PhD thesis

Cand.scient. Janny Marie Landegent Peterslund

FGF-signalling in the differentiation of mouse ES cells towards definitive endoderm

Academic advisors:

Dr. Berthe M. Willumsen, Department of Biology, Faculty of Science, University of Copenhagen – Denmark Dr. Palle Serup, Department of Stem Cell Biology, Hagedorn Research Institute – Denmark Submitted March 2010

"It is not birth, marriage, or death, but gastrulation, which is truly the most important time in your life." Lewis Wolpert (1986)

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Preface This thesis is based on experimental work performed in the Department of Stem Cell Biology (formerly the Department of Developmental Biology) at the Hagedorn Research Institute in Gentofte – Denmark, from August 2006 to March 2010 (including an 8 month leave). The project supervisors were Berthe M. Willumsen, Professor at the Institute of Biology, Faculty of Science, University of Copenhagen – Denmark, and Palle Serup, Ph.D., Scientific Director at the Hagedorn Research Institute, Gentofte – Denmark. The work was supported by grants from the Juvenile Diabetes Research Foundation and the Beta Cell Biology Consortium, National Institutes of Health – USA. This thesis is submitted in order to meet the requirements for obtaining the degree of philosophiæ doctor (Ph.D.) at the Faculty of Science, University of Copenhagen – Denmark. The thesis is based on two papers: Paper I:

‘A late requirement for Wnt and FGF signaling during Activin-induced formation of foregut endoderm from mouse embryonic stem cells’ Published in Developmental Biology, 2009; 330, pp. 286-304. Mattias Hansson, Dorthe R- Olesen, Janny M.L. Peterslund, Nina Engberg, Morten Kahn, Maria Winzi, Tino Klein, Poul Maddox-Hyttel and Palle Serup Paper II:

‘FGFR(IIIc)-activation induces mesendoderm but is dispensable for definitive endoderm formation in mouse embryonic stem cells’ Manuscript to be submitted. Janny Marie L. Peterslund and Palle Serup In addition, this thesis contains a general introduction to early mouse embryonic development and pancreas formation, directed differentiation of mES cells towards DE and pancreatic β-like cells, and FGF-signalling in relation to both development and differentiation. The reader will also find a short chapter expanding on the patterning of definitive endoderm described in paper I and a general discussion of my work in relation to the research field as a whole.

Janny Marie Landegent Peterslund March 2010

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Resumé Sukkersyge er en sygdom der på verdensplan påvirker mere end 200 mio. mennesker. Transplantation med insulinproducerende β-celler anses for en mulig fremtidig kur mod sukkersyge, men da antallet af donorer er begrænset er der brug for andre kilder til cellemateriale. Embryonale stamceller (ESC’er) er pluripotente celler, hvilket betyder at de har potentiale til at kunne differentiere til alle celle- og vævstyper i den voksne krop. De vil som sådan kunne være en ubegrænset kilde til in vitro-differentierede β-lignende celler. Det nærværende arbejde beskriver hvordan muse-ESC’er, der bruges som model for humane ESC’er, bliver induceret til at differentiere i de tidlige stadier mod β-lignende celler. Arbejdet fokuserer på fibroblast vækstfaktor (FGF)-signalering i forhold til dannelsen af mesendoderm, definitiv endoderm (DE) og ’posterior foregut endoderm’. Mesendoderm er en bipotent cellepopulation som kan differentiere til enten DE eller mesoderm ved activinA eller bone morphogenetic protein (BMP). DE og ’posterior foregut endoderm’ er mere modne celletyper der senere danner bugspytkirtlen og β-celler. Dette arbejde er baseret på et serum- og feederfrit cellekultursystem og viser hvordan induktion af mesendoderme celletyper med BMP4 eller forskellige activinA-koncentrationer afhænger af FGF-signalering. Blokering af FGF-signalering med opløselige FGF receptorer (FGFR’er) eller småmolekylære hæmmere hindrer dannelsen af mesendoderm, hvorimod tilsætning af en række FGF’er øger dens dannelse. Mere præcist er det aktivering af FGFR(III)c (FGFRc)-isoformer der giver effekten, mens aktivering af FGFRb-isoformer ikke har nogen effekt. Dette stemmer overens med udtrykket af især FGFRc-isoformer i tidlig differentiering af ESC’er. På den anden side reducerer høje FGF-koncentrationer antallet af mere modne DE celler, men aktiv FGFR-signalering generelt er dog nødvendig for DE-dannelse. En FGF4 knockout cellelinje, der tidligere har vist sig ikke at kunne differentiere til ektoderm og mesoderm kimlagstyper, kunne meget overraskende sagtens differentiere til DE uden tilsætning af FGF4-protein. Den heterozygote cellelinje FGF4+/– viser endda højere udtryk af DE markørgener end både vildtype og knockout cellelinjerne, hvilket tyder på at et mellemliggende niveau af FGF-signalering er fremmende for DE dannelse. Alt i alt tyder disse resultater på at DE opnås bedst i muse-ESC’er ved tidlig aktivering af FGFRc-isoformer i mesendodermdannelse efterfulgt af FGFRc-aktivering under endogene niveauer. Endeligt afhænger en videre differentiering af DE cellepopulationen af retinoic acid og FGFsignalering, helst ved FGF7 og FGF10, der aktiverer FGFRb-isoformer. En middelkoncentration af FGF resulterer i mange celler der udtrykker en ’anterior foregut’markør, få celler der udtrykker en ’posterior foregut’ markør og ingen celler der udtrykker en ’hindgut’-markør. Høje koncentrationer af FGF får DE til at danne ’hindgut’. Disse resultater giver ny viden om FGF-signalerings forskelligartede effekt på mesendoderm-, DE- og ’posterior foregut endoderm’-dannelse over tid. Det vil sandsynligvis være værdifuldt for slutmaterialet, nemlig β-lignende celler, at overføre denne viden til nuværende differentieringsprotokoller.

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Summary Diabetes is a metabolic disease affecting more than 200 million people worldwide. Therapeutic transplantation with insulin-producing β cells is envisioned as a cure for diabetes, but as donors are scarce, other sources of cell material are needed. Embryonic stem (ES) cells are pluripotent cells with the potential to differentiate into all cell and tissue types of the adult body. As such, they could serve as an unlimited source of in vitro differentiated β-like cells. The present work describes how mouse ES cells, which are used as a model for human ES cells, are induced to differentiate in the early steps towards β-like cells. The work focuses on fibroblast growth factor (FGF)-signalling in relation to the formation of mesendoderm, definitive endoderm (DE) and posterior foregut endoderm. The mesendoderm is a bipotent cell population, which can differentiate into either DE or mesoderm by activinA or bone morphogenetic protein (BMP). The DE and posterior foregut endoderm are more mature cell types which will later form the pancreas and β cells. This work is based on a serum and feeder-free cell culture system, and shows that induction of mesendoderm cell types by BMP4 or a range of activinA-concentrations depends on FGFsignalling. Blocking FGF-signalling by soluble FGF receptors (FGFRs) or small molecule inhibitors impedes formation of the mesendoderm, whereas addition of a range of FGFs increases its formation. More specifically, activation through FGFR(III)c (FGFRc)-isoforms elicits the effect, whereas activation of FGFRb-isoforms shows no effect. This correlates with the expression of especially FGFRc-isoforms in early differentiation of ES cells. On the contrary, high concentrations of FGFs reduce the number of more the mature DE cells, although active FGFR-signalling in general is necessary for DE-formation. Surprisingly, an FGF4 knockout cell line which has previously been reported to be unable to differentiate into ectoderm and mesoderm cell lineages can readily differentiate into DE without addition of ectopic FGF4. The heterozygote cell line FGF4+/– even shows higher expression of DE marker genes than both wild type and knockout cell lines, indicating that an intermediate level of FGF-signalling is beneficial to DE-formation. Together, these results suggest that activation of FGFRc-isoforms during early mesendoderm specification followed by FGFRc activation below endogenous levels is the most attractive recipe to obtain DE formation in mouse ES cells. Lastly, further differentiation of the DE cell population is dependent on retinoic acid and FGFsignalling, preferably by FGF7 and FGF10 which activate FGFRb-isoforms. An intermediate concentration of FGF results in a high number of cells expressing an anterior foregut marker, few cells expressing a posterior foregut marker and no cells expressing a hindgut marker. High concentrations of FGF posteriorize the DE to form hindgut. These results provide new insight into the differential effect of FGF-signalling on mesendoderm, DE and posterior foregut endoderm formation over time. Applying this knowledge to current protocols for differentiation may prove beneficial to the end-point material, namely β-like cells.

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Table of contents

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Abbreviations AP1 AIP AKT ALK4 A-P AVE BMP BSA Cdx2 CIP Cxcr4 DAPI DE Dkk1 DTT E EB EdU EpCAM EpiSC EPL ERK1/2 ES Evx1 ExE FACS FGF FGFR FGFRb/c Flk1 Foxa2 GAPDH GATA6 GFP Gp130 Gsc GSK3 hES cells Hesx1 Hex Hlxb9 HS

activator protein 1 anterior intestinal portal akt; protein kinase B activin receptor-like kinase 4 anterior-posterior anterior visceral endoderm bone morphogenetic protein bovine serum albumin Homeobox transcription factor Cdx2 caudal intestinal portal Chemokine (C-X-C motif) receptor 4 4′,6-diamidino-2-phenylindole definitive endoderm Dickkopf 1 dithiotreitol embryonic day embroid body 5-ethynyl-2’-deoxyuridine epithelial cell-adhesion molecule epiblast stem cell early primitive ectoderm-like extracellular signal-regulated kinase1/2 embryonic stem Even-skipped homeobox homolog 1 extra-embryonic ectoderm fluorescence-activated cell scanner fibroblast growth factor fibroblast growth factor receptor fibroblast growth factor(IIIb)/(IIIc) Fetal-like kinase 1 Forkhead transcription box 2 glyceraldehyde 3-phosphate dehydrogenase GATA-binding protein 6 green fluorescent protein glycoprotein 130 Gooseciod glycogen synthase kinase 3 human embryonic stem cells Homeobox expressed in ES cells 1 Hematopoietically expressed homeobox Homeobox transcription factor HB9 heparin sulfate

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ICM Id iPS JAK JDRF LIF MAPK m-Cer1 MEK mES cells MHC Mixl1 Ngn3 Nkx6.1 Oct4 Otx2 PCR PDGFR Pdx1 PE PLCγ PP PI3K Ptf1a qPCR RA RT-PCR S.D. S.E.M. Shh SMAD Sox SPRED STAT3 T TBP Tdh TE TGFβ VE WHO WNT3(a)

inner cell mass Inhibitor of differentiation induced pluripotent stem Janus kinase Juvenile diabetes research foundation leukemia inhibitory factor Mitogen-activated protein kinase mouse-Cerberus 1 a MAP kinase mouse embryonic stem cells major histocompatability complex Mix1 homeobox-like Neurogenin 3 NK homeobox transcription factor 6.1 Octamer-4 Orthodenticle homeobox 2 polymerase chain reaction platelet-derived growth factor receptor Pancreatic and duodenal homeobox factor 1 parietal endoderm phospholipase Cγ pancreatic polypeptide-producing phosphoinosityl 3 kinase Pancreas-specific transcription factor 1a quantitative polymerase chain reaction retinoic acid reverse transcriptase- polymerase chain reaction standard deviation standard error of the mean Sonic hedgehog proteins modulating the activity of TGFβ ligands; the name is a combination of the protein homologs ‘SMA’ (C. elegans) and ‘mosthers againts decapentaplegic’ (D. melanogaster) Sex determining region Y (SRY)-related HMG box Sprouty-related EVH1 protein signal transducer and activator of transcription 3 Brachyury TATA-binding protein Thermostable direct hemolysin gene trophectoderm Transforming growth factor β visceral endoderm World Health Organization wingless-type MMTV integration site 3(a)

