layers (ectoderm, mesoderm, endoderm) and terminally
42
differentiate into a wide variety of cell types, such as cardiomyocytes, hematopoietic cells,
neurons, pancreatic islet cells blood vascular and lymphatic endothelial cells (Risau et al. 1988;
Bain et al. 1995; Dang et al. 2002; Liersch et al. 2006). The in vitro culture of ES cells as EBs
affords opportunities to mechanistically study early differentiation events of 3D assemblies of
pluripotent cells. One advantage of in vitro differentiation studies is that ES cells can be studied
for gene mutations or knockouts that prove to be lethal during normal embryonic development
in vivo.
Global DNA microarray analysis indicates that EBs temporally express genes in a manner that
recapitulates the sequence of normal development from primitive ectoderm formation, to
gastrulation (Figure 2.2.2), and eventual early cell specification prior to organogenesis (Dvash
et al. 2004). EBs express most, if not all, factors necessary to induce and regulate
embryogenesis (Czyz and Wobus 2001; O'Shea 2004). Expression of phenotypic markers of
endoderm (such as Foxa2, Sox17, GATA 4/6, -fetoprotein, and albumin), mesoderm (such as
brachyury-T, Msp1/2, Isl-1, -actin, -globin, and Runx2), and ectoderm (such as Sox1, nestin,
Pax6, GFAP, Olig2, neurofilament, and -III tubulin) demonstrate the ability of EBs to generate
cells from all three germ layers (Itskovitz-Eldor et al. 2000). Furthermore, comparison of the
morphology of an egg cylinder stage mouse embryo (ca. embryonic day 6) with an attached day
7 EB suggests that early stage EBs do mimic early mouse embryogenesis in an organized
manner (Figure 2.2.2). However, time-wise it may very well be that embryonic–like in vitro
development is shifted or spread over time and space; in fact, continuous development of new
cells might disturb the differentiation processes.
43
Figure 2.2.2 Schematic representation of cells and
structures of an embryoid body on day 7 and an
early egg stage cylinder mouse embryo (ED 6). Both
present a parietal (yellow) and visceral (blue)
endoderm, primitive ectoderm (green) and mesoderm
(red). The EB does not present a trophoectoderm (light
pink). The EB central cyst represents the proamniotic
cavity while the blastocoel in the mouse embryos is
substituted with culture medium (pink stripes)
2.2.1
EB culture methods
Common EB culture practices include hanging drop and static suspension culture. The hanging
drop method of EB formation produces homogeneous cell aggregates by dispensing a defined
number of ES cells in physically separated droplets of media suspended from the lid of a Petri
dish (Hopfl et al. 2004). In contrast, static suspension culture is performed by simply adding a
suspension of ES cells to a bacteriological grade Petri dish, thereby allowing the cells to
spontaneously aggregate via cell–cell adhesions (Zhang et al. 2001). Static suspension cultures
produce a large number of EBs rather simply, but the size and shape of the resulting EBs are
highly variable due to the tendency of EBs to agglomerate after initial formation in static
suspension, often producing large, irregularly shaped masses of cells. Entrapment of a single
cell suspension or small clusters of ES cells in hydrogels, such as hyaluronic acid (Gerecht et al.
2007) represents a compromise between hanging drop and static suspension approaches to attain
physically separated EBs in a bulk semi-solid suspension medium; however, the efficiency of
EB formation from individual ES cells can be rather low. Alternative techniques for EB
44
formation and culture have also been developed recently, using multiwell and microfabrication
technologies. These induce aggregation more rapidly than hanging drops but still require
individual processing and manipulation of the resulting EBs (Ng et al. 2005).
2.2.2
Vasculogenesis, blood and lymphatic vascular development and
(anti-) lymph-angiogenesis research
Vasculogenesis is the formation of blood vessels by in situ differentiation of angioblast
precursors. Taking place right after gastrulation, it is one of the earliest and essential
differentiation processes during in vivo embryogenesis. EBs are avascular for the first 4 days of
development. This results in a gradient of oxygen and a deficit of nutrients toward the center of
the EB. As a consequence, hypoxia inducible factor-1α and concomitant VEGF-A become
upregulated, initiating vasculogenesis. At day 5 of culture, clusters of Flk1+, CD31+ (pecam),
MECA-32+, VE-cadherin+ and CD34+ endothelial cells are visible (Vittet et al. 1996). The
process of vasculogenesis is followed by angiogenesis, meaning proliferation and sprouting of
endothelial cells and vessel formation. Angiogenesis starts at day 8 of culture and 1-2 days are
needed to visualize the first blood vessel-like structures in the EBs (Vittet et al. 1996).
While the development of the blood vascular system has been long investigated using mouse
embryoid bodies, the differentiation of ES into lymphatic endothelial cells has been reported
only recently (Figure 2.2.3, (Liersch et al. 2006)). In the EBs, the lymphatic vessel-like
structures develop after the formation of the blood vessel-like structure networks, thus
mimicking the in vivo situation (Figure 2.2.4).
45
Figure 2.2.3 Development of lymphatic vessel-like structures in ES-derived EBs (modified from (Liersch et
al. 2006)). Double immunofluorescence stains of 21 days old EBs for CD31 (red; A, D), LYVE-1 (green, B;
arrows) and Prox1 (green, E; arrows) reveal CD31+ blood vessels and CD31+/LYVE-1+/Prox1+ lymphatic vessels
(C, F: merged images; arrows). Blood vessels expressed the blood vascular-specific marker MECA-32 (red, G)
whereas Prox1+ lymphatic vessels (H; green; arrow) were MECA-32- (I: merged image; arrow). Scale bars: 50 µm
Figure 2.2.4 Lymphatic vascular system development in mouse EBs largely mimics the in vivo situation. In
mouse embryos at embryonic day (ED) 5 the first progenitor endothelial cells develop. Later, the first blood vessels
and the heart form. At ED 9, as soon as the blood vessels mature, lymphatic competence and lymphatic
commitment are acquired by the endothelial cells of the anterior cardinal veins. Those lymphatic endothelial cells
will form the lymphatic vessels. In the embryoid bodies, both blood and lymphatic vessel-like structure formation
is delayed in time but the order of the in vivo developmental events is maintained (Liersch et al. 2006).
46
The evidence that CD31+/LYVE-1+/Prox1+ lymphatic endothelial cells can differentiate and
form lymphatic vessel-like structures in the EBs, and the fact that they respond to
lymphangiogenic factors (e.g. VEGF-C, (Kreuger et al. 2006; Liersch et al. 2006)) has opened a
plethora of possibilities to better understand the mechanisms that control lymphatic vascular
development and, in particular, lymphatic endothelial differentiation. To this end, we have
developed an EB-lymphatic vascular differentiation assay that is depicted in Figure 2.2.5
Figure
2.2.5
Embryoid
body
(EB)
lymphatic
differentiation and (anti-) lymphangiogenic assay. Mouse
embryonic stem cells colonies (red) are cultured in presence
of leukemia inhibitory factor (LIF) and a feeder layer of
mouse embryonic fibroblasts (grey). Then LIF and fibroblasts
are removed and single cells (yellow) are plated on a Petri
dish for suspension culture. After 3 days, EBs are formed
(green). EBs of same size are singularly picked and
transferred to 12multi well plates. After 14 days, treatment is
applied to the EBs (pink, yellow and blue wells). After 4
days, EBs are fixed and immunofluorescence staining (IF) is
performed for blood and lymphatic endothelial markers and
results are quantified.
Given the capacity of the EBs to form blood vessel-like structure, this in vitro model has been
used to screen for anti-angiogenic agents. For example, suramin, tamoxifen and
tetrahydrocortisol have been shown to inhibit angiogenesis in the EB system (Wartenberg et al.
1998; Wartenberg et al. 2001). As the lymphatic endothelial cells can differentiate and form
lymphatic vessel-like structures in the EBs, this in vitro model could serve to screen for novel
(anti)-lymphangiogenic agents as well. To this end, we have also developed an (anti)lymphangiogenic screening (Figure 2.2.5)
47
3 AIM AND OUTLINE OF THIS DISSERTATION
The aim of the present work was to identify the molecular mechanisms that regulate the earliest
phases of lymphatic vascular system development during embryogenesis, as well as novel
lymphangiogenic pathways. Revealing the molecular processes that control lymphatic vessel
formation and remodeling might open up new possibilities for the treatment of lymphatic vessel
associated pathologies, such as lymphedema and cancer metastasis.
We have used mouse embryonic stem cell-derived embryoid bodies that largely mimic mouse
embryogenesis, to identify novel molecular players regulating lymphatic competence and
commitment, which represent the earliest events in lymphatic vascular system development.
This approach identified all-trans-retinoic acid (RA) and, more potently, in combination with
cAMP, as inducer of LYVE-1 (competence) and Prox1 (commitment) expression. In situ and in
vivo studies on mouse and Xenopus embryos confirmed that RA upregulates, while a RARα
inhibitor (Ro 41-5253) or a PkA inhibitor (H89) downregulates, the expression of LYVE-1 and
Prox1 in the endothelial cells of the cardinal veins and developing lymphatics (Chapter 4.1).
The second part of this thesis has focused on the identification of novel anti-lymphangiogenic
factors. Mouse embryoid bodies and human lymphatic endothelial cell in vitro assays were used
to investigate anti-lymphangiogenic properties of active compounds. The screening identified
GW2974, a small molecule inhibitor of the EGFR/ErbB2 tyrosine kinase, as an inhibitor of
lymphatic vessel formation. Moreover, human lymphatic endothelial cell-based in vitro and
mouse in vivo studies have suggested the EGFR/ErbB2 signalling pathway as a novel regulator
of lymphangiogenesis (Chapter 4.2).
48
4 Identification of molecular mechanisms controlling
lymphatic vessel formation
4.1
A role for all-trans-retinoic acid in the early steps of
lymphatic vasculature development
4.1.1
Abstract
Despite recent progress in identifying lymphatic lineage-specific markers and growth factors,
the molecular mechanisms that regulate the earliest steps of lymphatic vascular system
development are unknown. To identify potential regulators of lymphatic competence and
commitment, we used an in vitro vascular differentiation assay with mouse embryonic stem
cell-derived embryoid bodies (EBs). We found that several growth factors, including VEGF-C,
growth hormone, insulin-like growth factor-1 and interleukin-7 moderately induced the
expression of the lymphatic competence marker LYVE-1 in the EBs. Importantly, incubation
with retinoic acid (RA) and, more potently, with RA in combination with cAMP, induced
LYVE-1 expression in the vascular structures of the EBs. This RA effect was dependent on
retinoid acid receptor (RAR)- and protein kinase A signalling. RA and cAMP incubation also
promoted the development of CD31+/LYVE-1+/Prox1+ cell clusters that arose independently
from vessel structures. In situ studies revealed that RAR-is expressed by endothelial cells of
the cardinal vein from ED 9.5 to 11.5 in mice, in areas of LYVE-1 expression. Timed exposure
of mouse embryos and Xenopus laevis tadpoles to excess of RA upregulated LYVE-1 and
VEGFR-3 on embryonic veins and increased formation of Prox1-positive lymphatic progenitors
and lymph sacs. These findings indicate that RA signalling mediates the earliest steps of
lymphatic vasculature development.
49
4.1.2
Introduction
The lymphatic vascular system has an important role in the maintenance of tissue fluid
homeostasis, intestinal lipid absorption, and immune surveillance in that it recruits and
transports immune cells from peripheral tissues to the regional lymph nodes (Alitalo et al. 2005;
Cueni and Detmar 2008). There have been many studies into the role of the lymphatic system in
promoting metastasis to lymph nodes and beyond (Hirakawa et al. 2005; Hirakawa et al. 2007;
Rinderknecht and Detmar 2008) and in modulating inflammatory diseases (Kerjaschki 2006;
Jurisic and Detmar 2009). Much progress has been made following the identification of specific
lymphatic markers that distinguish lymphatic endothelial cells from blood vascular endothelial
cells and novel molecular mediators of lymphatic vessel growth and differentiation, as well as
developmental studies in mouse models (Alitalo et al. 2005; Cueni and Detmar 2008).
There is considerable evidence that during mammalian embryonic development, the lymphatic
vascular system predominantly develops from pre-existing embryonic veins. In mice, expression
of the lymphatic vessel endothelial hyaluronan receptor-1 (LYVE-1) at embryonic day (ED) 99.5 by the endothelial cells that line the anterior cardinal veins is considered to be the first
morphological indication that venous endothelial cells have acquired the competence to respond
to an unidentified lymphatic-inducing signal (Oliver 2004). The biological function of LYVE-1
is unknown; LYVE-1–deficient mice have no major lymphatic or other abnormalities (Huang et
al. 2006; Gale et al. 2007). At approximately ED 9.5, a restricted subpopulation of endothelial
cells on one side of the cardinal vein express the transcription factor Prox1, indicating that this
is the stage of lymphatic commitment (Wigle et al. 2002). These Prox1-positive cells then bud
off from the cardinal veins and migrate away to finally form the primitive lymph sacs. Prox1deficient mice completely lack a lymphatic vascular system (Wigle et al. 2002). Vascular
budding and migration appear to occur under the guidance of local gradients of vascular
endothelial growth factor-C (VEGF-C), which activates its receptor, VEGFR-3, on lymphatic
50
precursor cells (Karkkainen et al. 2004). VEGF-C–deficient mice also lack lymphatic
vasculature and undergo prenatal death because of pronounced fluid accumulation in the tissues
(Karkkainen et al. 2004). It was recently, shown that the transcription factor Sox18 is expressed
in a subset of the cardinal vein cells that later become Prox1-positive lymphatic progenitor cells
and that Sox18 directly activates Prox1 transcription (Francois et al. 2008).
Studies in genetic mouse models indicate that following the formation of the lymph sacs, the
separation of the lymphatic and venous system is mediated by the tyrosine kinase Syk and the
adaptor protein SLP-76 (Abtahian et al. 2003; Sebzda et al. 2006), the sprouty-related
ena/VASP homology 1 domain-containing proteins (spred)-1 and 2 (Taniguchi et al. 2007), and
angiopoietin-like protein 4 (Backhed et al. 2007). Further lymphatic vessel maturation and
remodeling are controlled by a plethora of molecules (reviewed by (Cueni and Detmar 2008))
that includes the transcription factor Foxc2 (Petrova et al. 2004), angiopoietin-2 (Gale et al.