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1. General introduction Diabetes mellitus Diabetes mellitus (diabetes hereafter) is caused by a lack of insulin-production or insulinresponsiveness, resulting in high blood glucose levels in the patient. There are two types of diabetes, type I and type II, resulting in an absolute or a relative lack of β cells, respectively (Donath and Halban 2004). Type II is the most common form of diabetes, accounting for 90 – 95% of all cases. The World Health Organization (WHO) estimated a worldwide prevalence of 171 million diabetics in 2000 and the prognosis is 336 million people by the year 2030, calling it a pandemic. In Denmark alone, 226.000 people (or 5% of the total population) had diabetes in 2006 and it is estimated that the Danish health care system spends DKK 22 billion (or app. US$ 4 billion) per year on treatment of diabetes and diabetes-related illness (Juvenile Diabetes Research Foundation homage). Furthermore, it is estimated that the disease costs Denmark an extra DKK 9 billion per year due to loss-of-production. In type I patients, the disease is a result of an auto-immune attack on the insulin-producing β cells, resulting in the loss of β cell mass, followed by dependence on insulin-treatment for the patient (see (Lernmark and Falorni 1998; Madsen 2005) for reviews). This dependency is life-long, as the β cell mass does not regenerate to levels where it can sustain the body’s need for insulin. The onset of type I diabetes is due to genetic and/ or environmental factors, but a comprehensive knowledge of the aetiology of the disease is still lacking despite intense studies thereof. Along with the primary disease which is treated with insulin injections, patients develop severe secondary complications such as blindness, kidney failure, and amputations due to chronic vascular defects caused by their blood-glucose levels being irregular and difficult to stabilize. In type II patients, a gradual insulin resistance of the peripheral tissues leads to an increase in β cell mass as a compensation for this condition (Rhodes 2005). This condition is partly reversible, but will ultimately lead to type I diabetes and insulin-dependence if not treated. Type II diabetes is mainly caused by genetic predisposition and environmental factors such as obesity, physical inactivity, excessive calorie intake and aging (Ling and Groop 2009).

Cell replacement therapy in diabetes Although there is no cure for diabetes at present, research focusing on therapeutic treatment is envisioned as a palliative treatment or maybe even a cure for the disease. This section will focus on therapies directed against type I diabetes. In 2000, Shapiro and co-workers published the Edmonton protocol, as it is now referred to, providing proof of principle for curing diabetes by transplanting donor islets of Langerhans and re-establishing eu-glycaemia in seven patients suffering from type I (Shapiro et al. 2000). However, 85% of islet recipients needed insulin treatment after a 5-year period (Ryan et al. 2005). The obstacles of auto-immunity along with the scarcity of islet donor material, has directed focus towards other sources of β cell material to put to use in a similar treatment. In type I diabetics, there is evidence of a continuous β cell regeneration taking place (Meier et al. 2005). However, attempts to regenerate the β cell mass are most likely overruled by the autoimmune attack, the gross result being no insulin-production from the islets of Langerhans. There is evidence that β-cells can be generated from existing cells, but it is unclear whether this is through replication of existing β cells or by re-differentiation of other pancreatic cells such as duct cells or even from hepatic cells (Bouwens and Pipeleers 1998; Yang et al. 2002a; Dor et al. 2004; Hardikar 2004; Xu et al. 2008; Borowiak and Melton 2009). It is debated whether the pancreas hosts a pancreatic endocrine stem cell that can be stimulated to proliferate (Madsen 2005). By partial pancreatectomy or cellophane wrapping of the pancreas, it has been demonstrated that β

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cell mass regenerates probably through an ability to ‘sense’ the total mass of insulin-producing cells and self-regulate accordingly (Hardikar 2004). Still, the success of these studies is low and such treatments will probably not be sufficient for curing diabetes. Replacement therapy by external sources of β-like cells holds the potential of an infinite supply of donor material for transplantation. The initial cell material may come from either somatic (or adult) stem cells or embryonic stem (ES) cells, or may in time come from patient-specific induced pluripotent (iPS) cells. These cell types are described in detail in a later section. The immune system represents a major obstacle when discussing cell replacement-therapies. Patients would either have to be put on a life-long treatment of immune-suppressive drugs, with many complications to follow, or other methods will have to be developed to overcome the attack of cell material by the recipient immune system (Rossini et al. 1999). Encapsulation of therapeutic cells is one such method. Despite limitations in nutrient supply to and transport of effectermolecules from the transplanted cells, studies in rat show that they can convey normo-glycaemia in steptozotocin-induced diabetic rats (Omer et al. 2005). Another possibility is induction of immunological tolerance (mixed chimaerism), which has been shown possible by bone-marrow transplantation in mice (Rossini et al. 1999; Blaha et al. 2005).

Embryonic development To appreciate the challenges of directed in vitro differentiation of ES cells, one must acquire a thorough understanding of embryonic development. In this thesis, focus will be on mouse development from zygote to pancreatic β cells, especially concentrating on the early development, as the work presented here is on mouse ES cell differentiation in the early steps towards β-like cells. From zygote to blastocyst

The fertilized egg, the zygote, is a totipotent cell from which all tissues of the embryo proper, the germ line and extra-embryonic tissues arise. In mice, the zygote moves into the uterine tract of the female and matures along the way. By embryonic day 3.5 (E3.5), it has developed into an oval structure, the blastocyst, consisting of an inner cell mass (ICM) at one end and the blastocoel at the other end. The ICM gives rise to i) the epiblast (also called the primitive ectoderm), which will later form the embryo proper and ii) facing the blastocoel, the primitive endoderm, which will give rise to the visceral and parietal endoderm (PE), forming parts of the yolk sac and uterine wall lining, respectively (Gardner 1983; Rossant 2004). Surrounding these structures is the trophectoderm (TE), which will develop into the foetal parts of the placenta, the chorion. At E4.5 the blastocyst implants into the uterine wall, gaining access to maternal blood vessels and thereby a supply of nutrients and oxygen. The ICM expresses Octamer-4 (Oct4; encoded by Pou5f1), which is down-regulated in the TE (Nichols et al. 1998) and Nanog, which is maintained in the epiblast but down-regulated in the PE (Mitsui et al. 2003). Oct4–/– and Nanog–/– embryos die due to a failure to form the ICM or the epiblast, respectively. Also, Sex determining region Y (SRY)-box 2 (Sox2) is present in the ICM, possibly as remnants of maternally deposited protein (Avilion et al. 2003). It is expressed by the embryo in the later epiblast and TE, where it, together with OCT4 and nanog induces transcription of pluripotency-associated genes. The TE expresses Cdx2, a homeobox transcription factor, which is required to form the placenta and to suppress transcription of Oct4 and Nanog (Chawengsaksophak et al. 2004; Strumpf et al. 2005). The PE expresses GATA-binding protein 6 (GATA6; (Morrisey et al. 1998)). Recent studies have shown that cells in the ICM express both nanog and GATA6 proteins prior to segregation into either epiblast or PE cells in a mosaic pattern (Chazaud et al. 2006). It is the expression level of fibroblast growth factor (FGF) that regulates each cell’s fate in such that low levels of FGF generate epiblast cell formation and high levels of FGF generate PE formation (Yamanaka et al. 2010). At this early stage, FGF4 and FGF receptor 1 (FGFR1) are expressed in the ICM and knockout mice of both genes die prior to or at gastrulation (Deng et al. 1994; Yamaguchi et al. 1994; Feldman et al. 1995). Fgf4 expression is activated by

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SOX2 and OCT4. Later, the PE and epiblast transiently express Fgf5 (Haub and Goldfarb 1991; Hebert et al. 1991). Gastrulation and germ-layer formation

At E5.5 the epiblast is formed as a cup-like structure surrounded by the visceral endoderm (VE) which patterns the epiblast and initiates gastrulation (Figure 1-1A; (Rossant 2004)). It is at the posterior, proximal part of this cup that the primitive streak (PS) forms around E6.5 from where it starts to migrate distally (Figure 1-1A). There is an anterior-posterior (A-P) patterning of the epiblast prior to gastrulation. Onset of PS formation is initiated by a gradient of the transforming growth factor β (TGFβ)-family member nodal and wingless-type MMTV integration site 3 (WNT3) signalling from the posterior epiblast. Nodal generates a proximal-to-distal gradient and WNT3a forms a posterior-to-anterior gradient (Liu et al. 1999; Ben-Haim et al. 2006; Arnold and Robertson 2009). The nodal gradient moves distally, forming the PS along the way and finally ends at the distal-most part of the embryo, the node (Gadue et al. 2005). At the late streak stage, nodal expression is only found in the node where it forms a distal-to-proximal signal gradient. The WNT3 expression pattern is restricted to the posterior epiblast during PS-formation. At the same time, the TGFβ family member bone morphogenetic protein 4 (BMP4) from the extra-embryonic ectoderm (ExE) signals to the adjacent epiblast. Thereby a proximal-to-distal signal gradient opposing that of nodal is formed (Lawson et al. 1999). The shape of these morphogenetic gradients is modulated by the reciprocal expression of antagonists or inhibitors, these being lefty and cerberus-like inhibiting nodal-signalling, dickkopf-related protein 1 (DKK1) inhibiting WNT3-signalling, and chordin and noggin inhibiting BMP4-signalling (Gadue et al. 2005). Thereby, the extension of the PS is restricted to the posterior side of the embryo. The primitive streak expresses the T-box transcription factor Brachyury (T) in migrating cells of the PS and Mix1 homeobox-like (Mixl1), along with Even-skipped homeobox homolog 1 (Evx1) (Figure 1-1A; (Bastian and Gruss 1990; Dush and Martin 1992; Kispert and Herrmann 1994; Ng et al. 2005)). Goosecoid (Gsc) is expressed in the progressing PS and localises to the anterior streak (Blum et al. 1992).

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Figure 1-1: Early embryo development from blastocyst stage to development through the primitive streak. A) Schematic representation of mouse embryonic development and signal gradients of BMP4, WNT, and nodal from blastocyst to late-streak stages. The red line in the late streak-stage embryo marks a cross-section, which is shown in B). Names of marker genes expressed in a tissue-type are noted in brackets. B) Cross-section of embryo illustrating how epiblast cells (in blue) move through the PS and form the definitive endoderm (DE) germ layer (in yellow) displacing the visceral endoderm (VE; in green). In between the DE and epiblast/ ectoderm, the mesoderm germ layer (in red) forms. Ant, anterior; AVE, anterior visceral endoderm; DE, definitive endoderm; Dist, distal; ExE, extraembryonic ectoderm; ICM, inner cell mass; Post, posterior; Prox, proximal; PS, primitive streak; VE, visceral endoderm. Modified from (Gadue et al. 2005; Wolpert 2002).