2002; Shimoda et al. 2007), the non-kinase receptor neuropilin-2 (Yuan et al. 2002), ephrin B2
(Makinen et al. 2005) and the transmembrane glycoprotein podoplanin (Schacht et al. 2003).
Despite advances in our understanding of lymphatic vasculature development, the molecular
mechanisms that control the earliest stages of lymphatic competence (expression of LYVE-1 by
endothelial cells of the cardinal vein and lymphatic precursor cells) have not been determined.
To identify pathways that mediate lymphatic competence, we used a previously established
embryoid body (EB)-based vascular differentiation assay (Liersch et al. 2006) as a screening
model. This system assesses the ability of mouse EB cells to differentiate into lymphatic vessellike structures that express the panvascular marker CD31, as well as Prox1 and LYVE-1
(Liersch et al. 2006); it was previously used to characterize the ability of VEGF-C to promote in
vitro lymphangiogenesis (Kreuger et al. 2006; Liersch et al. 2006). Using this model system, we
51
investigated the potential effects of soluble factors that have been previously reported to be
potentially associated with activity on lymphatic endothelial cells in vitro or in vivo.
In our study, the growth factors VEGF-C, growth hormone, insulin-like growth factor (IGF)-1
and interleukin (IL)-7 were found to moderately induce the expression of LYVE-1 in EBs.
Incubation of EBs with retinoic acid (RA) and, more potently, a combination of RA and
cyclicAMP (cAMP) induced LYVE-1 expression in the vascular structures; this effect is
dependent on retinoid acid receptor (RAR)- and protein kinase A (PKA) signalling. In situ
studies revealed that RAR-is highly expressed by endothelial cells of the cardinal vein from
ED 9.5–11.5 in mice, in areas of LYVE-1 expression. Most importantly, timed exposure of
mouse embryos and of Xenopus laevis tadpoles to RA resulted in potent upregulation of LYVE1 and VEGFR-3 on embryonic veins and lymph sacs. Together, these findings indicate that RA
signalling could mediate the earliest steps of lymphatic vasculature development.
4.1.3
Materials and methods
4.1.3.1
Mouse embryonic stem cell culture, establishment and treatment of EBs
Murine C57BL/6x129SvEv-derived (passage 3–12) embryonic stem cells (mES cells; kindly
provided by N. Gale, Regeneron Pharmaceuticals, Tarrytown, NY, USA), were cultured on
mitotically inactivated primary mouse embryonic fibroblasts (PMEFs, passage 2-5, Institute of
Laboratory Animal Science,University of Zurich, Switzerland) in Dulbecco’s modified Eagle
medium (Gibco, Eggenstein, Germany), supplemented with 18% fetal bovine serum (Gibco),
100 nM sodium pyruvate (Sigma), minumum essential medium (MEM) vitamins, 2 mM Lglutamine, streptomycin and penicillin (all from Gibco), 10 mM 2-mercaptoethanol and 2000
U/ml recombinant leukaemia inhibitory factor (LIF; Chemicon International, Temecula, CA).
PMEFs and LIF were removed and mES cells were transferred to suspension culture for EB
formation as described (Kurosawa 2007). After 3 or 4 days, EBs of the same size were
52
transferred into 12-well dishes (BD Bioscience, San Diego, CA) and cultured for 14 days
without LIF. Then, EBs were incubated with or without the following factors for 2, 4, 6, 8, 10,
12 or 14 days: 20 ng/ml recombinant human vascular endothelial growth factor-A (VEGF-A
165; kindly provided by the National Cancer Institute, Bethesda, MD); 200 ng/ml recombinant
human VEGF-C (R&D Systems, Minneapolis, MN); 20 ng/ml human fibroblast growth factor-2
(FGF-2, kindly provided by the National Cancer Institute); 100 ng/ml recombinant human
insulin growth factor-1 (IGF-1, R&D Systems); 25 ng/ml recombinant human interleukin-3 (IL3; Chemicon International); 30 ng/ml human hepatocyte growth factor (HGF, R&D Systems);
20 ng/ml human platelet growth factor (PLGF, R&D Systems); 50 ng/ml human growth
hormone (R&D Systems); 20 ng/ml recombinant human interleukin-7 (IL-7, Chemicon
International); 100 µM S-nitroso-N-acetylpenicillamine (SNAP, Sigma-Aldrich, Buchs,
Switzerland); 10 mM human endothelin-3 (ET3, R&D Systems); 1, 2,5, 5 ,10, or 100 µM alltrans-RA (RA; Sigma); 10 µM 4-[E-2-(5,6,7,8-tetrahydro-5,5,8,8-tetramethyl-2-naphthalenyl)1-propenyl]benzoic acid (TTNPB, Sigma); 1 or 10 µM 13-cis-retinoic acid (13-cis-RA, Sigma);
0.5 mM 3',5'-cyclic monophosphate (cAMP; Fluka, Buchs, Switzerland); 10 µM Ro 41-5253
(BioMol International, Plymouth Meeting, PA); 10 µM N-[2-(p-bromocinnamylamino)ethyl]-5isoquinoline sulfonamide, Di-HCl salt (H89, CalbioChem, San Diego, CA). EBs were fixed in –
20°C cold 100% methanol or in 4% paraformaldehyde (PFA) at 4°C for 10 minutes.
4.1.3.2
Endothelial cell culture, real-time RT-PCR, FACS and immunostains
Human umbilical vein endothelial cells (HUVECs), obtained from ScienceCell Research Labs
(San Diego, CA) were seeded into fibronectin-coated culture dishes (10 µg/ml; BD Biosciences,
Bedford, MA) and were cultured in endothelial cell basal medium (EBM; Cambrex Bio Science,
Walkersville, MD) supplemented with 20% fetal bovine serum (FBS; Invitrogen, Grand Island,
NY), 2 mM L-glutamine, antibiotic-antimycotic solution, 10 µg/ml hydrocortisone and N6,2'-O-
53
dibutyryl-adenosine 3',5'-cyclic monophosphate (25 µg/ml; all from Fluka, Buchs, Switzerland).
Cells from passages 6 to10 were used. HUVECs were incubated for 24 hours with 10 µM RA
plus 0.5 mM cAMP or with dimethyl sulfoxide (DMSO) as negative control. Then real-time
RT-PCR, fluorescence-activated cell sorting (FACS) and immunostaining were performed. For
real-time RT-PCR analyses, total cellular RNA was isolated using the Trizol reagent
(Invitrogen) and was extracted with chloroform, precipitated with isopropanol, washed with
70% ethanol, and dissolved in DNase-free/RNase-free distilled water. The concentration of
RNA was measured using a NanoDrop ND-1000 spectrophotometer (Witec AG, Littau,
Switzerland). The expression of LYVE-1 and Prox1 mRNA was quantified by TaqMan realtime RT-PCR using the AB 7900 HT fast real-time PCR system (Applied Biosystems, Foster
City, USA). The following probes and primers were used: LYVE-1 forward primer (FP) 5’AGCTATGGCTGGGTTGGAGA-3’, reverse primer (RP) 5’ CCCCATTTTTCCCACACTTG3’, probe 5’ FAM-TTCGTGGTCATCTCTAGGATTAGCCCAAACC-BKH1-3’; Prox1: FP 5’ACAAAAATGGTGGCACGGA-3’, RP 5’-CCTGATGTACTTCGGAGCCTG-3’, probe 5’FAM-CCCAGTTTCCAAGCCAGCGGTCTCT-BKH1-3’. Each reaction was normalized for
the
expression
of
ß-actin
TAATGTCACGCACGATTTCCC-3’,
(FP
5’-TCACCGAGCGCGGCT-3’,
RP
5’-
probe
5’-JOE-CAGCTTCACCACCACGGCCGAG-
TAMRA-3’). Student t-test was performed in Office EXCEL. For fluorescence-activated cell
sorting (FACS) analysis, cells were stained with rabbit anti-human LYVE-1 antibody
(ReliaTech, Braunschweig, Germany), anti-rabbit fluorescein isothiocyanate (FITC) or isotype
control antibodies (CALTAG/Invitrogen, Basel, Switzerland), and FACS was performed using a
BD FACSCanto (BectonDickinson, Basel, Switzerland) and the FACSDiva software. Data were
analyzed with Flowjo software (Treestar, Ashland, TN). For immunostains, cells were stained
with human LYVE-1 antibody (ReliaTech) and human Prox1 antibody (kindly donated by Prof.
K.Alitalo, Helsinki, Finland), and corresponding secondary antibodies were labeled with Alexa-
54
Fluor 488 or 594 (Molecular Probes). Cell nuclei were counterstained with Hoechst
bisbenzimide (Sigma). All experiments were performed three times with comparable results.
4.1.3.3
Immunofluorescence and quantitative analysis of vessel development in EBs
EBs (n=9 per group) were stained with antibodies against mouse LYVE-1 (Angiobio, Del Mar,
CA and R&D Systems), CD31 (BD Bioscience) or Prox1 (kindly provided by Dr. Kari Alitalo),
and corresponding secondary antibodies were labeled with Alexa-Fluor 488 or 594 (Molecular
Probes, Eugene, OR). Cell nuclei were counterstained with Hoechst bisbenzimide (SigmaAldrich). LYVE-1/CD31 stained sections were examined with a Zeiss Axiovert 200M
microscope, and images were captured with a Zeiss AxioCam-MRm (Carl Zeiss; Oberkochen,
Germany). Image acquisition in the individual fluorescent channels was accomplished using
Axio Vision4.4 software (Zeiss). Adobe Photoshop CS3 (Adobe Systems, San Jose, CA) was
used to adjust image brightness and for image overlay. Computer-assisted morphometric vessel
analyses were performed using the IP-LAB software (Scanalytics; Fairfax, VA). The total EB
area was examined and the vessel area and number per EB were determined. The average vessel
area and vessel number per group was then calculated and statistical analysis (unpaired
Student’s t-test) was performed using MS Excel 2003. Prox1/LYVE-1/CD31 positive cell
cluster imaging and quantification was done with a Delta Vision microscope at 20x, and the 3dimensional images were captured with a Roper CoolSnap HQ camera and analyzed with
DeltaVision
Analysis
Software (Applied
Precision
SoftWoRx,
Issaquah,
WA)
for
deconvolution.
4.1.3.4
Immunohistochemistry of mouse embryo sections
FVB mice (12 to 16 weeks of age; 2 estrous females and 1 male per cage) were allowed to breed
overnight. Females with detectable vaginal plugs on the next morning were determined to be at
55
day 0 of pregnancy. Pregnant mice were housed individually. The pregnant mice were sacrificed
by CO2 at days 9.5, 10.5 and 11.5 of pregnancy. The embryos were removed by laparotomy. All
embryos were immediately fixed in 4% PFA at 4°C for 48 hours, then dehydrated in ethanol
series, cleared in xylene, and embedded in paraffin wax. Serial cross 10 µm-sections of the
embryos were cut and mounted on glass slides. After they were dewaxed in xylene, sections
were hydrated and processed for immunohistochemistry for LYVE-1 (goat biotinylated antimouse; R&D Systems) and RAR-(rabbit anti-mouse; Santa Cruz Biotechnology). In
additional experiments, RA (Sigma-Aldrich) or Ro 41.5253 (BioMol International) were
dissolved in corn oil right before use; pregnant mice (n=5 per treatment group) were given two
intra-peritoneal injections of 25 mg/kg of body weight of RA or 50 mg/kg body weight of Ro
41-5253 in corn oil on days 8 and 10 of pregnancy. Control mice (n=5) received an equal
volume of corn oil. Mice were sacrificed on day 11.5 of pregnancy. Embryos were embedded in
paraffin wax or frozen in OCT (Sakura FineTek, Torrance, CA). Serial cross sections of the
embryos were cut at a thickness of 10 µm (paraffin) or 15 µm (frozen sections) and were
mounted on glass slides. Paraffin sections were processed for LYVE-1 immunohistochemistry
as described above. The sections were exposed to AEC (3-amino-9-ethylcarbazole) chromogen
that forms a red end-product at the site of the antigen and the reaction was stopped after 1
minute incubation. Frozen sections were fixed in 100% methanol at –20°C for 10 minutes and
then processed for immunofluorescence staining for Prox1 (kindly donated from Prof. K
Alitalo) and CD31 (BD Bioscience), using corresponding secondary antibodies labelled with
Alexa-Fluor 488 or 594 (Molecular Probes, Eugene, OR). Cell nuclei were counterstained with
Hoechst bisbenzimide. The stained sections were examined with a Zeiss Axioskop 2 mot plus.
All images were captured with a Zeiss AxioCam-MRm. The quantification of LYVE-1
expression was performed using Photoshop C3; the number of selected LYVE-1 stained pixels
was determined and their percentage per total pixel number per picture (LYVE-1 area fraction)
was then determined. The jugular lymph sac area quantification was performed using the area
56
measurement tool of Photoshop C3. The quantification of CD31+/Prox1+ cells was performed
by counting the number of CD31+/Prox1+ cells per picture. Three sections per embryos and
four embryos from four different pregnant mice were analyzed.
4.1.3.5
Xenopus laevis embryos and in situ hybridization
In vitro fertilization, culture and staging of Xenopus laevis embryos (tadpoles) were performed
as previously described (Brandli and Kirschner 1995; Helbling et al. 1999). Embryos were
cultured in 0.1X MMR (0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES,
and 0.1 mM EDTA) at 22°C until stage 28. Thereafter, 20 µM RA plus 1 mM cAMP or DMSO
were added to the medium. In addition, tadpoles were also bathed with Ro 41-5253
(concentration range from 100nM to 20µM) but the treatment resulted in lethality within the
first 4 hours of exposure. Embryos were fixed at stage 39 in 4% formalin for 1 hour. Probe
synthesis, whole-mount in situ hybridization and bleaching of embryos were carried out as
described (Helbling et al. 1998; Helbling et al. 1999; Saulnier et al. 2002). For in situ
hybridization, probe synthesis, whole-mount in situ hybridization and bleaching of embryos
were carried out as described (Helbling et al. 1998; Helbling et al. 1999; Saulnier et al. 2002).
Digoxigenin probes were generated from linearized plasmids encoding for LYVE-1, Prox1,
VEGFR-3 and CD31. Digital photographs of stained embryos were taken with an AxioCam
Color camera mounted on a Zeiss Stereo Lumar.V12 stereoscopic microscope.
57
4.1.4
Results
4.1.4.1
RA promotes lymphatic differentiation of EBs
We used the EB vascular differentiation assay to identify regulators of the early steps of
lymphatic vascular system development (Liersch et al. 2006). We first investigated the effects
of several candidate molecules on the number and size of CD31+/LYVE-1+ vessel-like
structures. The factors were added to EBs, starting at day 14 after initiation, for up to 10 days. In
accordance with previous results (Kreuger et al. 2006; Liersch et al. 2006), VEGF-C promoted
the formation CD31+/LYVE-1+ vascular structures, compared with the vehicle control. The
maximum induction occurred on day 4 of exposure (data not shown), so this was chosen as the
study period for further investigations.
Incubation of EBs with VEGF-C, growth hormone, IGF-1 and IL-7 significantly promoted the
expression of LYVE-1 in CD31+ structures (Figure 4.1.1 A). Incubation of EBs with RA and,
even more potently, with a combination of RA and cAMP, resulted in an enlarged area of
CD31+/LYVE-1+ structures, compared to controls (Figure 4.1.1 A). Only VEGF-C and RA,
with or without cAMP, also significantly promoted the number of CD31+/LYVE-1+ structures
(Figure 4.1.1 B). In contrast, no major effects on the total area or number of CD31+/LYVE-1+
structures were detected after incubation with placental growth factor, hepatocyte growth factor,
IL-3 or the nitric oxide donor S-nitroso-N-acetyl-l,l-penicillamine (SNAP). cAMP did not
enhance VEGF-C effect and incubation with cAMP alone had no effect on EB vasculature
(Figure 4.1.1 A). In agreement with a specific role of retinoic receptors, we found that in
addition to RA, 13-cis-retinoic acid (+cAMP) and the synthetic retinoid analogue TTBNP (+
cAMP) also promoted significant formation of LYVE-1+ /CD31+ area (Figure 4.1.1 C).
Incubation of EBs with RA significantly increased the EB area covered by CD31+/LYVE-1+
structures (Figure 4.1.2 D-F) and, in agreement with the documented synergistic effect of RA
58
and cAMP in other systems (Lang et al. 1989; Drab et al. 1997; Hatzopoulos et al. 1998), the
combination of RA and cAMP for 4 days resulted in LYVE-1 expression by most of the CD31+
endothelial cells (78.6%+14%; Figure 4.1.2 G–L’), compared to 27%+14% in control EBs
(Figure 4.1.2 A–C). A quantitative analysis revealed that the RA+cAMP treatment significantly
increased the total CD31+ area and the CD31+/LYVE-1+ area, but not the CD31+/LYVE-1area, as compared with controls (Figure 4.1.2 M), thus excluding the possibility that the
observed increase of LYVE-1 expression might be a secondary effect due to an overall
increased amount of CD31-positive vascular structures.
59
Figure 4.1.1 The in vitro mouse embryoid body (EB) assay reveals that retinoic acid (RA) and cAMP induce
LYVE-1 expression. Mouse EBs were cultured for 14 days and were then incubated with compounds for 4 days.
10 µM all-trans-RA (RA) and RA+0.5 mM cAMP (RA+cAMP) increased the LYVE-1+ area among CD31+
vessel-like structures (A). VEGF-C, IGF-1, GH and IL-7 also showed a significant induction of LYVE-1
expression (A). 10 µM all-trans-RA + cAMP and VEGF-C induced the formation of several distinct LYVE1+
lymphatic vessel-like structures (B). 13-cis-RA + cAMP and the synthetic retinoid TTBNP (+ cAMP) increased the
area of LYVE-1 expression (C). Exposure of EBs to Ro 41-5253, an inhibitor of RAR-α, in combination with
RA+cAMP inhibited the expression of LYVE-1 in CD31+ vessel-like structures. H89, an inhibitor of PKA, also
inhibited the effects of RA + cAMP. Ro 41-5253 did not significantly inhibit the VEGF-C effect. EBs exposed to
the inhibitors alone expressed the same levels of LYVE-1 as controls. Data are expressed as mean values + SEM
(n= 5), *p<0.05; **p<0.01;*** p<0.001.
60
Figure 4.1.2 Incubation of EBs with RA and cAMP induces LYVE-1 expression in CD31+ vessel-like
structures. Differential immunofluorescence analysis of control EBs for expression of CD31 (red) and LYVE-1
(green) showed the formation of CD31+/LYVE-1- blood vessel-like structures whereas only a few endothelial cells
expressed LYVE-1 (A-C). Incubation with RA induced LYVE-1 expression in the CD31+ structures (D-F). RA in
combination with cAMP (RA+cAMP) further increased the CD31+/LYVE-1+ area (G-I, J-L’ and M). The total
CD31+ and the CD31+/LYVE-1+ area, but not the CD31+/LYVE-1- area was increased (M). Scale bars=100 µm;
data are expressed as mean values + SEM (n=9).
61
Importantly, incubation of EBs with RA and cAMP for up to 4 days significantly increased the
number of Prox1+/LYVE-1+/CD31+ cell clusters formed (Figure 4.1.3 E-J ; Figure 4.1.4, E-H),
whereas only a few individual Prox1+/LYVE-1+/CD31+ cells, but no cell clusters, were
detected in control EBs (Figure 4.1.3 A-D and J plus Figure 4.1.4 A-D).
Figure 4.1.3 Incubation of EBs with RA and cAMP increases formation of Prox1+/LYVE-1+/CD31+ cell
clusters. Double immunofluorescence analysis revealed that incubation with RA and cAMP induced expression of
Prox1 (green; F, G, H, I) in CD31+ (red; E, G, I) and in LYVE-1+ (Figure 4.1.4 E-H) endothelial cell clusters,
compared with control EBs where only a few Prox1+ (B, C and D) and CD31+ (A, C) cells were visible. Cell
nuclei are stained blue (D, H). Pictures were obtained after 2 days of exposure to RA and cAMP. Scale bars=50
µm. Quantitative analysis revealed that RA and cAMP exposure increased the number of CD31+/Prox1+ cell
clusters in EBs compared with negative controls, beginning at 1 day after treatment initiation (J). Data are
expressed as mean values + SEM (n= 5), *p< 0.05; **p<0 .01; ***p<0 .001.
62
Figure 4.1.4 Prox1 positive endothelial cell clusters in
the
EBs
are
also
LYVE-1
positive.
Double
immunofluorescent staining for LYVE-1 (red) and Prox1
(green) showed that EB treatment with RA+cAMP induced
expression of Prox1 (F, G) in LYVE-1+ (E, G) endothelial
cell clusters as compared with control (A-D) Cell nuclei:
blue (D, H). Scale bars: 20 µm.
We next investigated whether the effects of RA and cAMP on the formation of lymphatic
vessel-like structures was inhibited by the RAR-–specific antagonist Ro 41-5253. Incubation
of EBs with Ro 41-5253 alone did not affect the formation of CD31+/LYVE-1+ or
CD31+/LYVE-1+/Prox1+ structures, whereas Ro 41-5253 potently inhibited the induction of
lymphatic vessel-like structures by RA and cAMP (Figure 4.1.1 C and Table 4.1.1).
63
Day 16
Table 4.1.1
#EBs
Day 18
#Clusters
#EBs
#Clusters
Control
0/9
0
0/9
0
10 µM RA
0/9
0
0/9
0
10 µM RA+0.5 mM cAMP
9/9
3+1
9/9
1+0
10 µM RA+0.5 mM cAMP+10 µM Ro 41-5253
0/9
0
0/9
0
10 µM RA+0.5 mM cAMP+10 µM H89
0/9
0
0/9
0
0.5 mM cAMP
0/9
0
0/9
0
10 µM Ro 41-5253
0/9
0
0/9
0
10 µM H89
0/9
0
0/9
0
Table 4.1.1 Blockade of RAR- or of PKA inhibits the induction of Prox1+ cell clusters by RA and cAMP.
Four days of treatment with RA and cAMP induced the formation of 3+1 Prox1+/LYVE-1+/CD31+ cell clusters
per EB. These clusters were found in 9 out of 9 EBs analyzed. Ro 41-5253 (RAR-inhibitor) and H89 (PKA
inhibitor) completely prevented the induction of Prox1 endothelial cell clusters by RA and cAMP. No induction
was seen after incubation with RA alone, cAMP alone or with the inhibitors alone. #EB: number of analyzed EBs;
#Clusters: average number of CD31+/LYVE-1+/Prox1+ cell clusters per EB + SD.
The cAMP-dependent PKA inhibitor H89 also completely blocked the induction of
CD31+/LYVE-1+/Prox1+ structures by RA and cAMP; incubation of EBs with H89 alone had
no effects (Figure 4.1.1 C and Table 4.1.1). Together, these findings indicate a sufficient role of
the RAR–α and cAMP–PKA pathway in promoting the retinoid effects on lymphatic
differentiation in the mouse EB assay.
4.1.4.2
In vivo effects of RA in developing mouse embryos
We investigated whether RA could affect in vivo development of the mouse lymphatic vascular
system. Immunohistochemical analyses were used to determine whether RA receptors are
expressed by endothelial cells of the cardinal vein of mouse embryos at ED 9.5–11.5, when
64
expression of LYVE-1 and Prox1 is first observed. We found that RAR- was expressed
(Figure 4.1.5, B’, arrowheads) by the endothelial cells of the LYVE-1+ cardinal vein (Figure
4.1.5 A’, arrowheads) and by the developing lymph sacs at ED 11.5. In fact, RAR- was
expressed on/nearby the cardinal veins by EDs 10.5 (Figure 4.1.5 C) and 9.5 (Figure 4.1.5 D),
time-points at which the jugular lymph sacs had not yet formed.
Figure 4.1.5 RAR- is expressed by endothelial cells of the cardinal veins of mouse embryos.
Immunohistochemical analysis of mouse embryos for LYVE-1 (red; A-A’: ED 11.5) and RAR- (B, B’: ED11.5;
C: ED10.5; D: ED9.5) revealed that RAR- (B, B’, arrowheads) is expressed by/near the endothelial cells of the
LYVE-1+ (A, A’, arrowheads) cardinal veins and forming jugular lymph sacs of ED 11.5 as well as in/near the
cardinal veins of ED 10.5 (C, arrowheads) and ED 9.5 (D, arrowheads) mouse embryos. Two serial sections are
shown in panels A and B. Scale bars: A, B: 100 µm; A’, B’, C, D: 20 µm. Cell nuclei were counterstained with
hematoxylin. a, dorsal aorta; jls, jugular lymph sac; nt, neural tube; v, cardinal vein.
65
Based on the observed expression pattern of RAR- during lymphatic development, we
investigated whether RA also induced in vivo expression of LYVE-1 and Prox1. To this end, we
injected RA intraperitoneally into pregnant mice, to expose the developing embryos to an in
utero excess of RA. RA (25 mg/kg of weight) was injected on days 8 and 10 of pregnancy. ED
8 was chosen as the first injection time-point to ensure RA exposure before LYVE-1 and Prox1
were expressed by cardinal vein endothelium (at ED 9). ED10 was chosen for the second
injection to ensure that lymphatic-committed endothelial cells were exposed to excess RA as
they were budding from the cardinal veins.
At ED 11.5, when the first jugular lymph sacs are visible, immunohistochemistry with a short 1
min colorimetric reaction revealed strong LYVE-1 expression on the cardinal vein endothelial
cells of embryos that were exposed to RA in utero (Figure 4.1.6 C, C’ and D) but not of control
embryos (Figure 4.1.6 A, A’ and B). A quantitative analysis confirmed that LYVE-1 expression
was significantly upregulated upon treatment (Figure 4.1.6 G). We also observed an increase in
the size of the primary jugular lymph sacs in embryos exposed to RA (Figure 4.1.6 H; p=0.19).
In contrast, in utero exposure of the embryos to Ro 41-5253, an inhibitor of RAR-α, led to a
decrease in LYVE-1 expression by the ECs of the cardinal vein and of the jugular lymph sac
(Figure 4.1.6 E-F and G), as compared to control embryos (Figure 4.1.6 A, A’, B, G). The
lymph sac area was not affected by Ro 41-5253 (Figure 4.1.6 H). Importantly, differential
immunofluorescence analyses of CD31 and Prox1 expression revealed that embryonic exposure
to RA increased the number of Prox1+ endothelial cells in the cardinal veins and of sprouting
Prox1+ cells that form the lymph sacs (Figure 4.1.7 D-D’’’, E, and F), compared with control
embryos (Figure 4.1.7 A-A’’’, B and C). A quantitative analysis confirmed that the number of
CD31+/Prox1+ cells in the jugular area of the embryos significantly increased upon RA
treatment (Figure 4.1.7 I), whereas injection of Ro 41-5253 slightly decreased the number of
66
CD31+/Prox1+ cells (Figure 4.1.7 G-G’’’, H and I) as compared to controls (Figure 4.1.7 AA’’’, B, C and I).
Figure 4.1.6 In utero exposure of mouse embryos to excess of RA upregulates endothelial LYVE-1 expression
in the anterior cardinal veins and jugular lymph sacs. Immunohistochemical analysis of paraffin sections from
ED 11.5 mouse embryos for LYVE-1 (red) revealed that the endothelial cells of the anterior cardinal veins and of
the forming lymph sacs upregulated the expression of LYVE-1 after in utero exposure to an excess of RA (C, C’, D
and G), as compared with control embryos (A, A’, arrowheads, B and G). Treatment with the RAR antagonist Ro
41-5253 decreased LYVE-1 expression (E, E’, F and G). The jugular lymph sac area was enlarged after RA
treatment as compared to controls (H). A' - B, C' - D and E' - F are sections of embryos from two independent
pregnant mice each. Scale bars: A, C, E, 100 µm; A’-F, 50 µm. Jls, jugular lymph sac; nt, neural tube; v, cardinal
vein; data are expressed as mean values + SD (n=4) **p<0 .01.