Cells start gastrulation by undergoing an epithelial-to-mesenchymal transition, enabling them to move through the PS (Figure 1-1B, cross-section of the PS). FGF3, 4 and 8 are present in the PS and FGF4 and 8 are required for gastrulating cells to leave the PS (Sun et al. 1999). From chicken studies it is suggested that FGF4 and 8 act as a chemo-attractant and a chemo-repellent, respectively on the migrating cells, driving cell movements after ingression through the PS (Wilkinson et al. 1988; Hebert et al. 1991; Niswander and Martin 1992; Sun et al. 1999; Yang et al. 2002b; Bottcher and Niehrs 2005). When cells have moved through the PS, they become either definitive endoderm (DE), displacing the visceral endoderm, or mesoderm forming an intermediate germ layer between the DE and the epiblast (Lawson et al. 1991; Tam et al. 1993; Carey et al. 1995). The remaining cells of the former epiblast become the ectoderm germ layer. Cells moving through the mid- and posterior PS early become different types of mesoderm, including some extra-embryonic mesoderm, influenced by BMP4 and low concentrations of nodal (Parameswaran and Tam 1995; Kinder et al. 1999). The mesoderm expresses Fetal-like kinase 1 (Flk1) along with Noggin and Chordin. DE is formed from the anterior regions of the PS, influenced mainly by high concentrations of nodal. Here, Forkhead box A2 (Foxa2) is expressed during gastrulation and is later found in the DE (Tam and Beddington 1992; Robb and Tam 2004). A specific marker for the DE germ layer at this stage is the Sry-related HMG box gene 17 (Sox17), which is not expressed in the other two germ layers, but is found also in the VE (KanaiAzuma et al. 2002). The VE expresses the Sry-related HMG box gene 7 (Sox7), which is not found in DE (Futaki et al. 2004; Seguin et al. 2008). DE cells migrating through the PS early during gastrulation populate the anterior parts of the endoderm, the later foregut, and cells leaving the PS late in gastrulation populate the posterior endoderm, the later hindgut (Tam and Beddington 1992). Gut tube patterning and regionalization

By the end of gastrulation around E7.5 in the mouse, the embryo contains two major signalling centres, the anterior VE (AVE) and the node. The AVE is found at the anterior side of the embryo (Figure 1-1A) and acts to restrict the PS structure and is involved in the head fold (Wells and Melton 2000). The node is found in the distal-most part of the embryo cylinder, the anterior limit of the PS, from where it directs DE patterning among other activities. The DE is patterned along the A-P axis through interactions with the anterior ectoderm, cardiac mesoderm, and notochord to form the anterior gut tube and with the node, lateral plate mesoderm and PS to form the posterior gut tube (Wells and Melton 1999; Wells and Melton 2000). At E7.5 anterior and posterior gut tube can be distinguished from one another not only by their localization in the embryo, but also by their expression of marker genes. Mouse-Cerberus 1 (mCer1), Orthodenticle homeobox 2 (Otx2), Hematopoietically expressed homeobox (Hex) and Homeobox expressed in ES cells 1 (Hesx1) are expressed anteriorly and Cdx2 and certain members of the Hox family are expressed posteriorly. Dividing the forming gut tube into four regions can illustrate how they re-localize to form the proper gut tube (Figure 1-2A&B; E8.5): Region I, the most anterior part, seems to fold over region II and form the ventral foregut, giving rise to the lungs, stomach, liver and ventral pancreas. Region II gives rise to the oesophagus, stomach, duodenum and dorsal pancreas. Similarly, region IV seems to fold over region III, giving rise to posterior trunk endoderm and hindgut, forming the large intestine. Region III gives rise to the midgut and trunk endoderm, forming the small intestine (Wells and Melton 1999). Closure of the gut tube happens as the anterior intestinal portal (AIP) and caudal intestinal portal

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(CIP) move rostrally (arrows in Figure 1-2A&B). Around E8.5 – E8.75 the embryo starts to turn so that the endoderm becomes situated on the inside, and the ectoderm on the outside of the developing organism.

Figure 1-2: Formation, folding and branching of the gut tube. A) The endoderm (yellow and light green) separated from the mesoderm (red), ectoderm (blue) and PS (pink) in an E7.5 embryo. Roman numerals I–IV represent regions of E7.5 endoderm that fate map to regions I–IV of the E8.5 gut in B). B) The forming gut tube (yellow) is seperated from the remaining embryo. The foregut tube forms as region I folds over region II and migrates in a posterior direction, whereas the hindgut tube forms when region IV folds over region III and migrates in an anterior direction (arrows in A). C) The formation of organ buds in a E10.5 embryo. D) Branching and maturation of the pancreas. A, anterior; AIP, anterior intestinal portal; CIP, caudal intestinal portal; D, dorsal; d. Panc, dorsal pancreatic bud; int, duodenum/intestine; Lu, lung; Li, liver; P, posterior; PS, primitive streak; St, stomach; V, ventral; v. Panc, ventral pancreatic bud. Modified from (Wells and Melton 1999).

Instructive signals from the adjacent germ layers specify the A-P regions of the gut tube. The anterior part of the gut tube has been shown to adopt a more posterior fate when exposed to posterior mesoderm or ectoderm in mouse embryos (Wells and Melton 2000). This has been supported by studies in chicken, where anterior endoderm changes to a more posterior fate when transplanted to a posterior site, but it cannot change to a more anterior fate when transplanted there (Kumar and Melton 2003). These studies showed that increasing concentrations of FGF4 and retinoic acid (RA) posteriorize the gut tube in mouse and chicken (Wells and Melton 2000; Dessimoz et al. 2006; Bayha et al. 2009). In the chick embryo BMP, activinA (a surrogate for nodal) and RA signalling could induce expression of the pancreatic foregut marker Pancreatic and duodenal homeobox 1 (Pdx1) in endoderm anterior to the pancreas (Kumar and Melton 2003). Thus, signals originally implicated in the A-P specification of the neural tube were shown to have a similar function in the endoderm. The mouse gut tube can be specified by regional markers, including Foxa2 expressed throughout the gut tube; Sox2 expressed in the thyroid, trachea and stomach regions; Pdx1 expressed in the duodenal and pancreatic regions; and Cdx2 expressed in the posterior gut tube (as exemplified by the chicken cartoon in Figure 1-3; chicken CdxA = mouse Cdx2).

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Figure 1-3: Regional expression of transcription factors in the endoderm. Although their expression is here mapped in the chicken embryo, their homologs are expressed in a similar regional manner in the mouse. The transcription factors shown on the left, mostly homeobox genes, are mapped to specific regions of the endoderm, as shown on the right.. These genes are not only regionally expressed in already shaped organs (as shown in the E4 chicken gut tube), but also in the endodermal sheet prior to organ formation, with stable expression domains that can be used as markers of presumptive regions. The top left pink triangle shows Hex expression in the thyroid. The bottom left triangle refers to pancreas bud and the bottom right triangle to liver bud. BA1–4, branchial arches 1–4; Chicken CdxA = mouse Cdx2. Modified from (Grapin-Botton and Melton 2000).

Pancreas and β cell formation

Pdx1 is expressed in the region of the gut tube where the pancreatic primordia start to bud around E8.75 (Figure 1-2C) and all pancreatic cell types derive from this PDX1-positive (PDX1+ hereafter) domain (Jonsson et al. 1994). The ventral and dorsal buds form from the midline and lateral areas of the PDX1+ gut, respectively, then grow and branch extensively before fusing to become one pancreas by E12.5 (Figure 1-2D; (Jorgensen et al. 2007)). The ventral bud precedes the dorsal bud and expresses the Homeobox transcription factor HB9 (Hlxb9) and Pdx1 concomitantly whereas the dorsal bud expresses these sequentially and exclude sonic hedgehog (SHH)-signalling (Hebrok et al. 1998). SHH repression allows pancreatic budding in this region and is determined by the underlying notochord. Expression of the Pancreas-specific transcription factor 1a subunit (Ptf1a; or p48) is limited to the dorsal and ventral epithelium and is coexpressed with Pdx1. The ventral pancreas develops in close association with the adjacent hepatic and bile duct endoderm and restriction of the ventral pancreas is dependent on TGFβ (SMAD2/3), BMP (SMAD1/5/8) and FGF-signalling from the cardiac mesoderm (Deutsch et al. 2001; Rossi et al. 2001). TGFβ-signalling is stable whereas FGF and BMP-signalling are dynamic and can change within a few somite stages (Wandzioch and Zaret 2009). The dorsal pancreas does not show active BMP or FGF-signalling at this stage, but rather is specified by dynamic TGFβ-

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signalling. These dynamic signalling cascades possibly determine the boundaries of pancreatic endoderm, i.e. expression of Pdx1, and the closely related liver endoderm. As the pancreatic buds grow and branch, the surrounding mesenchyme secretes FGF10, which stimulates pancreatic epithelial proliferation. This mesenchymal stimulation is absolutely necessary for maintenance of Pdx1 expression and pancreatic development, as FGF10–/– show no Pdx1 expression at E10.5 (Bhushan et al. 2001). The epithelial expression of Pdx1 and NK homeobox transcription factor 6.1 (Nkx6.1) starts to deviate into NKX6.1+ cells found only in the central part of the epithelium, whereas PDX1+/NKX6.1– cells are found at the periphery and at E13.5 mark the acini (Jorgensen et al. 2007). These acini become the exocrine part of the pancreas that produces and secretes digestive enzymes into the connecting ducts, releasing them to the intestine (Slack 1995). Neurogenin 3 (NGN3)+ endocrine precursors delaminate from the epithelium and develop into Paired box gene 6 (Pax6)-expressing endocrine cells which form the islets of Langerhans. The endocrine cells in these islets produce hormones, which they secrete to the bloodstream. Islets consist of five cell types: α cells producing glucagon; β cells producing insulin; δ cells producing somatostatin; ε cells producing ghrelin and PP cells producing pancreatic polypeptide. These islets have a distinct morphology, with the β cells in the centre and the other cell types at the periphery.

Mouse ES cells in directed differentiation The initial cell population used to generate β-like cells for cell replacement therapy may come from either somatic (or adult) stem cells or from ES cells. Somatic stem cells are mono- or multipotent cells residing in most tissues and organs of the post-natal human. They are used in the treatment of leukaemia and other haematological malignancies through bone-marrow transplantation. Additional areas under investigation include treatment of strokes, myocardial infarctions, corneal regeneration and epidermal gene therapy (Pellegrini et al. 2009; McCall et al. 2010). There is no definitive evidence of a somatic stem cell in the pancreas and thus the focus of cell replacement-therapy for diabetes is currently on ES cells (Madsen 2005). ES cells are pluripotent cells, i.e. they can give rise to all tissues of the embryo proper (Ohtsuka and Dalton 2008). They are isolated from the inner cell mass of a blastocyst stage embryo and under the right culture conditions they can multiply indefinitely, while keeping their pluripotent phenotype (Evans and Kaufman 1981; Martin 1981; Ying et al. 2003a). They hold great potential due to their pluripotent nature, but as yet, no protocol applicable to human treatment has been developed. Modelling differentiation using mouse ES cells

Mouse ES (mES) cells are used for the study of directed differentiation towards definitive endoderm, pancreatic foregut and ultimately β-like cells because they have certain advantages. Working with mES cells holds fewer ethical concerns than working with human ES (hES) cells and there are many available tools, such as transgenic mES cell lines which can be used to test hypotheses that cannot be otherwise experimentally tested. Also, mouse embryonic development closely mimics human embryonic development, suggesting that conclusions may be extrapolated and applied to the human system. But in using mES cells for scientific purposes, it is important to bear in mind that the end product will always be a cell therapy based on hES cells, and findings will therefore always have to be confirmed in this system. The sections below will focus on mES cells with examples from hES cell work where relevant. Derivation of mESCs

The first mES cells were derived almost 30 years ago and the first hES cell line was derived 12 years ago (Evans and Kaufman 1981; Martin 1981; Thomson et al. 1998). mES cells (ES cells hereafter) are used either for generation of transgenic mice or for culturing and differentiation of (transgenic) cell lines. Mouse ES cells are derived from the ICM of the E3.5 blastocyst. They are grown on feeder cells in the presence of serum and leukemia inhibitory factor (LIF), which maintains them pluripotent. 11

The pluripotent state

The pluripotency of ES cells along with their capability to self-renew indefinitely are the most important characteristics of ES cells. The pluripotent state is characterised morphologically by tightly associated cells growing in rounded clusters. Molecularly, they are characterised by the expression of markers, some of which are found also in the ICM of the developing embryo. The most commonly used are Oct4, Nanog, Stage specific embryonic antigen-1 (SSEA-1), Sox2 and Alkaline phosphatase (Solter and Knowles 1978; Pease et al. 1990; Nichols et al. 1998; Avilion et al. 2003; Chambers et al. 2003; Mitsui et al. 2003). To test if ES cells have maintained their pluripotency over time when in culture, they are evaluated in several ways, each providing a more stringent test but at the same time taking more resources. They can be evaluated by i) morphology; ii) a positive stain with antibodies for the pluripotent markers; iii) subcutaneous injection in mice and formation of teratomes with cells of all three germ layers; iv) injection into the ICM of a developing embryo where they give rise to chimaeras with contributions to tissues of all three germ layers and the germ line (Ohtsuka and Dalton 2008). The last test is considered the ‘golden standard’ but is time-consuming and is therefore not routinely carried out except in the establishment and analysis of newly generated transgenic cell lines or by investigating whether a certain (manipulated) ES cell line is entirely pluripotent. Maintaining the pluripotent state in an ES cell culture is done through prevention of differentiation (or induction of self-renewal properties) and promotion of proliferation. LIF is added to the culture medium of pluripotent ES cells and is the main factor involved in keeping cells pluripotent (Figure 1-4A). It activates the JAK/STAT3-signalling pathway, resulting in transcription of genes involved in self-renewal, one of these being C-myc (Niwa et al. 1998; Cartwright et al. 2005). LIF also activates the important Phosphoinositol 3 kinase (PI3K), leading to activation of Ras/ mitogen-activated protein kinase (MAPK) and AKT pathways. The latter is involved in relief of C-myc repression by glycogen synthase kinase-3 (GSK-3), which seems to be very important in maintaining self-renewal capability (Umehara et al. 2007). BMP is a second important factor for maintaining self-renewal, its effect possibly depending on the culture conditions in which it acts (Ohtsuka and Dalton 2008). BMP activates SMAD1/5/8-signalling leading to expression of Inhibitor of differentiation (Id)-genes which block at least neural differentiation (Figure 1-4A; (Ying et al. 2003a)). BMPs also act by suppressing the p38 MAPK, which would otherwise promote differentiation (Qi et al. 2004; Kunath et al. 2007). FGF4 is expressed in pluripotent ES cells in an autocrine fashion and the activation of the ERK1/2signalling cascade by FGF must be suppressed in order to maintain cells in the plupipotent state (Ma et al. 1992; Kunath et al. 2007; Nichols et al. 2009). The LIF-gp130 receptor complex sees to this.