67
Figure 4.1.7 In utero exposure of mouse embryos to excess of RA increases the number of Prox1-positive
lymphatic progenitor cells in the cardinal veins and jugular lymph sac forming areas. Doubleimmunofluorescence analysis of ED 11.5 mouse embryos exposed to excess RA (D-D’’’, E and F) or vehicle (AA’’’, B and C) for CD31 (green) and Prox1 (red) revealed that RA increased the number of Prox1+ endothelial
cells in the anterior cardinal veins and in the area of the jugular lymph sacs as compared with control (I). Ro 415253 treatment (G-G’’’ and H) slightly reduced the number of Prox1+/CD31+ cells in the jugular area as compared
with control (I). A-B-C, D-E-F and G-H show representative sections of embryos from independent treated
68
pregnant mice, respectively. Cell nuclei: blue (A’, D’, G’). Scale bars: A, A’,D, D’,G, G’: 100 µm; A’’-F: 50 µm.
a, dorsal aorta; jls, jugular lymph sac; nt, neural tube; v, cardinal vein; data are expressed as mean values + SD
(n=4) ***p<0 .001.
The increase in CD31+/Prox1+ cells in the cardinal veins was not due to enhanced cell
proliferation, as revealed by the results of phosphohistone 3 stains (data not shown). ED 11.5
embryos, at gross examination, showed a slight increase in blood presence in the extraembryonic tissues, a lower amount of blood in the heart region as well as a lower heart beat
frequency. The treatment also caused a mild reduction of the caudal length as previously shown
(Chan et al. 2002) (data not shown).
4.1.4.3
RA upregulates expression of LYVE-1 and VEGFR-3 in the vasculature of
Xenopus laevis embryos
Due to their small size, transparency and easy maintenance, Xenopus laevis embryos are a
useful animal model for studying development (Dawid and Sargent 1988), particularly that of
the lymphatic vasculature (Ny et al. 2005). Xenopus embryos were incubated with RA and
cAMP from stage 28 on, when the Prox1 expression is first detected on the vasculature, until
stage 39, when lymphatic endothelial cells start to sprout from the lymph heart and the cardinal
vein to form the first lymphatic vessels. In situ hybridization analyses revealed that exposure of
the embryos to RA increased expression of VEGFR-3 (Figure 4.1.8 G, H and H’) and LYVE-1
(Figure 4.1.8 E, F and F’) throughout the developing vasculature, including the intersomitic
veins (arrowheads in Figure 4.1.8 F and H, H’), the cardinal veins (arrows in Figure 4.1.8 F and
H, H’) and the vitelline network (asterisks in Figure 4.1.8 H), indicative of enhanced lymphatic
competence. Expression of CD31 was downregulated throughout the entire vasculature (Figure
4.1.8 A, B, B’); this was an important observation because CD31 expression is reduced on
lymphatic vessels, compared to blood vessels, during normal development. Prox1+ cells were
69
found in the anterior and posterior cardinal veins, the tip of the tail, the lymph heart and the first
sprouts from the lymph heart in both the RA-exposed and control embryos (Figure 4.1.8 C, D,
D’). A quantitative analysis of the VEGFR-3 expression revealed a significant increase in the
number of VEGFR-3+ sprouts that developed from the anterior cardinal vein and the lymph
heart following exposure to the combination of RA and cAMP, compared with controls
(p=0.002; Figure 4.1.8 I). Only 16%+5% of the control embryos had > 5 VEGFR-3+ sprouts,
compared to 54%+5% of RA-exposed embryos. Moreover, in the embryos exposed to RA,
VEGFR-3+ sprouts were longer than those of the control group (Figure 4.1.8 G, H, H’).
Figure 4.1.8 Incubation of Xenopus laevis embryos with excess of RA downregulates CD31 and upregulates
the lymphatic markers LYVE-1 and VEGFR-3 in the developing vasculature. Xenopus laevis tadpoles were
bathed in RA in combination with cAMP (B, B’, D, D’, F, F’and H, H’) or in DMSO (A, C, E and G) from
embryonic stages 28 to 39. In situ hybridization (blue) for CD31 (A, B, B’), Prox1 (C, D, D’), LYVE-1 (E, F, F’)
or VEGFR-3 (G, H, H’) revealed that RA and cAMP reduced CD31 expression (B, B’) in the developing
vasculature, including intersomitic veins (isv), the cardinal vein (v), and the vitelline network (vn), compared with
controls (A). No effects on Prox1 expression and/or distribution in the lymph heart (lh), lymphatic sprouts (ls) or in
the lymphatic endothelial cells of venous origin (lv) were observed (C, D, D’). In contrast, LYVE-1 expression was
70
upregulated in the intersomitic veins and in the cardinal vein (F, arrowheads, F’) compared with controls (E). RA
incubation also upregulated VEGFR-3 expression in the cardinal vein and vitelline network (H, arrowheads and
asterisk respectively and H’) compared with control embryos (G, arrow heads). Scale bars: 50 µm; Scale bars of
insert pictures: 5 µm. isv, inter somatic veins; lh, lymph heart; ls, lymphatic sprouts; lv, lymphatic endothelial cells
of venous origin; v, cardinal vein; vn, vitelline network. (I) Quantitative analysis of the number of VEGFR-3+
sprouts from the anterior cardinal vein following exposure to DMSO (vehicle) or the combination of RA and
cAMP. Data are expressed as mean + SEM (n=150 per treatment group).
4.1.4.4
Human umbilical vein endothelial cells upregulate LYVE-1 expression upon
RA+cAMP treatment
We investigated whether RA in combination with cAMP could induce the expression of LYVE1 and Prox1 in human umbilical vein endothelial cells (HUVECs) in vitro.
Exposure of HUVECs to RA and cAMP for 12h, led to an upregulation of LYVE-1 mRNA
expression as shown by real-time RT-PCR (>2-fold; p=0.042; Figure 4.1.9 A). Expression of
Prox1 mRNA was unchanged (data not shown). Furthermore, the treatment resulted in an
upregulation of LYVE-1 at the protein level as shown by FACS analysis (Figure 4.1.9 B).
Double immunofluorescence stains for LYVE-1 and Prox1 of HUVECs treated or not with
RA+cAMP showed an upregulation of LYVE-1 but not Prox1 protein expression after treatment
(Figure 4.1.9 C).
These observations, showing that RA induces expression of LYVE-1 in human endothelial
cells, suggest that RA’s effects observed in the EBs in vitro and on embryos in vivo might be
directly mediated by endothelial cells.
71
Figure 4.1.9 HUVECs upregulate LYVE-1 expression after RA+cAMP treatment. (A) Exposure of HUVECs
to RA and cAMP for 12h, led to an upregulation of LYVE-1 mRNA expression as shown by real-time RT-PCR;
data are expressed as mean fold change + SEM (n=3). (B)Treatment resulted in an upregulation of LYVE-1 at the
protein level as shown by FACS analysis. Red profile indicates negative controls, whereas LYVE-1 antibody
reactivity is plotted as blue profile. X-axis represents the fluorescence intensity on a logarithmic scale whereas the
Y-axis shows number of events normalized to the highest cell count; one representative experiment out of three is
depicted; (C) Double immunofluorescence stains for LYVE-1 (red) and Prox1 (green) of HUVECs treated or not
with RA+cAMP showed an upregulation of LYVE-1 but not Prox1 protein expression after treatment; cell nuclei
blue; scale bar 20 µm.
72
4.1.5
Discussion
This is the first demonstration that all-trans-RA is involved in the earliest steps of lymphatic
vascular development, namely the acquisition of lymphatic competence and commitment by
endothelial cells of the embryonic cardinal vein; RA appears to do so by upregulating the
lymphatic markers LYVE-1 and Prox1.
Our studies with the mouse EB vascular differentiation assay found that VEGF-C, growth
hormone, IGF-1 and IL-7 increased the area of CD31+/ LYVE-1+ vessel-like structures in the
EB assay; these findings are in agreement with the reported lymphangiogenic activity of growth
hormone (Banziger-Tobler et al. 2008) and IGF-1 (Bjorndahl et al. 2005) and the lymphatic
reprogramming activity of IL-7 in cultured endothelial cells (Al-Rawi et al. 2005). Most
interestingly, however, we found that RA, alone and combination with cAMP, potently upregulated LYVE-1 expression in CD31+ vascular structures. The LYVE-1+, lymphatically
competent endothelial cells emerged from the pre-existing CD31+ blood vascular endothelium,
supporting the centrifugal theory of lymphatic vasculature development from pre-existing
embryonic veins originally proposed by Sabin in 1902 (Sabin 1902). However, the combination
of RA and cAMP also induced the formation of rare CD31+/LYVE-1+/Prox1+ cell clusters,
independently from the CD31+ vessel-like networks. These findings indicate that mesenchymal
progenitor cells might differentiate directly into lymphatically competent endothelial cells; this
is in agreement with the centripetal theory of mesenchyme-derived lymphatic progenitors
proposed by Huntington in 1910 (Huntington and McClure 1910). The mouse EB vascular
differentiation assay represents a suitable model for lymphatic vasculature development in vivo,
since lymphatic vessels in mice appear to be predominantly derived from pre-existing
embryonic veins, as suggested by recent lineage-tracing studies (Srinivasan et al. 2007),
although there might be a contribution by mesenchyme-derived progenitor cells (Wilting et al.
2003). EB cells appear to be more responsive to the induction of lymphatic commitment than
73
differentiated endothelial cells, since there was no induction of Prox1 by RA and cAMP in
cultured human umbilical vein endothelial cells (HUVECs). The absence of Prox1 induction in
HUVECs, in a 2-dimensional culture system, might indicate the need for additional inducing
factors derived from the microenvironment that are present in the EBs.
RA binds to nuclear retinoic acid receptors (RARs), which after forming heterodimers with
retinoid X receptors (RXRs), activate histone acetylation and gene transcription. Our finding
that Ro 41-5253 inhibited the induction of lymphatic vascular structures by RA indicated that
the RA effect was mediated via RAR-. Ro 41-5253 antagonizes the transactivation of RARs
by RA, with a high affinity for RAR-; 50- to 100-fold higher concentrations of Ro 41-5253 are
required to reduce the activation of RAR- and 1000-fold higher concentrations are required to
inhibit RAR-Apfel et al. 1992). The potentiation of RA’s effects by cAMP and the inhibition
by H89, a cAMP-dependent PKA inhibitor, indicate that RA’s effects are mediated by a
pathway that includes cAMP and PKA (Gaillard et al. 2006). Indeed, it has been found that RA
and cAMP have synergistic effects in several differentiation processes including neural (Lang et
al. 1989), smooth muscle (Drab et al. 1997) and endothelial progenitor cell (Hatzopoulos et al.
1998) differentiation, although the precise mechanisms of RA and cAMP interaction remain to
be established. This concept is supported by our in silico analyses; the AliBaba
(http://www.gene-regulation.com/pub/programs.html),
PROMO
(http://alggen.lsi.upc.es/cgi-
bin/promo_v3/promo/promoinit.cgi?dirDB=TF_8.3) and TESS (http://www.cbil.upenn.edu/cgibin/tess/tess) software indicates that RXRs and RARs, as well as cAMP response elementbinding proteins (CRE-BP), bind to the promoter regions of the mouse LYVE-1 and Prox1
genes (data not shown).
The role of retinoid signalling in the mediation of lymphatic competence is supported by
immunohistochemical analyses showing that RAR- is strongly expressed by the endothelial
74
cells of the embryonic cardinal veins at ED 9.5, when lymphatic competence and commitment
are first detected and at EDs 10.5 and 11.5, when LEC progenitors bud off from the cardinal
vein (Wigle et al. 2002; Oliver 2004). The finding that at EDs 9.5 and 10.5, RAR- was also
expressed by some cells in clusters near cardinal veins, raises the possibility that surrounding
cells might contribute, by indirect effects, to the induction of lymphatic competence by RA.
However, our observation that incubation of HUVECs with RA increased LYVE-1 mRNA
expression and also increased LYVE-1 protein levels, suggests that RA’s effects are directly
mediated by endothelial cells.
To investigate whether RA also promotes lymphatic competence and commitment in vivo, we
used established models of Xenopus and mouse lymphatic development. Previous studies have
shown that excess or lack of RA during early embryogenesis causes malformations (Lammer et
al. 1986; Radhika et al. 2002), through mispatterning of the embryonic tissue layers and the
anterior–posterior body axis. Several RAR knockout mice have been created (Clagett-Dame and
DeLuca 2002). Together, RAR deficiency and vitamin A deprivation lead to a plethora of
embryonic defects, including those of the eye, respiratory tract, heart, kidney, genital tract,
bones and limbs, as well as axial skeletal and neural malformations (Lohnes et al. 1995).
Importantly, however, vitamin A deficiency is also associated with cardiovascular problems and
enlargement of the anterior cardinal veins in rat embryos (White et al. 1998; White et al. 2000).
Zebrafish, that express mutant forms of the RA-metabolizing enzyme Cyp26A1, also display
patterning defects in multiple organs, including the common cardinal vein, pectoral fin, tail,
hindbrain, and spinal cord. Because of the multitude of developmental defects caused by RA
deficiency or exposure (Kochhar 1967; Moro Balbas et al. 1993; Ozeki and Shirai 1998;
Quemelo et al. 2007; Duester 2008), we investigated the in vivo effects of RA exposure over
only short time periods and during the earliest phases of lymphatic development.
Importantly, in mice increased levels of RA in utero upregulated LYVE-1 expression in the
endothelial cells of the anterior part of the cardinal veins and of the forming lymph sacs (which
75
also slightly increased in size). An increased number of CD31+/Prox1+ endothelial cells were
detected in the cardinal veins and the developing lymph sacs. In contrast, in utero injection of
Ro 41-5253, inhibiting RAR-α signalling (Apfel et al. 1992), reduced LYVE-1 expression and
decreased the number of CD31+/Prox1+ cells in the jugular area of the ED11.5 embryos.
However, the extent of inhibition was rather mild and the quantitative evaluation did not reach a
significance level of 0.05, in part due to the rather high variability among the different embryos.
Other possible explanations for these findings are 1) a potential compensatory role of RAR-β
and/or –γ (Manshouri et al. 1997), 2) non-canonical activity of RA via CREB or ERK (Canon et
al. 2004), and 3) an incomplete blockade of RAR-α activity by the two injections of Ro 415253.
Exposure of stage 28–39 Xenopus embryos to RA down-regulated CD31 expression but
enhanced expression of VEGFR-3 and LYVE-1 in the developing vasculature, mainly in the
intersomitic veins, the cardinal vein and the vitelline vein network. Because incubation of
Xenopus embryos with different concentrations of Ro 41-5253 induced lethality of stage 28
tadpoles (data not shown), the vascular effects of specific blockade of RAR-α could not be
evaluated in the Xenopus model. In addition, morpholino knockdown experiments would not
represent a suitable loss-of-function approach since it has been shown that knockdown of either
RAR- α1 or -α2 (or both) leads to a severe phenotype in Xenopus embryos, mainly regarding the
head formation, already at the early stage 20 (Koide et al. 2001) (the first Prox1 expression in
Xenopus is at stage 28).
All-trans-retinoic acid and its derivatives exert many different and often contradictory effects on
cells, depending on context, cell type, and other variables; its role in angiogenesis, for example,
remains so far controversial. It is has been found that RA induces in vitro tube formation of
HUVECs (Saito et al. 2007) via a possible paracrine effect by inducing endogenous VEGF-A
and FGF2 production, and angiogenesis in bovine aortic endothelial cells in vitro (Gaetano et al.
76
2001). On the other hand, RA caused endothelial cells to become refractory to stimulation by
either tumor-conditioned media or various angiogenic factors without interfering with cell
proliferation (Lingen et al. 1996). In addition, there is evidence that the anti-tumor activity of
retinoids involves inhibition of tumor-induced angiogenesis (Oikawa et al. 1989; Majewski et
al. 1995; Lingen et al. 1996).
Interactions between retinoid signalling and Prox1 expression have been described during other
aspects of development. RA modulates Prox1 expression in the dorsal endoderm of mouse
embryos (Burke and Oliver 2002) and promotes neuronal progenitor cell differentiation into
retinal (Zhao et al. 2002) or cochlear cells (Bermingham-McDonogh et al. 2006), which express
Prox1 (Tomarev et al. 1996; Wigle et al. 2002; Dyer 2003; Bermingham-McDonogh et al.
2006). Development of the murine liver also appears to be RA-dependent (Ogura et al. 2004)
and Prox1 is a marker for rat and mouse hepatocytes (Burke and Oliver 2002; Dudas et al.
2006). Moreover, RA, Prox1 and RALDH2 all have important roles in pancreas development
(Burke and Oliver 2002; Molotkov et al. 2005).
Now, retinoid signalling is shown to mediate the earliest steps of lymphatic vasculature
development-the acquisition of lymphatic competence and commitment via expression of
LYVE-1 and Prox1, respectively. It will be important to determine whether retinoids also
modulate post-natal lymphangiogenesis, such as the growth of new lymphatic vessels during
tissue regeneration and cancer progression (Cueni and Detmar 2008).
77
4.2
Activation of the epidermal growth factor receptor
promotes lymphangiogenesis in vitro and in vivo
4.2.1
Abstract
The lymphatic vascular system regulates tissue fluid homeostasis and has an important role in
immune surveillance, inflammation and cancer metastasis. However, the molecular
mechanisms involved in lymphangiogenesis remain poorly characterized. We used our
previously established mouse embryonic stem cell-derived embryoid body vascular
differentiation assay to investigate the effects of candidate small molecules on blood and
lymphatic vessel formation. We found that inhibition of the epidermal growth factor (EGF)
receptor (EGFR) inhibited the formation of lymphatic vessel-like structures. In vitro studies
with human dermal lymphatic endothelial cells (LECs), that were found to express EGFR,
revealed that EGF promotes lymphatic vessel formation; this effect was inhibited by
incubation with an EGFR blocking antibody and with small molecules that target the EGFR or
its associated tyrosine kinase. Incorporation of EGF into a matrigel plug assay revealed that
EGF also promotes lymphangiogenesis in vivo. Moreover, transgenic mice with skin-specific
overexpression of amphiregulin, which also activates EGFR/ErbB2, showed an enhanced
density of lymphatic vessels in the skin. Together, our findings reveal, for the first time, a role
for EGFR activation in lymphangiogenesis and suggest that specific EGFR antagonists might
be used to inhibit pathological lymphangiogenesis.
78
4.2.2
Introduction
The lymphatic vascular system plays an essential role in the maintenance of normal fluid
homeostasis as well as in several pathological conditions, including inflammation and tumor
metastasis. Identification of lymphatic endothelium-specific markers, such as podoplanin and
the lymphatic vessel endothelial hyaluronan receptor-1 (LYVE-1), and studies of lymphatic
system development in a range of genetic mouse models have significantly increased our
understanding of how lymphatic endothelial cell (LEC) differentiation, growth and function
are regulated (Cueni and Detmar 2006). Vascular endothelial growth factor-C (VEGF-C) is,
thus far, the best characterized lymphangiogenic factor that activates VEGF receptor
(VEGFR)-3. Under normal conditions, VEGFR-3 is specifically expressed by LECs but not
by blood vascular endothelial cells (BECs). Activation of VEGFR-3 promotes LEC
proliferation and migration in vitro (Goldman et al. 2007), and lymphatic vessel formation in
vivo (Veikkola et al. 2001). In addition, VEGF-A has also been shown to stimulate
lymphangiogenesis (Hong et al. 2004; Bjorndahl et al. 2005; Hirakawa et al. 2005).
Furthermore, additional growth factors, including fibroblast growth factor-2, hepatocyte
growth factor, angiopoietin-1 and -2, and platelet-derived growth factor, have been recently
shown to promote lymphangiogenic processes (Cueni and Detmar 2008). Because of the
emerging role of the lymphatic vascular system in human disease, including cancer metastasis,
chronic inflammation, organ transplant rejection and hypertension, the identification of
molecular mediators and/or inhibitors of lymphangiogenesis is of immense interest.
Based on the results of a recent chemical library screening for compounds that interfere with
vascular development in Xenopus laevis tadpoles (Kalin et al. 2009), we selected several
compounds to investigate for their possible anti-lymphangiogenic activity in our previously
established mouse embryonic stem cell-derived embryoid body (EBs) vascular differentiation
79
assay (Liersch et al. 2006). In particular, we investigated the effects of calcium channel
inhibitors and of inhibitors of the EGFR/ErbB2 tyrosine kinase, as well as an inhibitor of
VEGF receptor tyrosine kinases. GW2974, a small molecule that targets the EGFR/ErbB2
tyrosine kinase, strongly inhibited the formation of CD31+/LYVE-1+ lymphatic vessel-like
structures in EBs. Conversely, its ligand EGF promoted the vessel formation in the EBs and
also the in vitro migration and tube formation of human lymphatic endothelial cells (LECs),
that we found to express the EGFR in vitro and in vivo. This effect was inhibited by
incubation with an EGFR blocking antibody and with small molecules that target the EGFR or
its associated tyrosine kinase. Incorporation of EGF into a matrigel plug assay revealed that
EGF also promotes lymphangiogenesis in vivo. Moreover, transgenic mice with skin-specific
overexpression of amphiregulin, which also activates EGFR/ErbB2, showed an enhanced
density of lymphatic vessels in the skin. Together, our findings reveal, for the first time, a role
for EGFR activation in lymphangiogenesis and suggest that specific EGFR antagonists might
be used to inhibit pathological lymphangiogenesis.
4.2.3
Materials and methods
4.2.3.1 Mouse embryonic stem cell culture, establishment and treatment of EBs
Murine C57BL/6x129SvEv derived embryonic stem cells (mES cells; passage 3-12; kindly
provided by N. Gale, Regeneron Pharmaceuticals, Tarrytown, NY), were cultured on
mitotically inactivated primary mouse embryonic fibroblasts (PMEFs, passage 2-5, Institute of
Laboratory Animal Science, University of Zurich, Switzerland) in Dulbecco’s modified Eagle
medium (Gibco, Eggenstein, Germany), supplemented with 18% fetal bovine serum (Gibco),
100 nM sodium pyruvate (Sigma-Aldrich, Buchs, Switzerland), MEM vitamins, 2 mM Lglutamine, streptomycin and penicillin (all from Gibco), 10 mM 2-mercaptoethanol and 2000
U/ml recombinant leukemia inhibitory factor (LIF; Chemicon International, Temecula, CA).
80
PMEFs and LIF were removed and mES cells were transferred to suspension culture for
embryoid body (EB) formation as described (Kurosawa 2007). After 3 or 4 days, EBs of the
same size were transferred into 12-well dishes (BD Bioscience, San Diego, CA) and cultured
for 14 days without LIF. Then, EBs were incubated or not with the following factors for 4
days: 100 ng/ml human recombinant epidermal growth factor (EGF, BD Biosciences); 10 µM
all-trans-retinoic acid (RA; Sigma-Aldrich); 0.5 mM 3',5'-cyclic monophosphate (cAMP;
Fluka, Buchs, Switzerland); 200 ng/ml recombinant human VEGF-C (R&D Systems,
Minneapolis, MN) in combination or not with the following substances (all from SigmaAldrich) that were added at 10 µM concentrations: plendil, 4-(2,3-dichlorophenyl)-1,4dihydro-2,6-dimethyl-3,5-pyridinecarboxylic acid ethyl methyl ester (felodipine); YC-93, 1,4dihydro-2,6-dimethyl-4-(3-nitrophenyl)methyl-2-[methyl(phenylmethyl)amino]-3,5pyridinedicarboxylic acid ethyl ester hydrochloride (nicardipine); 4′,5,7-trihydroxyisoflavone,
5,7-dihydroxy-3-(4-hydroxyphenyl)-4H-1-benzopyran-4-one (genistein); N4-(1-benzyl-1Hindazol-5-yl)-N6,N6-dimethyl-pyrido[3,4-d]pyrimidine-4,6-diamine (GW2974). Medium only
and dimethyl sulfoxide (DMSO) were used as negative controls. EBs were fixed in 20°C cold
methanol for 10 minutes.
4.2.3.2 Immunofluorescence analysis of vessel development in EBs
EBs (n=9 per group) were stained with antibodies against mouse LYVE-1 (Angiobio, Del
Mar, CA and R&D Systems), CD31 (BD Bioscience), and corresponding secondary
antibodies labelled with Alexa-Fluor 488 or 594 (Molecular Probes, Eugene, OR). Cell nuclei
were counterstained with Hoechst bisbenzimide (Sigma-Aldrich). CD31/LYVE-1 stained
sections were examined with a Zeiss Axiovert 200M microscope, and images were captured
with a Zeiss AxioCam-MRm (Carl Zeiss; Oberkochen, Germany). Image acquisition in the
individual fluorescent channels was accomplished using Axio Vision4.4 software (Zeiss).
81
Adobe Photoshop CS3 (Adobe Systems, San Jose, CA) was used for image overlay.
Computer-assisted morphometric vessel analyses were performed using the IP-LAB software
(Scanalytics; Fairfax, VA). The total EB area was examined, and the vessel area and number
per EB were determined. The average vessel area and vessel number per treatment group was
then calculated and statistical analysis (unpaired Student’s t-test) was performed using MS
Excel 2003.
4.2.3.3 Proliferation, migration and tube formation of human lymphatic endothelial cells
Human dermal microvascular lymphatic endothelial cells (LECs) were isolated from neonatal
human foreskins by immunomagnetic purification as described (Kajiya et al. 2005). Cells were
cultured in endothelial basal medium (Cambrex, Verviers, Belgium) supplemented with 20%
fetal bovine serum (Gibco, Paisley, UK), antibiotic antimycotic solution (Fluka, Buchs,
Switzerland), L-glutamine (2 mmol/L, Fluka), hydrocortisone (10 µg/ml, Fluka), and N6,2'-Odibutyryladenosine-3',5'-cyclic monophosphate sodium salt (25 µg/ml, Fluka) for up to 11
passages. Cells were grown in a humidified atmosphere at 37°C and 5% CO2.
Proliferation, migration, and tube formation assays were performed as described (Kajiya et al.
2005). Cells were incubated with 100 ng/ml human recombinant EGF (BD Biosciences), 20
ng/ml recombinant human VEGF-A165 (kindly provided by the National Cancer Institute,
Bethesda, MD), 1.2 µg/ml Pichia Pastoris-derived VEGF-C (kindly provided by Dr. K.
Ballmer-Hofer, Paul Scherrer Institute, Switzerland); 10 µM GW2974 (Sigma-Aldrich) or
equal amounts of DMSO as negative control. In additional experiments, cells were incubated
with 10 µg/ml of recombinant humanized monoclonal antibody 2C4 (Pertuzumab, kindly
provided by Roche Diagnostics GmbH, Mannheim, Germany) or 10 µg/ml of an EGFR
blocking
antibody
(Calbiochem,
Darmstadt, Germany).
Mouse
IgG
(Santa
Cruz
Biotechnology, Heidelberg, Germany) was used as control.
82
For proliferation assays, LECs (1.25 to 1.5 x 103) were seeded into fibronectin (Chemicon
International, Temecula, CA) coated 96-well plates. Cells were treated with the different
compounds. After 72 hours, cells were incubated with 5-methylumbelliferylheptanoate
(MUH, Sigma-Aldrich) as described (Detmar et al. 1992). The intensity of fluorescence,
proportional to the number of viable cells, was measured using a SpectraMax Gemini EM
microplate reader (Bucher Biotec AG, Basel, Switzerland). Ten wells per condition were
analyzed. For migration assays, two parallel lines and one orthogonal line were scratched in
confluent LEC cultures, to remove LECs without damaging the fibronectin coating. Pictures
of the two crosses per well were taken immediately after scratching (T0) and after 16 hours of
compound incubation (T16) with a Zeiss AxioCam-MRm (Zeiss). The quantitative analysis of
the non-closed area was performed using Photoshop C3. Three wells per condition were
analyzed.