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Figure 1-4: Key signalling pathways required for maintaining pluripotency and for directed differentiation into the three germ layers. A) LIF signalling activates JAK–STAT3 and PI3K pathways to induce target genes essential for pluripotency. BMP signals activate SMAD1/5/8-Id gene and suppress p38 MAPK. Activin/ nodal have been shown to contribute mESCs proliferation but not pluripotency. B) Illustration of signalling demonstrated induce to ES cell self-renewal and germ layer specification, through the EPL-state. BMP, bone morphogenetic protein; EPL, early-primitive ectoderm-like; ESCs, embryonic stem cells; FGF, fibroblast growth factor; ICM, inner cell mass; LIF, leukemia inhibitory factor. Modified from (Ohtsuka and Dalton 2008; Gadue et al. 2005).

Recent studies have suggested that epigenetic processes are required for repression of developmental pathways through the actions of e.g. polycomb-group complex proteins. How important epigenetic control and regulation of the pluripotent state is compared to the addition of growth factors and cytokines is yet to be determined (Niwa 2007). Growing ES cells as a pluripotent culture can be done on a feeder-layer of e.g. mouse embryonic fibroblasts in the presence of serum and LIF. Here, serum contains BMP and feeder cells may be substituted entirely by LIF, which they contribute to the culture condition (Smith et al. 1988; Ying and Smith 2003). Alternatively, pluripotent ES cells can be maintained in feeder-free serum replacement media containing LIF, where N2, B27 and BMP4 are added to replace serum in general, and BMP in particular (Ying et al. 2003a). Directed differentiation of ES cells

Removal of LIF (and BMP4) initiates differentiation of ES cells by increasing ERK activity. FGF5 is up-regulated and pluripotency markers are down-regulated along with PI3K and AKTsignalling pathways (Rathjen et al. 1999; Ohtsuka and Dalton 2008). Intact FGF4-signalling has been shown to be necessary for initiation of differentiation for at least ectoderm and, less convincingly, mesoderm lineages (Kunath et al. 2007). As little as a 24-hour pulse of ectopic

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FGF4 can initiate differentiation in an FGF4–/– cell line, but differentiation cannot be rescued by the addition of ectopic FGF5, which binds one of the same FGFRs as does FGF4 (Kunath et al. 2007; Stavridis et al. 2007). Based on these data, it was suggested that FGF4-signalling is necessary for leaving the pluripotent state and initiate differentiation in general. Along those lines, an Ext–/– ES cell line deficient in heparan sulphate (HS), a proteoglycan necessary for FGF-FGFR complex formation (see a later section), has recently been shown inadequate to differentiate (Kraushaar et al. 2010). Rathjen and co-workers showed that when ES cells leave the pluripotent state, they initially turn into early primitive ectoderm-like (EPL) cells (Figure 1-4B; (Rathjen et al. 1999)). This early differentiation is reversible, and EPL cells are thereby distinct from the embryonic epiblast cells, as these cannot reverse to the pluripotent state or contribute to chimaeras (Nichols and Smith 2009). The direction of differentiation taken by ES cells leaving the pluripotent state depends on the signals they receive through the growth medium (Figure 1-4B). Neural differentiation is often termed the ‘default pathway’ as it seems to be initiated solely by the removal of LIF and BMP4 (Ying et al. 2003b). However, recent work suggests that it is dependent on at least FGF and RAsignalling ((Ying et al. 2003b); Nina Engberg, unpublished data). TGFβ-signalling through nodal/ activinA (activin hereafter) and BMP4 or WNT-signalling all inhibit neural differentiation (Finley et al. 1999; Kubo et al. 2004; Vallier et al. 2004; Watanabe et al. 2005). Removing LIF, but not BMP4, induces mesoderm formation either on gelatine or on Collagen-IV coated plates, as does low concentrations of activin (Nishikawa et al. 1998; Gadue et al. 2005). DE differentiation is obtained when growing cells in high concentrations of nodal or activin, its surrogate in ES culture (Kubo et al. 2004; Tada et al. 2005; Yasunaga et al. 2005; Gadue et al. 2006). Differentiation can be done in either monolayer culture or via embryoid body (EB) formation, a 2-dimensional or 3dimensional differentiation process, respectively, with increasing secondary signalling. Endoderm differentiation and patterning

The generation of β-like cells from ES cells is hypothesized to be a stepwise differentiation and maturation process (Figure 1-5A; (Madsen and Serup 2006)). These steps recapitulate developmental stages of the embryo in vitro as such mimicry is believed to be the best way to obtain functional β-like cells capable of functionally replacing the patient’s β cells. Although culture conditions for maintaining mES and hES cells in the pluripotent state differ significantly, directed differentiation towards β-like cells has proven somewhat similar and data from the human system are therefore included in this section.

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Figure 1-5: Stepwise differentiation path from ES cells to β-like cells exemplified in the human system. A) hES cells undergo a stepwise differentiation through definitive endoderm, primitive gut tube, posterior foregut and pancreatic endoderm to become endocrine cells as β-like cells within 15-18 days in culture. B) A protocol for this stepwise differentiation with culture conditions depicted in the top panel, number of days in each step, intermediate cell type and finally marker expression in the bottom panel. ES, embryonic stem; ME, mesendoderm; DE, definitive endoderm; PG, primitive gut tube; PF, posterior foregut endoderm; PE, pancreatic endoderm; EN, endocrine precursor. Modified from (Madsen and Serup 2006; D’Amour et al. 2006).

High concentrations of activin (up to 100 ng/ml) are routinely used to differentiate both mES and hES cells to DE (Kubo et al. 2004; D'Amour et al. 2005; Gadue et al. 2006). This differentiation has an efficiency of app. 40 – 80% by day 5 without sorting of the cell culture and is dependent on WNT and FGF-signalling (Funa et al. 2008; Morrison et al. 2008; Willems and Leyns 2008). The DE population has been shown in hES cell cultures to be expandable, a feature much desired for putative future usage in cell replacement therapies where expandable intermediary stages may prove very cost-efficient (Morrison et al. 2008; Seguin et al. 2008). When differentiating cells to endoderm, it is crucial to ensure that it becomes definitive and not visceral endoderm, as the gut tube and its derivate organs stem from the DE germ layer. DE and VE share common markers such as Sox17, E-cadherin and Foxa2 but differ in other markers, DE expressing Cxcr4 and VE expressing Sox7 and Thermostable direct hemolysin (Tdh) among others (Yasunaga et al. 2005; Sherwood et al. 2007). A detailed analysis of marker expression is therefore necessary to ensure that the right type of endoderm is obtained. This has proven not so much a practical issue as a theoretical one, as differentiation in high concentrations of activin leads to formation of DE and not VE. Having established DE cells, a stepwise protocol based on embryonic development studies in mouse and chicken are employed (Madsen and Serup 2006; Van Hoof et al. 2009). Exemplified

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in hES cells by one of the most successful protocols reported to date, these steps go from DE via primitive gut tube, posterior foregut endoderm, pancreatic endoderm and endocrine precursors to hormone-expressing endocrine cells (Figure 1-5B; (D'Amour et al. 2006)). Cells at each of these steps express a specific set of genes which can be used as markers to analyse the efficiency and quality of the differentiation process (see Figure 1-5B for specific markers). This protocol was based on the fact that induction of gut tube markers and posterior foregut PDX1+ cells includes suppression of SHH (by KAAD-cyclopamine) and presence of FGF10. Other groups have found that RA and FGF4 could induce app. 30% PDX1+ cells from hES cell culture and in a study using FGF2, intermediary concentrations of thereof determine Pdx1-induction whereas higher concentrations determine posterior small intestinal fates and inhibits hepatocyte differentiation (Johannesson et al. 2009; Ameri et al. 2010). D’Amour and co-workers generated insulinproducing cells and in a later protocol they showed how pancreatic epithelial cells could become fully glucose-responsive β-like cells when transplanted into the epididymal fat pad and left to mature in vivo (Kroon et al. 2008). Thus as a cell replacement therapy it may not be necessary to fully differentiate the transplanted material, as long as these are differentiated to a degree where the in vivo conditions can mature them, provided that they do not show unwanted side effects such as teratoma-formation which has been observed in a few transplants (Kroon et al. 2008). In the mouse ES cell field there has been little success in obtaining PDX1+ cells in monolayer culture. Although the more diffuse 3-dimensional culture or EB formation as the starting point holds some potential (Schroeder et al. 2006; Wang and Ye 2009), a reproducible, growth factordriven protocol in not available for now. Accordingly, the hES cell field seems ahead of the mES cell field at the present time. However, several laboratories have tried to copy the ‘D’Amour protocol’ and have found it difficult, partly due to the success of the protocol being highly dependent on the cell line used and due to high induction of hepatic cell types (Mfopou et al. 2007; Cho et al. 2008; Ricordi and Edlund 2008; Semb 2008).

FGFs, FGFRs and signalling in mES cells Fibroblast growth factors (FGFs) were initially discovered as having a mitogenic effect on fibroblast cells (Armelin 1973; Gospodarowicz 1974) and are involved in multiple functions in the organism. During development they play key roles in germ layer formation and ES cell differentiation (as explained above), in the formation of most neural processes, in skeletal formation and are crucial for limb formation and development (Ornitz and Itoh 2001; Bottcher and Niehrs 2005; Mason 2007). In the adult organism, they are involved in cell migration, differentiation, proliferation, chemotaxis, survival and apoptosis (Bottcher and Niehrs 2005). FGFs and FGFRs

FGFs are a family of growth factors comprising 22 members in mouse and human, named FGF123 (FGF15 is the mouse ortholog of the human FGF19). They are between 17 – 34 kDa in size and share a 16-65% identity in a conserved 120 amino acid sequence (Bottcher and Niehrs 2005; Eswarakumar et al. 2005). It is this conserved portion of the protein which interacts with the FGF receptors (FGFRs) and heparan sulfate (HS; see the section below). They are arranged into several families based on their sequence homology (Figure 1-6A; (Itoh and Ornitz 2008)). Overall there are three types of FGFs: i) FGF11 – 14 are intracellular FGFs (iFGFs), which do not act through binding to the FGFRs, but rather seem to interact with downstream signalling pathways intracellularly (Goldfarb 2005); ii) FGF15, 21 and 23 are hormone-like FGFs (hFGFs), which have low-affinity HS-binding sites and act in an endocrine manner (Tomlinson et al. 2002; Kharitonenkov et al. 2005; Fukumoto and Yamashita 2007; Itoh and Ornitz 2008); iii) the remaining (canonical) FGFs are secreted proteins with high HS-binding affinities meaning they are restricted to the vicinity of FGF-producing cells and act locally (Itoh and Ornitz 2004; Itoh and Ornitz 2008). FGF knockout mice show very different phenotypes from embryonic lethality (FGF4, 8 and 15), postnatal lethality (FGF9, 10, 18) and viable mutants with various phenotypes (FGF2, 3, 5, 6, 7, 12, 14, 17) to no phenotype at all (FGF1; (Ornitz and Itoh 2001)). 16

There are four membrane-bound FGFRs consisting of three Ig domains (domains I – III) on the extracellular side and of two tyrosine kinase domains on the intracellular side of the cell surface. The Ig-domains convey FGF-binding, determine FGF ligand selectivity (domain III) and interact with HS (Bottcher and Niehrs 2005). The C-terminal half of Ig-domain III in FGFR1-3 demonstrates alternative splicing of exon 7 to either exons 8 or 9, generating FGFR(III)b or FGFR(III)c isoforms, respectively (FGFRb or FGFRc hereafter; Figure 1-6C; (Johnson and Williams 1993; Groth and Lardelli 2002; Eswarakumar et al. 2005)). The expression of specific FGFR-isoforms is tissue-specific, meaning ligand-receptor interactions can be regulated across tissues, giving a high range of interaction-potential to modulate downstream signalling pathways and gene expression. The intracellular kinase domains are responsible for tyrosine kinase activity leading to auto-phosphorylation and recruitment of downstream signalling components eliciting the cellular response to ligand binding. FGFR4, which has no splice variants, resembles FGFRcisoforms both structurally and in FGF binding-affinities (Vainikka et al. 1992). A secreted third isoform, FGFR1-3(III)a lacks the trans-membrane and intracellular kinase domains, and although they are present in blood, their function is somewhat unclear (Johnson et al. 1990; Johnson et al. 1991; Johnson and Williams 1993; Hanneken 2001).

Figure 1-6: The evolution of the FGF family, binding of FGF to FGFR and alternative splicing of the FGFR. A) The evolutionary relationships within the fibroblast growth factor (FGF) gene family. The twenty-two FGF encoding genes are arranged into seven subfamilies. Branch lengths are proportional to the evolutionary distance between each gene. FGF19 is the human ortholog of mouse FGF15. The FGF11 subfamily is also referred to as the intracellular FGFs (iFGFs), and the FGF15 (FGF19) family as the hormone-like FGFs (hFGFs). B) Ribbon diagram of the ternary FGF2/heparin/FGFR1 complex showing FGF2 in yellow, D2 and D3 domains of the ligand-binding portion of FGFR1 in green and blue, respectively, and heparin in red. C) The two isoforms of FGFR1-3 are generated by

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alternative splicing of exons 8 and 9. The C-terminal half of the DIII domain is encoded by exon 8 to generate the FGFR(III)b isoforms while the C-terminal half of DIII is encoded by exon 9 to generate the FGFR(III)c isoforms. Modified from (Itoh and Ornitz 2004; Eswarakumar et al. 2005).

HS, downstream signalling pathways and regulation

Heparan sulfates (HSs) are proteoglycans consisting of repeated subunits of D-glucosamine and disaccharides and are embedded in the extracellular matrix on the cell surface. Mutations in genes involved in the synthesis or modulation of HS results in developmental defects in mice, most likely due to impaired FGF-signalling (Bullock et al. 1998; Ornitz 2000; Kraushaar et al. 2010)). HS stabilizes FGFs to thermal denaturation, proteolysis and limit their diffusion and release to interstitial spaces (Moscatelli 1987; Flaumenhaft et al. 1990). This leads to increased 1:1 FGF:FGFR complex formation which then results in a transient receptor dimerization and signalling from this 2:2 FGF:FGFR complex (Figure 1-6B; (Hsu et al. 1999; Plotnikov et al. 1999; Pye and Gallagher 1999)). Intracellular phosphorylation of the kinase-domains leads to activation of intracellular signal transduction pathways. Most commonly, the Ras/MAPK pathway is activated upon FGFsignalling, but also the PLCγ/Ca2+ and PI3K/AKT pathways are activated (Bottcher and Niehrs 2005). For activation of the Ras/MAPK pathway, FRS2 is recruited and forms a complex activating Ras. Pathway activity ultimately leads to nuclear translocation of MAPK and activation of target genes, such as AP1, c-myc and ETS transcription factors (Wasylyk et al. 1998). Activation of PLCγ leads to intracellular release of Ca2+ and activation of phosphokinase C (PKC; (Pawson 1995)). Finally, activation of PI3K through either direct interaction with the FGFR or by components of the Ras/ MAPK pathway leads to activation of AKT (Carballada et al. 2001). FGF-signalling is regulated by members of the sprouty and sprouty-related EVH1 protein (SPRED) families by a feedback mechanism that regulates MAPK-signalling through receptor tyrosine kinase binding (Hacohen et al. 1998; Lim et al. 2002). Furthermore, FGF-activity is modified through a tight regulation of HS-synthesis and certain transmembrane regulators (SEF and FLRT), which interrupt FGF-FGFR complex formation or downstream signalling (Bottcher and Niehrs 2005). Synthetic small molecules such as SU5402 and PD173074 inhibit most FGFRsignalling with some or little secondary effects, respectively (Mohammadi et al. 1997; Mohammadi et al. 1998). To date, the role of FGF-FGFR signalling in directed differentiation towards cell types of the endoderm germ layer has been investigated as a general activation or inhibition of FGF-FGFR signals. The role of individual FGF- and FGFR family members in differentiation towards DE remains to be elucidated.

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2. Aims The aim of this study was to investigate the role of FGF-signalling in directed differentiation of mouse ES cells towards i) mesendoderm formation, and ii) definitive endoderm formation and patterning. We applied a mono-layer, serum-free culture system in which differentiation of mouse ES cells towards mesendoderm was achieved by addition of BMP4 and a range of activin-concentrations to the culture medium. Formation of definitive endoderm was reached using high concentrations of activin and further patterning hereof was done using various growth factors and inhibitors. We investigated the influence of FGFs on this system by addition of a range of FGFs, small molecule FGF-inhibitors and soluble FGFRs. We analysed the resulting cell populations by using green fluorescent protein (GFP)-linked reporter cell lines and knockout cell lines, immunecytochemistry, RT-PCR and qPCR.

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3. Paper I A late requirement for Wnt and FGF signaling during activin-induced formation of foregut endoderm from mouse embryonic stem cells Published in Developmental Biology, 2009, 330, p. 286 – 304. Mattias Hanssona,1, Dorthe R- Olesena,b,1, Janny M.L. Peterslunda, Nina Engberga, Morten Kahna,b, Maria Winzia, Tino Kleina, Poul Maddox-Hyttelb and Palle Serupa,* a

Department of Developmental Biology, Hagedorn Research Institute, Niels Steensens Vej 6, DK-2820 Gentofte, Denmark. b Department of Animal and Veterinary Basic Sciences, Faculty of Life Sciences, University of Copenhagen, DK-1870 Frederiksberg C, Denmark. 1 These authors contributed equally to this work. * Corresponding author. Author contributions

The majority of the experiments were performed in collaboration between Mattias Hansson and Dorthe R. Olesen with equal contributions. I worked on the effect of FGF-signalling upon foregut endoderm formation, and in collaboration with Nina Engberg looked at the further patterning of activin-induced definitive endoderm towards Pdx1-expressing pancreatic endoderm. The paper was published in Developmental Biology, issue 330, 2009. Stated below is each individual’s contribution to the paper (numbers indicate figures). Mattias Hansson Dorthe R. Olesen Janny M. L. Peterslund Nina Engberg Morten Kahn Maria Winzi Tino Klein Poul Maddox-Hyttel Palle Serup

1; 2; 3; 4A; 7; 10; 11; S1; S2; S3B,C; S4; Wrote paper draft 1; 2; 3; 4A,C,F; 5; 8B; S1; S2 8A; 9; 10; 12; S4 4B,D,E; 12 S3A; S5B,C 6; S6 S5A Supervisor Principal investigator and supervisor; paper revision

For co-authorship declaration, see Appendix A.

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Developmental Biology 330 (2009) 286–304

Contents lists available at ScienceDirect

Developmental Biology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / d e v e l o p m e n t a l b i o l o g y

A late requirement for Wnt and FGF signaling during activin-induced formation of foregut endoderm from mouse embryonic stem cells Mattias Hansson a,1, Dorthe R. Olesen a,b,1, Janny M.L. Peterslund a, Nina Engberg a, Morten Kahn a,b, Maria Winzi a, Tino Klein a, Poul Maddox-Hyttel b, Palle Serup a,⁎ a b

Department of Developmental Biology, Hagedorn Research Institute, Niels Steensens Vej 6, DK-2820 Gentofte, Denmark Department of Animal and Veterinary Basic Sciences, Faculty of Life Sciences, University of Copenhagen, DK-1870, Frederiksberg C, Denmark

a r t i c l e

i n f o

Article history: Received for publication 16 July 2008 Revised 18 March 2009 Accepted 30 March 2009 Available online 7 April 2009 Keywords: Embryonic stem cell Gastrulation Endoderm Mesendoderm Anterior–posterior patterning TGF-β Wnt FGF

a b s t r a c t Here we examine how BMP, Wnt, and FGF signaling modulate activin-induced mesendodermal differentiation of mouse ES cells grown under defined conditions in adherent monoculture. We monitor ES cells containing reporter genes for markers of primitive streak (PS) and its progeny and extend previous findings on the ability of increasing concentrations of activin to progressively induce more ES cell progeny to anterior PS and endodermal fates. We find that the number of Sox17- and Gsc-expressing cells increases with increasing activin concentration while the highest number of T-expressing cells is found at the lowest activin concentration. The expression of Gsc and other anterior markers induced by activin is prevented by treatment with BMP4, which induces T expression and subsequent mesodermal development. We show that canonical Wnt signaling is required only during late stages of activin-induced development of Sox17-expressing endodermal cells. Furthermore, Dkk1 treatment is less effective in reducing development of Sox17+ endodermal cells in adherent culture than in aggregate culture and appears to inhibit nodal-mediated induction of Sox17+ cells more effectively than activin-mediated induction. Notably, activin induction of GscGFP+ cells appears refractory to inhibition of canonical Wnt signaling but shows a dependence on early as well as late FGF signaling. Additionally, we find a late dependence on FGF signaling during induction of Sox17+ cells by activin while BMP4-induced T expression requires FGF signaling in adherent but not aggregate culture. Lastly, we demonstrate that activin-induced definitive endoderm derived from mouse ES cells can incorporate into the developing foregut endoderm in vivo and adopt a mostly anterior foregut character after further culture in vitro. © 2009 Elsevier Inc. All rights reserved.