For tube formation assays, confluent LEC monolayers were overlaid with collagen type I gels
(1 mg/ml; Cohesion, Palo Alto, CA) as described (Kajiya et al. 2005), containing the studied
compounds or their solvent only. Tube-like structure formation was evaluated for up to 20
hours. Three pictures of hot-spots were taken per well, and tube length was analyzed using the
IP Lab software as described (Kajiya et al. 2005). Three wells per condition were used. For all
in vitro studies, three independent experiments were performed. Statistical analyses were
performed using the two-tailed unpaired Student’s t-test (Graph Pad Prism 4).
4.2.3.4 Immunostaining of cultured human LECs
Dermal lymphatic endothelial cells were fixed in 4% paraformaldehyde (PFA) at 4°C, washed
in PBS, stained with rabbit anti-human EGFR antibody (Abcam, Cambridge, UK) and and a
secondary antibody conjugated with Alexa 594. Cell nuclei were counterstained with
83
bisbenzimide. Specimens were examined with a Zeiss Axioskop 2 mot plus. All images were
captured with a Zeiss AxioCam-MRm.
4.2.3.5 Immunoblotting
For Western blot analyses of EGFR protein expression, confluent cultures of LEC were
homogenized in lysis buffer (Kajiya et al. 2005). The protein concentrations were determined
using the NanoOrange Protein Quantification Kit (Molecular Probes, Eugene, OR). The
lysates (100 µg of total protein) were then subjected to SDS-polyacrylamide gel
electrophoresis using NuPAGE 10% BT gels, 1.0 mm, 12 well, and NuPAGE MES SDS
running buffer (20×; Invitrogen). The proteins were transferred onto Trans-Blot Transfer
Medium pure nitrocellulose membranes (BioRad, Hercules, CA) for immunoblot analysis.
Blocking was performed with 5% non-fat dry milk in 0.1% Tween20 (Sigma) in PBS,
followed by immunoblotting with a rabbit anti-human EGFR antibody (Abcam). Specific
binding was detected by the ECL Plus Western Blotting Detection System (GE Healthcare,
Buckinghamshire, UK). Equal loading was confirmed using an antibody against β-actin
(Sigma).
4.2.3.6 In vivo matrigel lymphangiogenesis assay
FVB wild-type mice (female, 9-weeks old, n=5 per group; Charles River, Sulzbach, Germany)
were subcutaneuosly injected with 500 µl of growth factor reduced Matrigel (Becton
Dickinson), containing 1 µg/ml of VEGF-C or 1 µg/ml of EGF (BD Biosciences), or an equal
volume of PBS. After 15 days, tissues were excised, embedded and frozen in optimal cutting
temperature
(OCT)
compound
(Sakura
Finetek,
Zoeterwoude,
Netherlands).
Immunofluorescent staining for mouse LYVE-1 (Angiobio, Del Mar, CA) and CD31 (BD
Pharmingen) was performed using corresponding secondary antibodies labeled with Alexa84
Fluor 488 or 594 (Molecular Probes). Cell nuclei were counterstained with Hoechst
bisbenzimide. The stained sections were examined with a Zeiss Axioskop 2 mot plus. All
images were captured with a Zeiss AxioCam-MRm. Vessel area, density and size were
evaluated with Photoshop C3. For statistical analyses, the two-tailed unpaired student’s t-test
was used (Graph Pad Prism 4). The animal study was approved by the Veterinaeramt des
Kantons Zuerich, Zuerich, Switzerland, permission number 123/2005.
4.2.3.7 Immunohistochemistry
Tail and ear skin paraffin sections from 7-days old keratin14-amphiregulin transgenic mice
(K14-ARtg) (Cook et al. 1997) and from wildtype FVB mice were incubated at 60°C for four
hours, dewaxed in xylene, hydrated and processed for immunohistochemistry for LYVE-1
(goat biotinylated anti-mouse LYVE-1 antibody; R&D Systems). The stained sections were
examined with a Zeiss Axioskop 2 mot plus. All images were captured with a Zeiss AxioCamMRm. Vessel area, density and size were evaluated with Photoshop C3. For statistical
analyses, the two-tailed unpaired Student’s t-test was used (Graph Pad Prism 4). Human skin
sections were fixed in acetone for 10 minutes. Double immunofluorescent staining for
podoplanin (D2-40, Signet, Dedham, MA) and EGFR (Abcam) was performed using
corresponding secondary antibodies labeled with Alexa-Fluor 488 or 594 (Molecular Probes).
Cell nuclei were counterstained with Hoechst bisbenzimide or hematoxylin.
85
4.2.4
Results
4.2.4.1
Blockade of the EGF-EGFR/ErbB2 pathway inhibits the formation of
CD31+/LYVE-1+ vessel-like structures in embryoid bodies
We used our previously established mouse embryoid body (EB) vascular differentiation assay
to investigate the effects of several candidate small molecules on the development of
CD31+/LYVE-1+, lymphatic vessel-like structures. Treatment was started at day 14 after
initiation of the EBs, and was performed for four days. Compounds were added alone or in
combination with all-trans-retinoic acid (RA), cAMP and VEGF-C, which we have recently
found to potently promote lymphatic differentiation in EBs (Marino et al. manuscript
submitted).
EBs incubated with the mixture of RA, cAMP and VEGF-C showed an increased area and an
increased number of CD31+/LYVE-1+ vessel-like structures as compared to control EBs
(Figure 4.2.1 A-B). Incubation with SU5416, a VEGFR-2 tyrosine kinase inhibitor, with the
calcium channel inhibitors felodipine and nicardipine, or with the EGFR/ErbB2 tyrosine
kinase inhibitors genistein and GW2974, significantly reduced the CD31+/LYVE-1+ area of
the EBs. These effects were seen after incubation with the inhibitors alone and after
incubation with the inhibitors together with the mixture of RA, cAMP and VEGF-C (Figure
4.2.1 A). All of the five compounds tested also reduced the number of CD31+/LYVE-1+
vessel-like structures (Figure 4.2.1 B) when added to the EBs alone. In combination with RA,
cAMP and VEGF-C, only genistein, GW2974 and SU5416 showed significant inhibitory
activity (Figure 4.2.1 B). All compounds, alone or in combination with RA/cAMP/VEGF-C,
also slightly reduced the CD31+/LYVE-1- area and structure number (data not shown).
86
Figure 4.2.1 The EGFR/ErbB2 pathway is involved in the formation of CD31+/LYVE-1+ vessel like
structures in EBs. Small molecules (10 µM) were added to the EBs, starting at day 14 after initiation, for four
days in combination or not with a mixture of all-trans-retinoic acid (RA), cAMP and VEGF-C. EBs exposed to
RA+cAMP+VEGF-C showed an increase of the CD31+/LYVE-1+ area and the number of vessel-like structures
as compared to control EBs (A and B). SU5416, felodipine, nicardipine, genistein and GW2974, alone or in
combination with RA+cAMP+VEGF-C, significantly reduced the LYVE-1+/CD31+ area (A). All 5 small
molecules also inhibited vessel-like structure formation as compared to medium only (B). In contrast, only
genistein, GW2974 and SU5416 significantly inhibited vessel like structure formation when used in combination
with RA+cAMP+VEGF-C (B). Data are expressed as mean values + SEM (n=9), *p<0.05; **p<0.01;***
p<0.001.
Whereas incubation of EBs with the mixture of RA, cAMP and VEGF-C increased the
formation of CD31+/LYVE-1+ vessel-like structures (Figure 4.2.2 B-B’’), compared with
control EBs (Figure 4.2.2 A-A’’), addition of GW2974 to the RA/cAMP/VEGF-C mixture
almost completely prevented the formation of CD31+/LYVE-1+ vessel-like structures, and
only clusters of CD31+/LYVE-1+ single cell were detected (Figure 4.2.2 C-D’’).
87
Because inhibition of the EGFR/ErbB2 tyrosine kinas reduced the formation of
CD31+/LYVE-1+ vessel-like structures in EBs we next investigated whether epidermal
growth factor (EGF), an activating legend of the EGFR/ErbB2, might promote vessel
formation. EBs exposed to EGF for 4 days showed a significant increase in the EB area
occupied by CD31+/LYVE-1+ structures (Figure 4.2.3 A) and in the number of these
structures (Figure 4.2.3 B), as compared with controls. VEGF-C, used as a positive control,
showed a similar effect. The EGF effect was completely prevented by addition of GW2974
(Figure 4.2.3 A, B). Microscopical imaging revealed that both VEGF-C and EGF promoted
the formation of CD31+/LYVE-1+ vessel-like structures (Figure 4.2.3 D-E'') as compared to
controls (Figure 4.2.3 C-C’’). Addition of GW2974 together with EGF prevented the
formation of CD31+/LYVE-1+ vessel-like structures and only single CD31+/LYVE-1+ cells
clusters were found (Figure 4.2.3 F-F’’). Taken together, these data indicate a possible role of
EGF via EGFR/ErbB2 signalling in the formation of lymphatic vessel-like structures.
88
Figure 4.2.2 GW2974 inhibits the formation of CD31+/LYVE-1+ vessel-like structures in EBs. Double
immunofluorescent staining for CD31 (red) and LYVE-1 (green) showed the formation of CD31+/LYVE-1+
vessel-like structures in control EBs (medium, A-A’’). Four days of EB incubation with RA+cAMP+VEGF-C
increased the formation and the complexity of structure networks (B-B’’) as compared with control. Cotreatment of EBs with RA+cAMP+VEGF-C and GW2974 strongly inhibited the formation of those structures;
only single CD31+/LYVE-1+ cell clusters are present in the EBs (C-D’’). Scale Bars: 100 μM.
89
Figure 4.2.3 EGF promotes the formation of CD31+/LYVE-1+ vessel-like structures in EBs. VEGF-C,
EGF, GW2974 or DMSO were added or not to EBs, starting at day 14 after initiation, for four days. Exposure to
EGF, as well as to VEGF-C, strongly increased the CD31+/LYVE-1+ area (A) and the number of vessel-like
structures (B) as compared with control. The EGF effect was inhibited by GW2974 but not by DMSO (A, B).
Double immunofluorescent staining for CD31 (red) and LYVE-1 (green) showed that EGF (E-E’’), together with
VEGF-C (D-D’’) promoted CD31+/LYVE-1+ vessel-like structure formation as compared to controls (C-C’’).
Exposure to a combination of EGF and GW2974 inhibited CD31+/LYVE-1+ vessel formation; only
CD31+/LYVE-1+ single cells clusters are visible (F-F’’). Data are expressed as mean values (n=9) + SEM;
*p<0.05; **p<0.01;*** p<0.001Scale bar: 50 µm.
90
4.2.4.2 EGFR is expressed by human LECs in vitro and in vivo
The expression of EGFR on human blood vascular endothelial cells has been previously
reported (Sini et al. 2005). In contrast, EGFR expression on human lymphatic endothelial cells
has not been studied. Immunofluorescence and immunoblot analyses of human LECs in vitro
revealed that LECs express EGFR, although at rather low levels (Figure 4.2.4 A-C). The
expression of EGFR was slightly induced following 24h of EGF exposure (Figure 4.2.4 A-C).
Double immunofluorescent staining of human skin for EGFR and podoplanin revealed that
podoplanin-positive dermal lymphatic vessels express low levels of EGFR in vivo (Figure
4.2.4 D-F; arrowhead). Low levels of EGFR expression were also detected in podoplaninnegative blood vessels (Figure 4.2.4 D-F; arrow).
Figure 4.2.4 Human LECs express EGFR in vitro and in vivo. Immunofluorescent analysis (A-B) and
immunoblot (C) for EGFR showed that human LECs express low levels of EGFR in vitro. The expression of the
receptor was slightly induced following 24h of EGF exposure (A-C). Double immunofluorescent analysis of
human skin for EGFR (red) and podoplanin (green) revealed that dermal lymphatic vessels express low levels of
EGFR (D-F arrow heads). Podoplanin negative blood vessels also express low levels of EGFR (D-F arrows).
Scale bars 50 μm.
91
4.2.4.3
EGF-EGFR/ErbB2 signalling promotes in vitro lymphangiogenesis
Lymphangiogenesis is a complex process that involves proliferation, migration and vessel
formation of LECs. To further characterize the potential role of EGFR/ErbB2 signalling in
these processes, we next treated human LECs with GW2974 and performed in vitro
proliferation, migration and tube formation assays. Migration was evaluated in an in vitro
"scratch" wound healing assay. We found that GW2974 did not inhibit LEC proliferation nor
migration (Figure 4.2.5 A, B), but that it potently inhibited tube formation in vitro (Figure
4.2.5 C), as compared to untreated controls. VEGF-A or VEGF-C, used as positive controls,
showed a significant activation of these processes (Figure 4.2.5 A-C).
Figure 4.2.5 GW2974 inhibits human lymphatic endothelial cell tube formation in vitro. Human dermal
LECs (hLECs) were used to study cell proliferation (A), migration (B) and tube formation (C). Incubation with
VEGF-A significantly promoted hLEC proliferation and migration (A, B). Incubation with GW2974 did not
inhibit hLECs proliferation (A) or migration (B). Incubation with VEGF-C significantly promoted hLEC tube
formation (C). In contrast, GW2974 significantly inhibited tube formation as compared with control (C). Data are
expressed as mean values (n=3) + SEM; t-test: *p<0.05; *** p<0.001. AFU: absolute fluorescent units.