Introduction Directed differentiation of embryonic stem (ES) cells into mesoand endodermal derivatives is intensely studied due to their potential clinical applications. Meso- and endoderm is formed by epiblast cells that ingress through the primitive streak (PS) during gastrulation (reviewed in Tam and Loebel, 2007). Fate mapping studies have shown that cells that migrate through different anterior–posterior regions of the streak give rise to different mesodermal and endodermal components (Carey et al., 1995; Lawson, 1999; Lawson and Pedersen, 1992). At early stages, mesodermally fated cells ingress alongside endodermally fated cells but it is unclear when and how ingressing cells acquire their ultimate fate. The definitive endoderm (DE) is derived from progenitors migrating through the anterior PS at early and mid-streak stages (Carey et al., 1995; Lawson, 1999; Lawson

⁎ Corresponding author. Fax: +45 44438000. E-mail address: [email protected] (P. Serup). 1 These authors have contributed equally to this work. 0012-1606/$ – see front matter © 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.ydbio.2009.03.026

and Pedersen, 1987; Lawson and Pedersen, 1992). Moreover, recent evidence suggests that a common progenitor population, the mesendoderm, exists in the PS (Kinder et al., 2001; Lawson et al., 1991; Tada et al., 2005) and that the cumulative exposure to nodal signaling determines mesendodermal fates such that increasing exposure to nodal shifts the fate from posterior mesoderm through anterior mesoderm and posterior endoderm to anterior DE at the largest dose (Ben-Haim et al., 2006). The use of mouse ES (mES) cell lines with the green fluorescent protein (GFP) targeted to the PS- and early mesodermal-specific genes Brachyury (T), Mix1 homeobox-like 1 (Mixl1), and Goosecoid (Gsc) has made it possible to quantify mesendoderm induction and isolate and characterize different mesodermal and endodermal populations (Fehling et al., 2003; Gadue et al., 2006; Kubo et al., 2004; Ng et al., 2005; Tada et al., 2005; Yasunaga et al., 2005). Anterior PS fates and endoderm was induced with high concentrations of activin A (activin hereafter) that activates Smad2/3 signaling through binding to the same receptor as nodal. Recent studies have extended the endoderminducing properties of activin to human ES cell differentiation cultures (D'Amour et al., 2005, 2006). However, as nodal signaling in the early

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embryo interacts with other signaling pathways such as the BMP, Wnt, and FGF pathways (reviewed in Tam and Loebel, 2007), we address here the role of these signals and their potential to modulate activininduced mesendodermal differentiation of mES cells grown under standard conditions in feeder- and serum-free adherent monoculture (Ying et al., 2003a,b). We use ES cells containing reporter genes (lacZ or GFP) targeted to the T (Fehling et al., 2003), Mixl1 (Hart et al., 2002), Gsc (Tada et al., 2005), Flk1 (Shalaby et al., 1995), Sox17 (Kim et al., 2007), and Sox2 (Li et al., 1998) loci to monitor, over time, the effects of different growth factors on the expression of markers specific to different anterior and posterior regions of the PS and derivatives thereof. We confirm and extend previous findings on the ability of increasing concentrations of activin to progressively induce more ES cell progeny to an anterior PS fate. Remarkably, while the number of Gsc- and Sox17-expressing cells increases with increasing activin concentration, the highest number of T-expressing cells is found at the lowest activin concentration, similar to the activin response seen in Xenopus animal cap cells. Furthermore, expression of Gsc and other anterior markers induced at high activin doses is prevented by simultaneous treatment with BMP4 which redirects development towards mesodermal fates, also similar to results from Xenopus. Extending previous work, we find that inhibition of canonical Wnt signaling by treatment with Dkk1 is able to prevent activin-induced development of endodermal cells but only at late stages of differentiation. Dkk1 also inhibits activin-induced Mixl1expression and consistent with this finding Wnt3a and activin act additively on Mixl1 expression but not on Gsc expression. Wnt3a by itself appears to induce only posterior PS fates depending, however, on endogenous Smad2/3 signaling. Additionally, we demonstrate that induction of anterior and posterior PS fates by activin or BMP4, respectively, is dependent on FGF signaling. Lastly, we demonstrate for the first time that activin-induced DE derived from mES cells can incorporate into the developing foregut endoderm when implanted into chicken embryos but respond only to a limited degree to posteriorizing cues in vitro by initiating expression of regional foregut markers. Materials and methods Cell culture and differentiation of ESCs Mouse ES cells (40,000 cells/cm2) were kept undifferentiated on gelatin-coated cell culture plastic (Nunc) in serum-free medium; KODMEM supplemented with N2, B27, 0.1 mM nonessential amino acids, 2 mM L-glutamine, Penicillin/Streptomycin (all from Invitrogen), 0.1 mM 2-mercaptoethanol (Sigma-Aldrich), 1500 U/ml leukemia inhibitory factor (LIF, Chemicon) and 10 ng/ml BMP4 (R&D Systems), essentially as described by Ying et al. (2003a). ES cells were passaged every second day with daily media changes for at least three passages (6 days) prior to initiation of differentiation studies. For differentiation experiments cells grown as described above were dissociated to single cells and differentiation was induced by seeding 2000 cells/cm2 on gelatin-coated cell culture plastic in KODMEM supplemented with N2, B27, 0.1 mM nonessential amino acids, 2 mM L-glutamine, Penicillin/Streptomycin (all from Invitrogen), 0.1 mM 2-mercaptoethanol (Sigma-Aldrich) without LIF and BMP4. The medium was supplemented with one or more of the following growth factors, soluble receptors, and small molecule compounds: activin (3, 10, 30 or 100 ng/ml), Wnt3a (5 or 100 ng/ml), Nodal (1 µg/ml), BMP4 (10 ng/ml), Dkk1 (320 ng/ml; all from R&D Systems), and FGF2 (100 ng/ml; Invitrogen). Soluble FGF receptors (all from R&D Systems) were first used to achieve inhibition of ligands specific for both b and c splice forms by mixing sFGFR1IIIc, sFGFR2IIIb, and l sFGFR4 (12, 8 and 24 ng/ml, respectively). To achieve selective inhibition of the b or c splice form specific FGFs we mixed sFGFR1IIIb and sFGFR2IIIb (both at 250 ng/ml) or sFGFR1IIIc and sFGFR4 (both at

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250 ng/ml), respectively. The medium containing FGF2 or sFGFRs was supplemented with 10 μg/ml heparan sulfate (Sigma-Aldrich), 1 μM SB431542 (Inman et al., 2002), 10 μM SU5402 or 100 nM PD173074 (Calbiochem). The cells were cultured for up to 7 days and the medium was changed daily, beginning at the second day of differentiation. It should be noted that our B27 supplement contain retinyl acetate which is a precursor during RA synthesis. However, experiments using B27 supplement without retinyl acetate (Invitrogen) did not affect the number of activin-induced Sox17-GFPHi endodermal cells. Further differentiation of day 5 activin-induced cultures was done by 3 days of additional culture in serum-free medium (KO-DMEM, N2, B27, 0.1 mM nonessential amino acids, 2 mM L-glutamine, Penicillin/Streptomycin, 0.1 mM 2-mercaptoethanol) supplemented with Wnt3a (5 ng/ml), FGF4 (10 ng/ml) and/or 0.1 μM all-trans retinoic acid (Sigma). Differentiation in embryoid bodies (EBs) was carried out using the hanging drop method. Cells were dissociated to single cells using nonenzymatic Cell Dissociation Solution (Sigma-Aldrich) and diluted in N2B27 medium containing the relevant growth factors to yield 100 cells per 20 μl drop. Approximately 150 drops were applied to the lid of a 14 cm cell culture dish (Nunc), and placed upside down over autoclaved Millipore water. Drops were left overnight and EBs were washed down with HBSS w/o Ca2+ and Mg2+ and left to sediment for 3–4 min before removing the supernatant and transferring EBs to 50 mm Petri dishes (Sterilin) containing N2B27 medium with the relevant growth factors. The medium was changed daily. We frequently observed that 3–5 individual aggregates would form in the hanging drop yielding aggregates composed of 20–30 cells. Flow cytometry The cells were dissociated in 0.05% Trypsin-EDTA (Invitrogen) and a percentage of GFP+ cells was analyzed on a FACSCalibur flow cytometer (BD Biosciences) at days 2–6 in at least three independent experiments. Mean % GFP+ cells ± standard deviation (S.D.) was calculated and statistical analyses were performed using a two-tailed Student's t-test for paired samples, unless we had a clear expectation of the outcome in which case a one-tailed test was used. Sorting of GFP+ cells for RNA extraction was performed on a FACSAria (BD Biosciences). CXCR4 expression was analyzed on a FACSCalibur flow cytometer using a human anti-CXCR4 monoclonal antibody (MAB172; R&D Systems). Cells were dissociated using Collagenase (SigmaAldrich) for 5 min. The cell suspension was fixed and stained as described below without permeabilization. Visual inspection of the stained cells by confocal microscopy confirmed surface localization of the antigen. Immunofluorescence and X-gal staining The cells were cultured on gelatin-coated chamber slides for 3, 5 or 8 days and fixed at room temperature for 30 min in 4% formaldehyde solution (Mallinckrodt Baker) for immunofluorescence or 5 min in 0.2% glutaraldehyde for X-gal staining. For immunofluorescence, the cells were permeabilized in graded ethanol followed by blocking in 10% donkey serum for 1 h and incubation with primary antibody for 1 h at room temperature or overnight at 4 °C. The following antibodies were used: goat anti-Foxa2 (Santa Cruz Biotechnology), goat anti-T (R&D Systems), rat anti-E-cadherin (E-cad; Zymed/Invitrogen), goat anti-Sox17 (R&D Systems), Alexa 448 conjugated rabbit anti-GFP (Molecular Probes/Invitrogen), rabbit anti-β-galactosidase (MP Biomedicals), rabbit anti-Lhx1 (Chemicon), goat and rabbit antiPdx1 (a kind gift from C. Wright), mouse and rabbit anti-Nkx6-1 (Jensen et al., 1996; Pedersen et al., 2006), rabbit anti-Sox2 (Chemicon) and mouse anti-Cdx2 (BioGenex). The cells were incubated with Cy2-, Cy3-, Texas Red- or Cy5-conjugated species-

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specific secondary antibodies (Jackson ImmunoResearch Laboratories) and 4′,6-diamidino-2-phenylindole (DAPI, MP Biomedicals). The Lhx1 staining used tyramide signal amplification (PerkinElmer) according to the manufacturer's recommendations. Negative controls, where the primary antibodies were omitted, were included for all stainings. These controls showed no unspecific staining of the secondary antibodies (data not shown). β-galactosidase activity was visualized by adding X-gal stain solution for 4 h at 37 °C. The slides were analyzed using an LSM 510 META laser scanning microscope (Carl Zeiss) or a BX60 epifluorescence microscope equipped with a DP71 camera (both from Olympus). RT-PCR and qPCR Cells were dissociated using 0.05% Trypsin-EDTA and collected by centrifugation. Total RNA was isolated using the RNeasy kit with DNAse treatment (Qiagen) following the manufacturer's protocol. cDNA was prepared from 5 or 100 ng RNA using MMLV Reverse Transcriptase (Invitrogen). PCR reactions were performed using 1 μl cDNA, 1 μl 20 μM primer mix and 23 μl Reddy Mix PCR master mix (Abgene). The PCR was carried out with an initial denaturation step at 96 °C for 2 min, followed by 32–38 cycles of 96 °C for 30 s, 55 °C for 30 s and 72 °C for 1 min. The PCR was finished with a final extension step at 72 °C for 5 min. QPCR was performed using the standard SYBR® Green program with dissociation curve of the Mx3005P (Stratagene). PCR reaction was run in duplicates using 5 μl Brilliant® SYBR® Green QPCR Master Mix (Stratagene), 1 μl cDNA, 1 μl 10 μM primer mix and 3 μl DEPC-treated water. Quantified values for each gene of interest were normalized against the input determined by the housekeeping genes G6pdh and Tbp. The results are expressed as the relative expression level compared with the vehicle control condition or the scrambled control siRNA in the vehicle condition. Primer sequences are available on request. siRNA transfection The sequence effective for mouse β-catenin knock-down was designed using software available at the web site of Invitrogen (http://rnaidesigner.invitrogen.com/rnaiexpress/). The β-catenin (accession number NM_0079614.2) target sequences of the STEALTH siRNAs were 5′-GCCTTCATTATGGACTGCCTGTTGT-3′ (siRNA1) and 5′GAGCAAGGCTTTTCCCAGTCCTTCA-3′ (siRNA2). The STEALTH negative control siRNA (scrambled; Invitrogen) has been used as negative control and is labeled “scrambled”. The cells were cultured on gelatin-treated 24-well plates as previously described with a starting density of 4000 cells/cm2. After 1 day of differentiation the cells were transfected with 100 nM STEALTH siRNAs using Lipofectamine2000 according to the manufacturer's instructions. The transfected cells were grown further and analyzed for β-catenin expression by western blot at days 3 and 5 of differentiation. QPCR and flow cytometry analysis was performed at day 5. Western blot Total cell lysates were obtained on days 3 and 5 of differentiation culture using RIPA lysis buffer. 20 μg of each protein sample were loaded and analyzed by western blot using a rabbit anti-β-catenin monoclonal antibody (Lab Vision) and a mouse anti-β-Actin monoclonal antibody (Sigma-Aldrich) as primary antibodies as well as secondary HRP-conjugated antibodies (Santa Cruz Biotechnology).

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Generation of stable Wnt-reporter ES cell lines and chimera formation The SuTOP-CFP construct was generated by cutting the luciferase gene from the Super8XTOPFLASH Wnt reporter (generously provided by Dr. R. Moon, University of Seattle, WA) with Fse and Nco1 and replacing it with Cerulean PCR product from the pmCerulean-C1 vector (Rizzo et al., 2004). To generate stably transfected cell lines, E14 cells (Hooper et al., 1987) of low passage number were co-transfected with SuTOP-CFP and a ploxP-Neo vector, conferring resistance to Neomycin. The relationship between plasmids was 10:1 (reporter: ploxP-Neo). Transfection was carried out using Lipofectamine2000 (Invitrogen) and 24 h after transfection, selection was added (200 μg/ ml G418, Invitrogen). Medium was changed daily for 9 days and 0.5 μM BIO (GSK3β inhibitor, Calbiochem), which activates canonical Wnt signaling, was added to the medium for the last 2 days of selection to identify Wnt-responsive colonies. Fluorescent colonies were then picked and expanded and one clone was selected for chimera formation via blastocyst injection. E3.5 blastocysts were harvested by flushing the uteri of mature, time mated NMRI mice (Taconic). Chimeric embryos were generated by injection of approximately 10 SuTOP-CFP ES cells into the blastocoel using a paraffin-oil driven manual injector (Cell Tram Vario, Eppendorf) and a Narishige micromanipulator. Following 3 h of culture in M16 medium, embryos were transferred to the uterus of E2.5 pseudo-pregnant, 7 week old NMRI foster mothers. Care of the animals was done according to institutional guidelines. Embryos were harvested at E10.5 and fixed in Lilly's fixative (4% phosphate buffered formaldehyde) for 30 min before being analyzed for native Cerulean fluorescence. All animal experiments were performed in accordance with institutional and national regulations. Chick embryo grafting Fertilized eggs from white leghorn chicken were purchased from Triova and incubated at 38 °C to Hamburger and Hamilton (HH) stages 8–10 (Hamburger and Hamilton, 1951). The embryos were explanted as previously described (Chapman et al., 2001). E14 ES cell progeny was prepared for grafting by labeling with fluorescent CMTMR CellTracker dye (Molecular Probes/Invitrogen). Clumps of cells were scraped off, washed in PBS and inserted between endoderm and mesoderm of chicken embryos via a small incision in the endoderm. Grafted embryos were incubated for 48 h in a humidified incubator at 38 °C. The embryos were isolated, washed in PBS, fixed in 4% PFA at room temperature for 2 h and stored in methanol at − 20 °C until the time of analysis. Whole-mount immunofluorescent analyses of grafted chicken embryos were performed as previously described (Ahnfelt-Ronne et al., 2007). Results Dose-dependent effects of activin on the expression of PS genes are modulated by BMP and Wnt signaling Previous studies have demonstrated induction of Brachyury (T) and Goosecoid (Gsc) by activin in mES cells (Gadue et al., 2006; Kubo et al., 2004; Tada et al., 2005; Yasunaga et al., 2005). However, differences in media compositions and the culture methods used make a direct comparison of the response of these two genes to varying doses of activin difficult. Prior to the induction of differentiation by addition of growth factors, we culture the undifferentiated ES cells under defined conditions (Ying et al., 2003a). As activin is known to dosedependently regulate T and Gsc expression in Xenopus animal cap

Fig. 1. Activin, BMP and Wnt signaling control the dynamic expression of primitive streak genes in differentiating ES cells. Flow cytometric analysis of TGfp/+ (A), GscGfp/+ (B), or Mixl1Gfp/+ (C) cells grown in adherent culture for up to 6 days in serum-free media containing 0, 3, 10, 30 or 100 ng/ml activin in the presence or absence of 10 ng/ml BMP4 or 100 ng/ml Wnt3a. The mean % GFP+ cells ± standard deviation of three independent experiments is presented.

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Fig. 2. BMP4 but not Wnt3a inhibits the expression of Foxa2 and E-cadherin, and promotes expression of Flk1 in the presence of activin. The expression of Foxa2, E-cad (Cdh1), and Flk1 (β-gal) was analyzed by immunofluorescence in Flk1LacZ/+ ES cells cultured for 5 days in media containing 0, 3 or 100 ng/ml activin, 100 ng/ml Wnt3a, 10 ng/ml BMP4, 100 ng/ml activin + 100 ng/ml Wnt3a, or 100 ng/ml activin + 10 ng/ml BMP4.

cells such that low doses will activate T and high doses will activate Gsc (Dyson and Gurdon, 1998; Green et al., 1992; Gurdon et al., 1994, 1999; Latinkic et al., 1997), we initially tested if exposure of mES cells to increasing doses of activin would lead to a shift from T to Gsc expression. ES cell lines carrying T-Gfp (TGfp/+) and Gsc-Gfp (GscGfp/+) knock-in alleles (Fehling et al., 2003; Tada et al., 2005) were cultured with increasing amounts of activin (from 3 to 100 ng/ml) and the number of GFP+ cells was analyzed by flow cytometry. Examples of primary flow cytometry diagrams are shown in Fig. S1. When GFP+ cells were quantitated we found that 3 ng/ml activin transiently induced 21 ± 15% T-GFP+ cells (mean % ± S.D., n = 3) peaking at day 4 (Fig. 1A). However, this induction was not statistically significant. At higher activin concentrations the number of T-GFP+ cells declined gradually such that the highest dose (100 ng/ml) resulted in only

5 ± 4% T-GFP+ cells at day 4, comparable to the control samples cultured in the absence of activin (6 ± 5% T-GFP+ cells, Fig. 1A). In contrast, flow cytometric analyses of GscGfp/+ cells showed that the expression of this anterior PS marker (Blum et al., 1992) was induced by activin in a dose-dependent manner with expression peaking at days 5–6 (Fig. 1B). 3 ng/ml activin induced 25 ± 4% Gsc-GFP+ cells at day 5, and this number increased with increasing concentration of activin reaching 43 ± 11% Gsc-GFP+ cells in cultures treated with 100 ng/ml activin (Fig. 1B). Control samples grown in the absence of activin contained 3 ± 4% Gsc-GFP+ cells at this time point. The induction of Gsc expression at day 5 was statistically significant for all activin concentrations tested (p b 0.05). Thus, similar to the case in Xenopus animal cap cells, low doses of activin support T expression while high doses stimulate Gsc expression. Intriguingly, while

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expression of the PS marker Mixl1 (Pearce and Evans, 1999) was induced by activin, the number of Mixl1-GFP+ cells was independent of the activin concentration (Fig. 1C). In vivo, BMP4 is a ventralizing agent, acting during gastrulation to induce T and repress Gsc expression (Fainsod et al., 1994; Jones et al., 1996; Steinbeisser et al., 1995). We found a strong but transient induction of T expression in response to BMP4 (Fig. 1A). Peaking at day 3, we observed 48 ± 15% T-GFP+ cells, which was significantly higher than detected in vehicle-treated cells (p b 0.05). The induction of T expression by BMP4 was observed regardless of the presence or absence of activin. However, the activin-mediated induction of GscGFP+ cells at day 5 was strongly inhibited by BMP4 (p b 0.05), irrespective of the activin concentration (Fig. 1B). Cultures containing BMP4 never contained more than 10 ± 6% Gsc-GFP+ cells, which is comparable to vehicle-treated cells. BMP4 also stimulated Mixl1 expression peaking at day 3 with 26 ± 12% Mixl1-GFP+ cells but further addition of activin did not affect the number of Mixl1-GFP+ cells (Fig. 1C). In Xenopus, Wnt molecules have both organizer-inducing and posteriorizing activities (Niehrs, 2004) and in mice Wnt3 is required for proper axis formation and induction of the primitive streak (Barrow et al., 2007; Liu et al., 1999). We therefore tested the ability of Wnt3a by itself or in combination with different doses of activin to induce PS markers. 5 ng/ml Wnt3a induced 23 ± 4% T-GFP+ cells (data not shown), whereas 100 ng/ml induced 30 ± 9% T-GFP+ cells at day 3, significantly higher than the control cells (Fig. 1A; p b 0.05). Activin did not have a significant effect on Wnt3a-induced T expression although we observed a tendency to reduced numbers of T-GFP+ cells with the highest doses of activin. The prominent induction of T expression seen with both BMP4 and Wnt3a was confirmed by immunofluorescence at day 3 (Fig. S2A). Wnt3a also induced Mixl1 expression. Cultures containing 100 ng/ml Wnt3a contained 11 ± 4% Mixl1-GFP+ cells at day 4. Notably, the combination of 100 ng/ml activin and 100 ng/ml

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Wnt3a induced 22 ± 3% Mixl1-GFP+ cells at day 4, approximately twice that achieved by either factor alone (Fig. 1C; p b 0.05). When examining co-expression of T and GFP at day 3 by immunofluorescence we found that most, if not all, Mixl1 expressing cells also expressed T, while the converse was not the case (Fig. S2B). While Wnt3a stimulated T and Mixl1 expression, it did not affect the number of Gsc-GFP+ cells (Figs. 1B and S2). The induction of Gsc expression by activin in the presence or absence of Wnt3a and its inhibition by BMP4 was confirmed by immunofluorescence at day 5 (Fig. S2C). Notably, the few T-expressing cells present in activin-treated GscGfp/+ cells after 5 days did not express GFP. Considering that T is found not only in the PS, but also in the emerging mesoderm at the late gastrula stage (Inman and Downs, 2006), this may indicate that mesoderm is also formed in activin-treated cultures. Collectively, the different ES lines all responded similarly to growth factor treatment (Fig. S2). Overall, our results indicate an anteriorizing role of activin during ES cell differentiation that can be modulated by the posteriorizing factors BMP4 and Wnt3a, consistent with the roles of these factors before and during gastrulation (reviewed in Tam and Loebel, 2007). BMP4 induces mesoderm and blocks activin-mediated induction of definitive endoderm To establish if our cultures contained embryonic or extraembryonic cell types we first examined expression of CXCR4 (chemokine (C-X-C motif) receptor-4), which is expressed in embryonic but not in extraembryonic tissues (McGrath et al., 1999; Sherwood et al., 2007). Using FACS analysis we compared surface expression of CXCR4 on cells isolated from dissociated E11 mouse embryo heads with that of differentiated ES cell progeny. Two distinct CXCR4-expression populations could be detected among cells from mouse embryos, a CXCR4lo and a CXCR4hi population (Fig. S3A). When ES cell progeny from either vehicle or activin-treated cultures was analyzed it was clear that the vast

Fig. 3. Activin-induced expression of the anterior primitive streak marker Gsc is inhibited by BMP4 but not by Dkk1. GscGfp/+ ES cells were grown in serum-free medium supplemented with one or more of the following growth factors or inhibitor; 10 ng/ml BMP4, 100 ng/ml Wnt3a, 320 ng/ml Dkk1, 3 or 100 ng/ml activin as indicated. (A) Triple-label immunofluorescence was performed to analyze the co-expression of E-cad (Cdh1), Gsc (GFP), and Sox17 in cells grown for 5 days under the indicated conditions. Note Cdh1+GFP−Sox17− cells (white arrows), Cdh1−GFP+Sox17− cells (red arrows), Cdh1+GFP−Sox17+ cells (white arrowheads), and Cdh1+GFP+Sox17+ cells (with yellow nuclei, red arrowheads) (B) Triple-label immunofluorescence was also performed to analyze co-expression of Cdh1, Gsc (GFP), and Oct4 on day 5. Note Cdh1+GFP−Oct4+ cells (white arrows), Cdh1−GFP+Oct4− cells (red arrows), and Cdh1+GFP+Oct4− cells (red arrowheads).