92
We next studied the in vitro effects of EGF on LECs. Incubation with EGF had no major
effect on LEC proliferation (Figure 4.2.6 A) but significantly stimulated migration and tube
formation (Figure 4.2.6 B-C). Because small molecule tyrosine kinase inhibitors may inhibit a
number of tyrosine kinases, in addition to their most specific target, we next used a specific
EGFR blocking antibody (EGFRbAB) and an ErbB2 heterodimeryzation blocking antibody
(pertuzumab). The stimulating effects of EGF on cell migration were significantly inhibited by
EGFRbAB and by pertuzumab, and even more potently by a combination of both antibodies
(Figure 4.2.6 B). GW2974 also inhibited the EGF effect (Figure 4.2.6 B). No major effect on
LEC migration was observed after incubation with either the blocking antibodies alone, with
GW2974 or with the IgG controls (Figure 4.2.6 B). Similarly, the stimulating effect of EGF
on LEC tube formation was inhibited by EGFRbAB and by pertuzumab (Figure 4.2.6 C).
GW2974 also inhibited tube formation alone or in combination with EGF (Figure 4.2.6 C).
The blocking antibodies alone or the IgG controls had no major effects on tube formation
(Figure 4.2.6 C).
93
Figure
4.2.6
EGF,
via
EGFR/ErbB2,
promotes human lymphatic endothelial cell
migration and tube formation in vitro.
Human dermal lymphatic endothelial cells
(hLECs) were used to study cell proliferation
(A), migration (B) and tube formation (C).
Incubation with VEGF-A or VEGF-C promoted
hLEC proliferation/migration (A, B) and tube
formation (C), respectively, as compared with
controls. EGF did not induce cell proliferation
(A) but it significantly induced hLEC migration
and tube formation as compared with controls
(B, C). The EGF effect was inhibited by
GW2974, by an EGFR blocking antibody
(EGFRbAB), by pertuzumab and by the
combination of the two antibodies (B, C).
GW2974 alone significantly inhibited tube
formation (C). The blocking antibodies alone as
well as the IgG control did not affect hLEC
migration (B) or tube formation (C). Data are
expressed as mean values (n=3) + SEM;
*p<0.05;
**p<0.01;***
p<0.001.
AFU:
absolute fluorescent units.
94
4.2.4.4 EGF promotes lymphangiogenesis in vivo
We next investigated whether EGF might also have an effect on lymphatic vessel formation in
vivo. To this end, matrigel plugs containing EGF, VEGF-C or vehicle alone were injected
subcutaneously into the left inguinal area of wild type FVB mice. After 2 weeks, the matrigel
plugs and the associated skin were dissected and analyzed. We found a significantly increased
lymphatic vessel area (p=0.014), increased lymphatic vessel size (p=0.53) and increased
lymphatic vessel density (p=0.073) in the skin surrounding matrigel plugs that contained EGF
(Figure 4.2.7 A-C), as compared to those that contained PBS. The effects of EGF were largely
comparable to those of VEGF-C (Figure 4.2.7 A-C).
Figure 4.2.7 EGF promotes lymphangiogenesis in vivo. A matrigel plug containing PBS, VEGF-C or EGF
was injected subcutaneously into the left inguinal area of wild type mice. After 2 weeks, the plug and the
associated skin were dissected and quantitatively analyzed. VEGF-C significantly increased lymphatic vessel
area (A) and size (B) as compared with control PBS; the lymphatic density was not significantly increased (C).
EGF treatment significantly induced lymphatic vessels area (A) and it slightly increased size (B) and density (C).
Data are expressed as absolute values per matrigel plug; bar represents the median value.
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4.2.4.5 Transgenic
overexpression
of
amphiregulin
in
mouse
skin
promotes
lymphangiogenesis in vivo
Amphiregulin is another specific ligand for EGFR/ErbB2. Recently, it has been shown that
transgenic mice that express amphiregulin under control of the K14 promoter in the skin,
develop a chronic, psoriasis-like skin inflammation early during life (Cook et al. 1997). We
next investigated whether chronic transgenic overexpression of amphiregulin in the mouse
skin might also promote lymphangiogenesis. We found an increased lymphatic vessel area
(Figure 4.2.8 A), size (Figure 4.2.8 B) and density (Figure 4.2.8 C) in both the ear skin and the
tail skin of seven-days old transgenic mice, as compared with wild type mice of the same age.
Immunohistochemical analysis of LYVE-1 expression in the tail skin (Figure 4.2.8 D, E) and
ear skin (Figure 4.2.8 F, G) revealed increased numbers of enlarged dermal lymphatic vessels
in K14/amphiregulin transgenic mice (Figure 4.2.8 E, G) as compared with wild type mice
(Figure 4.2.8 D, F).
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Figure 4.2.8 Transgenic overexpression of amphiregulin in mouse skin promotes lymphangiogenesis.
Seven-day old K14-amphiregulin (AR) transgenic mice presented an increased lymphatic vessel area (A), size
(B) and density (C) in both ear skin and tail skin, as compared with wild type (WT) mice (the increase is not
significant for lymphatic size in the tail sample, B). Immunohistochemical analysis for LYVE-1 in the tail (D, E)
and ear (F, G) skin showed enlarged and more dense dermal lymphatics in the transgenic mouse samples (E, G)
as compared with wild type animals (D, F arrow heads. Data are expressed as absolute values per animal, bar
represents the mean value. Scale bars 50 μm.
97
4.2.5
Discussion
In a search for novel pathways involved in lymphatic vessel growth, we have investigated a
number of small molecular organic compounds for their effects on vascular development in
mouse embryonic stem cell-derived embryoid bodies (EBs). The results indicated that
epidermal growth factor receptor (EGFR/ErbB2) signalling might regulate lymphatic vessel
formation and conversely that EGF, via EGFR/ErbB2, promotes vessel growth. We also found
that human dermal LECs express low levels of EGFR both in vitro and in vivo and that
signalling of EGF via EGFR/ErbB2 promotes human LEC migration and tube formation.
Importantly, two independent in vivo studies reveal that EGFR signalling also promote
lymphangiogenesis in vivo.
Recently, a forward chemical genetics approach, using a chemical library containing 1,280
small organic molecules with well-defined pathway specificities, was applied to Xenopus
laevis tadpoles (Kalin et al. 2009). A phenotypic screen revealed compounds affecting either
blood vessel formation, lymphatic vessel formation or both. Among the compounds with
effects on vascular development, SU5416 (targeting the tyrosine kinase associated with
VEGFR-2) potently inhibited blood vessel formation, whereas GW2974, an EGFR/ErbB2
tyrosine kinase inhibitor, and the calcium channel blockers felodipine and nicardipine,
predominantly inhibited lymphatic vessel development. Genistein, another EGFR/ErbB2
tyrosine kinase inhibitor, affected both blood and lymphatic vessel formation. The lymphatic
phenotypes were associated with impaired lymph sac and lymph heart formation, and with
hypoplasia of the posterior lymphatic vessels (Kalin et al. 2009). These results indicated a
role of EGFR signalling in developmental lymphangiogenesis. However, the small organic
tyrosine kinase inhibitors also target other tyrosine kinases than EGFR, and the potential role
98
of EGF/EGFR signalling for the growth and function of lymphatic vessels after
embryogenesis has remained unknown.
Mouse embryonic stem cell-derived EBs have been used to study (anti-)angiogenic properties
of small molecules and drugs (Wartenberg et al. 1998). Recently, we found that also
lymphatic vessel-like structures develop in EBs and that incubation of EBs with the
lymphangiogenic growth factor VEGF-C promotes formation of CD31+/LYVE-1+ lymphatic
vessel like structures (Liersch et al. 2006), indicating that EBs might be useful tools for
identifying compounds potentially modulating lymphangiogenesis. We tested the selected
compounds alone or in combination with a mixture of retinoic acid (RA), cAMP and VEGFC, since we recently found that RA in combination with cAMP potently promote the
development of lymphatic vessel-like structures in EBs (Marino et al., submitted). Although
the calcium channel blocking compounds felodipine and nicardipine, as well as genistein, also
reduced the number of CD31+/LYVE-1+ vessel-like structures in EBs, incubation with
GW2974 showed the strongest inhibitory effect in the EBs. In fact, GW2974 almost
completely prevented the formation of lymphatic vessel like structures, and only single
CD31+/LYVE-1+ cells were observed, even when EBs were incubated in the
RA/cAMP/VEGF-C mixture.
EGFR family proteins (Riese and Stern 1998) play important roles in the regulation of cell
growth, proliferation, and differentiation of various cell types. In particular, mice lacking the
EGFR (ErbB1) die before implantation or at mid-late gestation (depending on the genetic
background), due to placental malformations and severe defects of various epithelial cells
(Wong 2003). In particular, it has been shown that EGFR may regulate angiogenesis both
directly (Baker et al. 2002) and indirectly (Kumar and Yarmand-Bagheri 2001). Moreover,
99
EGFR ligands such as EGF and TGFα are potent pro-angiogenic factors (Schreiber et al.
1986), in part via upregulation of VEGF-A expression in epithelial cells (Detmar et al. 1994)
The EGFR has been reported to be expressed by quiescent and by tumor-associated blood
vascular endothelial cells (Sini et al. 2005; Amin et al. 2006). Importantly, we detected EGFR
expression in human dermal lymphatic endothelial cells both in vitro and in vivo. These
findings indicate that the effects of EGF (and of amphiregulin) on lymphatic vessel activation
are, at least in part, mediated directly via EGFR activation. Recently, it has been reported that
proliferation of cultured mouse mesenteric LEC was induced by EGF (via Akt and ERK1/2)
and inhibited by AEE788, a small molecule that targets EGFR and VEGFR2 TKs (Rebhun et
al. 2006). EGF has also been reported to stimulate proliferation of bovine LECs (Liu and He
1997). In contrast, we did not observe any significant effect of EGF treatment, or of
incubation with the EGFR inhibitor GW2974, on the in vitro proliferation of human dermal
lymphatic endothelial cells in the present study. These differences might be due to the
different culture conditions, to different tissue origin of the LECs, or to species-specific
differences in the response of lymphatic endothelium to growth stimulation. Importantly,
however, our in vitro studies revealed that activation of the EGFR potently promotes LEC
migration and tube formation, and that - conversely - blockade of EGFR signalling via direct
antibody inhibition and via tyrosine kinase inhibition reduce migration and tube formation.
Our study is the first to reveal that activation of the EGFR promotes lymphangiogenesis in
vivo. We have used two independent experimental mouse models to investigate the role of this
receptor in lymphatic vessel formation. Implantation of an EGF containing matrigel plug
resulted in an increased number of enlarged lymphatic vessels in the surrounding skin after 14
days. The matrigel plug assay is a widely used and established (lymph)-angiogenic assay.
However, implantation also leads to an inflammatory response of the host tissue, with possible
100
confounding effects by the inflammatory cells. Thus, we investigated a second mouse model
with chronic overexpression of another EGFR ligand, amphiregulin, by transgenic
keratinocytes within the epidermis. We initially studied older K14/amphiregulin mice at an
age of 3 weeks, and found dramatic enlargement of the cutaneous lymphatic vasculature (data
not shown). However, at this time point, the mice had developed a strong, psoriasis-like skin
inflammation, in agreement with the published phenotype (Cook et al. 1997). Because
psoriasis skin lesions in humans, and psoriasis-like skin lesions in mice are associated with
massive lymphatic hyperplasia (Kunstfeld et al. 2004), we then studied the vasculature of
young, 1-week old K14-amphiregulin transgenic mice (at a time point when there was no
massive inflammation detectable). These investigations revealed that overexpression of
amphiregulin promoted dermal lymphangiogenesis. Taken together, the results indicate a role
of EGFR signalling in lymphatic vessel growth in vivo.
Is the EGF effect on lymphatic endothelial cells mediated directly or indirectly? We were
unable to detect EGFR expression neither on mRNA level in ex vivo isolated mouse colon
lymphatic endothelial cells nor on protein level in mouse dermal lymphatic vessels (data not
shown); thus, the pro-lymphangiogenic effect of EGF and amphiregulin in vivo might be - at
least partially - due to an indirect effect mediated by other cells types that might also be
present in the EB assay. In fact, it has been shown that EGFR/ErbB2 signalling leads to the
expression of angiogenic growth factor, such as VEGF-A in keratinocytes and cancer cells
(Detmar et al. 1994; Dvorak et al. 1995; Kumar and Yarmand-Bagheri 2001; Al-Nedawi et al.
2009). In contrast, the effect of EGF on the migration and tube formation of human dermal
LECs was most likely mediated directly since these cells express the EGFR and since no other
cell type was present in these cultures. Moreover, the levels of VEGF-A and VEGF-C were
below detection limit in LEC cultures treated with EGF, as evaluated by ELISA (data not
shown).
101
EGFR and ErBb2 are frequently upregulated in human carcinomas and support proliferation
and survival of cancer cells (Yarden and Sliwkowski 2001). Small molecule inhibitors such as
erlotinib and blocking antibodies such as cetuximab, that target EGFR, are in clinical use for
some types of cancer such as lung cancers (Herbst and Sandler 2008). Inhibition of tumor
associated-angiogenesis by EGFR blockade has been reported (Petit et al. 1997; Perrotte et al.
1999), and combination therapies blocking EGF and VEGF signalling are currently under
investigation as a potential strategy to overcome resistance to anti-angiogenic therapy.
Importantly, patients who undergo EGFR blocking treatment often develop an acute skin rush
and skin inflammation (Tsimboukis et al. 2009), and development of peripheral edema has
been reported (Perez-Soler and Saltz 2005; Von Minckwitz et al. 2005). Based upon our
results, analysis of patient biopsies would be of great interest to identify possible effect of
EGFR inhibition on tumor-associated lymphangiogenesis.
In conclusion, the present study indicates that small inhibitory molecules (e.g. erlotinib) or
blocking antibodies (e.g. cetuximab, pertuzumab) targeting EGFR/ErbB2 signalling might be
potential candidates for the treatment of pathological lymphangiogenesis in cancer and other
diseases.
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5 CONCLUSIONS AND OUTLOOK
The present work has aimed at identifying molecular regulators of the earliest steps of lymphatic
vascular development. So far, this task has proven challenging using classical in vivo genetic
manipulation techniques due to the fact that several candidate gene mutations have resulted in
mouse embryo lethality. The use of mouse embryonic stem cell-derived embryoid bodies, an in
vitro model that largely mimics mouse embryogenesis, has allowed us to circumvent this
problem.
Using the embryoid body vascular differentiation assay, we have identified retinoic acid (RA) as
a novel mediator of lymphatic competence and commitment. RA alone, and in combination with
cAMP, potently up-regulated LYVE-1 expression in CD31+ vascular structures. Indeed, these
LYVE-1+, lymphatically competent endothelial cells emerged from the pre-existing CD31+
blood vascular endothelium, supporting the centrifugal theory of lymphatic development
proposed by Sabin in 1902 (Sabin 1902). However, the combination of RA and cAMP also
induced the formation of rare CD31+/LYVE-1+/Prox1+ cell clusters that emerged
independently from the CD31+ vessel-like networks. Thus, also supporting the centripetal
theory proposed by Huntington in 1910 (Huntington and McClure 1910). Our studies also
demonstrated that retinoic acid receptor (RAR)-was expressed by the endothelial cell of the
embryonic cardinal veins in mouse embryos and that Ro 41-5253, a RAR- antagonist
inhibited the induction of lymphatic vascular structures by RA, suggesting that RA activity is
most likely mediated via RAR-α. The potentiation of RA’s effects by cAMP and the inhibition
by H89 (a cAMP-dependent PkA inhibitor) indicate that RA’s effects are mediated by a
pathway that also involves cAMP and PkA. Importantly, in mouse embryos increased levels of
RA in utero upregulated LYVE-1 expression in the endothelial cells of the cardinal veins and
103
jugular lymph sacs, and increased the number of CD31+/Prox1+ lymphatic endothelial cells. In
contrast, in utero injection of Ro 41-5253 decreased expression of LYVE-1 and the number of
CD31+/Prox1+ lymphatic endothelial cells in the same vascular regions.
Moreover, exposure of Xenopus embryos to RA induced the expression of VEGFR-3 and
LYVE-1, thus conferring lymphatic competence to the developing vasculature.
Retinoids, such as 13-cis-RA, are widely used via both topical (Orfanos et al. 1997) and
systemic administration (DiGiovanna 2001) for the treatment of various conditions such as
acne, psoriasis, postinflammatory hyperpigmentation, photodamaged skin, leukemia, disorders
of keratinisation, premalignant skin lesions, cutaneous T-cell lymphoma, and dissecting
cellulitis of the scalp (Saurat 1998). Recently, retinoids are being considered as agents to treat
autoimmune diseases, such as multiple sclerosis (Pino-Lagos et al. 2008) and Alzheimer’s
disease (Shudo et al. 2009), due to the fact that they can, in general, exert immunosuppressive
effects. Importantly, therapeutic efficacy of retinoids has been reported for the treatment of
persistent solid facial edema, a complication of acne vulgaris (Thielitz and Gollnick 2008;
Manolache et al. 2009). In contrast, retinoic acid syndrome, a clinical syndrome that occurs after
treatment of leukemia (Leelasiri et al. 2005) with all-trans-retinoic acid, is very often associated
with edema formation and weight gain. Since lymphatic vessels are known to be involved in
inflammatory processes, as well as in edema formation and obesity (Cueni and Detmar 2008),
this clinical evidence suggests a possible link between RA treatment and lymphatic vessel
growth and function. Indeed, preliminary results in a pilot study on adult mice indicate that
intraperitoneal injection of RA induces dermal lymphangiogenesis (data not shown). Thus, it
will be of great interest to determine whether retinoids might modulate the growth of new
lymphatic vessels in pathological conditions like lymphedema. In this respect, mouse models of
e.g. tail lymphedema (Jin et al. 2008) might represent a suitable in vivo approach to evaluate the
effect of RA on the regression of the pathology.
104
The second part of this thesis focused on using the embryoid body model to identify novel
(anti)-lymphangiogenic pathways. To this end, embryoid bodies were treated with small organic
molecules with well-defined pathway specificities, and the formation of lymphatic vessel-like
structures was evaluated. The screening identified the small molecule GW2974, which targets
the tyrosine kinase associated with epidermal growth factor receptor (EGFR/ErbB2), as an
inhibitor of lymphatic vessel formation. Conversely, embryoid bodies treated with EGF, an
activating ligand for EGFR/ErbB2, displayed increased number of lymphatic vessel networks as
compared to control embryoid bodies. We found that human dermal lymphatic endothelial cells
(hLECs) express low levels of EGFR. GW2974, as well as EGFR/ErbB2 blocking antibodies,
inhibited in vitro migration and tube formation of hLECs whereas EGF significantly induced
those processes. In vivo analysis of mice subcutaneously implanted with EGF containing
matrigel plugs, as well as of mice expressing amphiregulin (another EGFR/ErbB2 ligand) under
the control of K14 promoter, revealed that the activation of the EGFR/ErbB2 signalling pathway
induces dermal lymphangiogenesis. Taken together, our results suggest that targeting EGFR
might represent a novel strategy to interfere with lymphangiogenesis in pathological conditions.
Further in vivo studies, such as blocking EGFR in tumor bearing mice, could prove valuable in
defining the therapeutic potential of EGFR antagonists to inhibit tumor metastasis. Moreover,
since small molecules such as erlotinib and blocking antibodies such as cetuximab that target
EGFR are already in clinical use for some types of cancer such as lung cancers (Herbst and
Sandler 2008), the analysis of patient tumor biopsies would be of great interest to identify a
possible effect of EGFR inhibition on tumor-associated lymphangiogenesis.
In conclusion, mouse embryonic stem cell-derived embryoid bodies have allowed us to discover
novel processes regulating lymphatic differentiation and lymphangiogenesis. These findings
open up the way for future research projects with the aim of better understanding the molecular
mechanisms that control growth and function of the lymphatic vascular system.
105
6 ACKNOWLEDGMENTS
I wish to thank Michael Detmar for giving me the opportunity of doing my PhD in such an
interesting field and at such an honourable institution as ETH. Thanks for patiently
accompanying me through the fabulous and exciting world of science.
Many thanks go to Prof. Cornelia Halin for being co-referee of my thesis and for her always
helpful suggestions.
I thank Prof. André Braendli and Dr. Vasilios Debouras for providing the Xenopus techniques,
helping in performing the experiments and for revising the manuscript: “A role for all-transretinoic acid in the early steps of lymphatic vascular system development”.
I would like to thank my two students Yvonne Angehrn and Sabrina Riccardi for their
dedication to the “EB factories” and the great time together inside and outside the lab!
For the study: “Activation of the epidermal growth factor receptor promotes lymphangiogenesis
in vitro and in vivo” I thank Dr. Baenziger-Tobler, Yvonne Angehrn and Sabrina Riccardi for
their contribution, Prof. M. Pittelkow for providing the K14-AR mouse samples and Mr. M.
Hasmann (Roche Diagnostics GmbH) for providing the human ErbB2 antibody, pertuzumab.
Special thanks go to Deena and Steven, our precious native English speaking post docs that
“volunteered” to correct this thesis, fishing and hunting out Italian expressions and misplaced
commas!
Immense thanks go to the wonderful “Detmar&Halin lab”. All present and former members
have not just been my colleagues: they have been and are my “Swiss Family”, my irreplaceable
106
friends. Exceptional thanks go to Giorgia, Jay, Christian, Matthias, Tanja, Max, Steven, Martin,
Benni and Masha. Those four years with you guys have been unforgettable. Moreover,
uncountable thanks go to one lab member above all: my office-mate and best friend Milica that
intensively shared with me Ph.D. happiness and frustration as well as very precious life
moments.
At last but not least I would like to thank my beloved fiancé Gianluca, my little sister Roberta
and my parents for always believing in me and supporting any of my (crazy or intelligent)
decisions and ideas. Without them I would not be who, what and where I am! I love you all so
much!
107
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8 CURRICULUM VITAE
Daniela Marino
Neunbrunnenstrasse 215, 8046 Zurich, Switzerland
Telephone: +41 44 633 7326 (Office); +41 76 455 31 54 (Mobile)
Email: [email protected]; [email protected]
Personal Data
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Date of birth: November 17, 1981
Place of birth: Agrigento, Italy
Nationality: Italian
Marital status: Unmarried
Education
09/2005-present
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Swiss Federal Institute of Technology (ETH) Zurich, Switzerland
07/2005
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Bachelor in Biotechnology
University of Milan, Italy
Thesis: “An hybrid biologic-electronic neural network experiment and analysis”
Stem Cell Research Institute (SCRI), San Raffaele Hospital, Milan, Italy
07/2000

Master in Biotechnology, Functional Genomics and Bioinformatics
University of Milan, Italy
Thesis: “Study and analysis of learning, differentiation and proliferation of human neural stem cells”
Stem Cell Research Institute (SCRI), San Raffaele Hospital, Milan, Italy
07/2003
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PhD studies under supervision of Prof. Michael Detmar
High school graduation
Liceo Scientifico Statale, A. Sciascia, Canicattì, Agrigento, Italy
Work Experience
09/2005-present


PhD thesis project in the group of Prof. Michael Detmar
Swiss Federal Institute of Technology (ETH) Zurich, Switzerland
Thesis: “Identification of molecular mechanisms controlling lymphatic vessel formation by use of the
embryonic stem cell-derived embryoid body assay”.
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08/2004-09/2004

Teaching of molecular and cell biology, genetics, evolution and anatomy for high
school students
Study center, G. Toniolo, Canicattì, Agrigento, Italy
Further Education and Workshops
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BD-Swiss Tissue Culture Society workshop, “Cell culture models in Pharma Research”. Basel,
Switzerland. November, 2008.
“Genetics, breeding and monitoring of mutant mice”. University of Zurich, Switzerland. November,
2007
Laboratory Animal Science, theoretical and practical training in laboratory animal handling.
University of Zurich, Switzerland. September, 2006.
PALM MicroBeam workshop. Istituto Oncologico della Svizzera Italiana (IOSI), Bellinzona,
Switzerland. February, 2006.
“In vitro culture, Embryos, ES –cells, Primary Cells”. University of Zurich, Switzerland. October, 2005.
Other Professional Activities
2006-2009
Head of the social committee of the Pharmaceutical Scientists’ Association (PSA) at the
Institute of Pharmaceutical Sciences, ETH Zurich, Switzerland
2006-2008
Teaching assistant, yearly one-week practical course in medicinal chemistry for Bachelor
students in Pharmaceutical Sciences at the Institute of Pharmaceutical Sciences, ETH Zurich,
Switzerland
2007
Supervisor of semester- and diploma-projects of students in Pharmaceutical Sciences
Publications and Patent

R.M.R. Pizzi, D. Rossetti, G. Cino, D. Marino, A.L.Vescovi and W. Baer (2009). A cultured human
neural network operates a robotic actuator. Biosystems 95, 137-144.

D. Marino, V. Dabouras, A.W. Braendli and M. Detmar. A role for all-trans-retinoic acid in the early
steps of lymphatic vascular development. Submitted

D. Marino, Y. Angehrn, S. Riccardi, N. Baenziger-Tobler, M.R. Pittelkow and M. Detmar. Activation of
the epidermal growth factor receptor promotes lymphangiogenesis in vitro and in vivo. Manuscript in
preparation.

D. M. Leslie-Pedrioli, V. Dabouras, T. Karpanen, G. Jurisic, J.W. Shin, D. Marino, R. E. Kalin, S.
Leidel, P. Cinelli, S. Schulte-Merker, A. W. Braendli and M. Detmar. miR-31 inhibits lymphatic lineagespecific differentiation in vitro and lymphatic vessel development in vivo. Submitted.

L. Cueni, L. Chen, D. Marino, R. Hunggenberger, A. Alitalo and M. Detmar. Podoplanin-Fc reduces
lymphatic vessel formation in vitro and in vivo and causes disseminated intravascular coagulation when
transgenically expressed in the skin. In revision.
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
Pending patent: Multiwell culture plate for three-dimensional cultures. Priority date: 20.02.08.
http://www.wipo.int, patent database, code: WO/2009/103416
Conference Presentations
With ORAL presentation:

SSCN - Swiss Stem Cell Network annual meeting
D. Marino, Y. Angehrn, M. Detmar. “Lymphatic endothelial cell differentiation of mouse embryonic
stem cells”
Organizer: Prof. Lukas Sommer; University of Zurich, Switzerland. December 14, 2007.
With POSTER presentation:

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SSCN- Swiss Stem Cell Network annual meeting
D. Marino, V. Dabouras, A.W. Braendli and M. Detmar. “All-trans-RA is involved in the early
processes of lymphatic vascular system development”.
Organizers: Brigitte Galliot, Marcos Gonzalez Gaïtan; University of Geneva, Switzerland. February 6,
2009.
ESDR – European Society for Dermatological Research
D. Marino, R. Liersch, M. Detmar. “Induction of lymphatic endothelial cell differentiation in mouse
embryonic stem cells”
Organizer: Reinhard Dummer; University of Zurich, Switzerland. September 5–8, 2007.
KEYSTONE Symposium - Stem Cells and Cancer
D. Marino, R. Liersch, M. Detmar. “Induction of lymphatic endothelial cell differentiation in mouse
embryonic stem cells”
Organizers: Irving L. Weissman and Michael F. Clarke; Keystone Resort, Keystone, Colorado, USA.
March 2-7, 2007.
EuroStemCell International Conference - Advances in Stem Cell Research
D. Marino, R. Liersch, M. Bruderer, C. Bader, M. Detmar. “Lymphatic endothelial cell differentiation of
mouse embryonic stem cells”
Organisers: Yann Barrandon, Austin Smith, Elena Cattaneo; Ecole Polytechnique Fédérale de Lausanne,
Switzerland. September 8-10, 2006.
ESH - European School of Haematology, Euroconference on Angiogenesis
D. Marino, R. Liersch, M. Bruderer, C. Bader, M. Detmar. “Lymphatic endothelial cell differentiation of
mouse embryonic stem cells”
Organizers: K. Alitalo, Ch. Betsholtz, A. Bikfalvi, C. Rüegg, G. Tobelem; Mandelieu/Cannes, France.
May 13-16, 2006.
Outside Interestes and Activities

Sports (aqua-fitness, yoga, volleyball, kickboxing, snowboarding), travelling, cooking, reading, singing,
new technologies.
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