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majority of the cells were CXCR4hi (Fig. S3A), indicating that most cells, in both vehicles and activin-treated cultures, are embryonic rather than extraembryonic in nature. Furthermore, significant levels of Sox7 (SRYbox containing gene 7) transcripts, which is exclusively expressed in the extraembryonic part of the endoderm in the gastrula stage embryo (Kanai-Azuma et al., 2002), could only be detected in Wnt3a-treated cultures but not in vehicle, BMP4-, or activin-treated cultures (Fig. S3B). Having established the embryonic nature of the ES cell progeny in our cultures we next examined the expression of a number of germ layer specific markers. We initially analyzed expression of the transcription factor gene Foxa2 and the epithelial marker E-cadherin (E-cad; Cdh1), both of which are expressed in developing endoderm (Ang et al., 1993; Sasaki and Hogan, 1993). Immunofluorescent staining of cells grown in 3 or 100 ng/ml activin showed that these cultures contained many Foxa2+Cdh1+ cells compared to vehicletreated samples (Fig. 2). Notably, addition of BMP4 (10 ng/ml) but not Wnt3a (100 ng/ml) was able to drastically reduce the number of Foxa2+Cdh1+ cells induced by activin (Fig. 2). Analysis of VEGF receptor-2 (Kdr or Flk1) expression using Flk1-LacZ ES cells revealed that both BMP4 (with or without 100 ng/ml activin) and Wnt3a were capable of inducing Flk1-expressing cells (Fig. 2), indicative of mesoderm formation (Ema et al., 2006).

Wnt signaling augments the development of Sox17-expressing definitive endoderm induced by activin Based on the analysis of Foxa2 and Cdh1 expression it was not clear if the concentration of activin used influenced subsequent differentiation towards DE. Furthermore, analyses of Foxa2 and Cdh1 expression cannot distinguish between DE from different A–P positions. We therefore examined the number of GscGfp/+ cells that co-expressed GFP, Cdh1, and Sox17 by ICC as an indicator of anterior DE (ADE), in response to varying doses of activin with or without additional BMP4, Wnt3a, or Dkk1 treatment. Notably, we found that 100 ng/ml activin resulted in higher numbers of Cdh1+GFP+Sox17+ triple positive cells than seen with 3 ng/ml activin (Fig. 3A, compare panels b and c), supporting that efficient formation of ADE depends on the activin concentration (Yasunaga et al., 2005). Treatment with 3 ng/ml activin resulted in many Cdh1+ cells but the majority of these were not coexpressing GFP or Sox17 and most likely represent undifferentiated ES cells (see below). Most of the GFP+ cells generated in response to 3 ng/ ml activin were Cdh1−Sox17−, suggesting that they may represent mesoderm (Fig. 3A, panel b). Similarly, after treatment with Wnt3a alone (100 ng/ml) most GFP+ cells were Cdh1−Sox17− (Fig. 3A, panel d). We tested if Wnt signaling was required for the development of

Fig. 4. The requirement for canonical Wnt signaling during activin-induced Sox17 expression is more pronounced in aggregate culture than in adherent culture, and Dkk1 inhibits nodal-induced Sox17 expression more than activin-induced Sox17 expression. (A) Nodal/activin and Wnt signaling interactions were analyzed in Mixl1Gfp/+ and GscGfp/+ cells at days 2–6 of differentiation using flow cytometry. (B) GscGfp/+ cells cultured in the presence of Dkk1 prior to and during activin induction were analyzed by flow cytometry. (C) GscGfp/+ cells were induced to form embryoid bodies in the presence of the indicated growth factors and analyzed for GFP expression by flow cytometry after 5 days of culture. (D) Sox17Gfp/+ cells cultured for 5 days were analyzed for GFP expression by flow cytometry. The mean % GFP+ cells ± S.E.M. of three independent experiments is presented. (E) Sox17Gfp/+ cells cultured for 2–7 days in the presence of activin (30 or 100 ng/ml) were analyzed for GFP expression by flow cytometry at the indicated time points. (F) Sox17Gfp/+ cells were induced to form embryoid bodies in the presence of the indicated growth factors and analyzed for GFP expression by flow cytometry after 5 days of culture. The mean % GFP+ cells ± standard deviation of three independent experiments is presented for all flow cytometric analyses unless otherwise noted.

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Cdh1+GFP+Sox17+ triple positive cells in response to activin and found that such cells were still generated efficiently in response to 100 ng/ml activin if Dkk1 was included (Fig. 3A, panel f). In contrast, addition of BMP4 completely prevented the development of such cells (Fig. 3A, panel g). We often found clusters of Cdh1+GFP−Sox17− cells in cultures treated with different combinations of activin and Wnt3a (Fig. 3A, panels b–e). Immunocytochemistry revealed that many Cdh1+Gsc− cells were Oct4+ and thus likely represent undifferentiated ES cells (Fig. 3B). Attempts to quantify the relative number of Gsc-GFP+Cdh1+ and Gsc-GFP−Cdh1+ cells by FACS under the various conditions failed due to problems with achieving reliable FACS data using the Chd1 monoclonal antibodies available. Although it was not evident from the above experiments with the GscGfp/+ cells that canonical Wnt signaling influenced the expression of PS markers or the formation of DE in response to activin we wanted to examine closer if Wnt activity was required for expression of other PS markers and DE formation in our ES cell cultures since gastrulation and thereby also endoderm formation requires Wnt3 activity in vivo (Barrow et al., 2007; Liu et al., 1999) and because we detected expression of Wnt3 and Wnt3a in our differentiating ES cell cultures (Fig. S4). To test this notion, we first cultured Mixl1Gfp/+ cells with activin or Wnt3a in the presence or absence of 320 ng/ml Dkk1 or 1 μM of the ALK4/5/7-specific inhibitor SB431542, respectively. The concentration of the inhibitors was titrated by using a Wnt- or activinresponsive luciferase assay (data not shown), choosing the concentration that blocked the response to exogenous Wnt3a or activin, respectively, without causing non-specific toxicity in ES cells. When analyzing the number of Mixl1-GFP+ cells after 4 days of activin treatment (30 ng/ml) we found that these were significantly reduced when Dkk1 was included (26 ± 3% vs. 4 ± 3%; p b 0.05; Fig. 4A). Similarly, the number of Mixl1-GFP+ cells induced by 100 ng/ml Wnt3a was reduced from 11 ± 4% on day 4 to 3 ± 4% by simultaneous SB431542 treatment (Fig. 4A). Thus, activin and Wnt3a act cooperatively to induce Mixl1 expression and both signaling pathways are required for Mixl1 expression in ES cell progeny. Although Wnt3a only induced low numbers of Gsc-expressing cells, these were dependent on endogenous nodal/activin signaling as SB431542 significantly inhibited the development of Gsc-GFP+ cells in response to Wnt3a (Fig. 4A). In contrast, FACS analyses confirmed that activin-induced Gsc expression was not inhibited by Dkk1 treatment. Cultures of GscGfp/+ cells contained 17 ± 4% Gsc-GFP+ cells at day 5 when grown in 30 ng/ml activin and 23 ± 12% when cultured in the presence of activin and Dkk1 (Fig. 4A) consistent with the above mentioned immunofluorescent analysis that demonstrated that Gsc-GFP+Sox17+ Cdh1+ triple positive cells were efficiently generated in response to activin treatment, regardless of the presence of Dkk1. Undifferentiated ES cells have a low level of active canonical Wnt signaling despite the lack of exogenously added Wnt factors (Sato et al., 2004). To test if this low level of Wnt signaling has any influence on the later activation of Gsc expression we cultured undifferentiated GscGfp/+ cells for three passages in media containing Dkk1 before differentiation was induced by removing LIF and BMP4 and adding activin and Dkk1 for 5 days. We found that inclusion of Dkk1 prior to differentiation did not prevent activin from efficiently inducing Gsc expression (Fig. 4B). Furthermore, Dkk1 failed to prevent activin from inducing Gsc-GFP+ cells in aggregate culture (Fig. 4C). To obtain quantitative data on the number of DE cells after growth factor treatment we subjected a Sox17Gfp/+ reporter line (Kim et al., 2007) to our differentiation protocol. At mid-streak stage Sox17 expression marks the definitive endoderm forming adjacent to the anterior end of the PS. Simultaneous with the movement of DE to the anterior region of the gastrula, the Sox17 expression domain expands to include the endoderm underlying the neural plate of the early tailbud-stage embryo (Kanai-Azuma et al., 2002). Thus, Sox17 is an early marker that similar to genes such as Hex, Foxa2, and Cer1, are expressed simultaneously in the anterior visceral endoderm and the

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DE in the embryonic part of the gastrulating embryo. Sox17Gfp/+ cells express a low level of GFP (Sox17-GFPLo) when kept undifferentiated in the presence of LIF and BMP4 (not shown). Differentiation under neural promoting conditions (Ying et al., 2003a) results in the development of a Sox17-GFP− population and some remaining Sox17-GFPLo cells while treatment with activin for 5 days induces Sox17-GFPHi and Sox17-GFP− populations in addition to a remaining Sox17Lo population (Fig. S1). Increasing concentrations of activin resulted in development of progressively more Sox17-GFPHi cells with the highest numbers reached with 30 and 100 ng/ml of activin (Fig. 4D). We observed an increase in Sox17-GFPHi cells over time, peaking at day 5, followed by a modest decrease at days 6 and 7 (Fig. 4E). The induction of Sox17-GFPHi cells was at least partly dependent on Wnt signaling as treatment with Dkk1 reduced the number of GFPHi cells by ∼ 50% at the highest activin concentration (Fig. 4D). Notably, the number of Sox17-GFPHi cells induced by 1 μg/ml nodal appeared more strongly reduced in response to Dkk1 treatment than did a similar number of Sox17-GFPHi cells induced by 30 and 100 ng/ml of activin (Fig. 4D). Furthermore, when differentiation was performed in aggregate culture, which may rely more on endogenous signaling (Sachlos and Auguste, 2008; ten Berge et al., 2008), the number of activin-induced Sox17-GFPHi cells were strongly reduced by Dkk1 treatment (Fig. 4F). As expected, the addition of BMP4 prevented activin-induced formation of Sox17-GFPHi cells (p b 0.001), while addition of Wnt3a resulted in a marginal, but significant (p b 0.05) increase in the development of Sox17-GFPHi cells (Fig. 4D). To further define the time at which canonical Wnt signaling was required for the formation of T-, Gsc- and Sox17-GFPHi cells, we cultured the cells with activin for three (TGfp/+) or five (GscGfp/+ and Sox17Gfp/+) days and added either Wnt3a or Dkk1 for shorter periods

Fig. 5. Wnt signaling is required during late stages of activin-induced definitive endoderm formation. ES cells were cultured in serum-free medium with 100 ng/ml activin and supplemented with 100 ng/ml Wnt3a or 320 ng/ml Dkk1 for a variable number of days. TGfp/+ cells (A) were cultured for 3 days and GscGfp/+ (B) and Sox17Gfp/+ (C) cells were cultured for 5 days before being analyzed for GFP expression by flow cytometry. The mean % GFP+ cells ± standard deviation of three independent experiments is presented.

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during the 5 day activin treatment. To monitor the effectiveness of Dkk1 treatment in reducing canonical Wnt signaling we first generated an ES cell line stably transfected with a SuperTOP-Cerulean reporter (SuTOP-CFP) and established its ability to report Wnt signaling in E10.5 chimeric embryos after injection into E3.5 blastocysts and implantation into pseudo-pregnant females. As shown in Fig. S5A, native Cerulean fluorescence can be observed in several sites known to harbor active Wnt signaling at this stage, including the peripheral aspects of the otic vesicles (Maretto et al., 2003). When SuTOP-CFP cells were cultured in activin we found that CFP+ colonies developed at day 3 even in the absence of exogenous Wnt addition and that additional treatment with Dkk1 reduced or abolished reporter activity depending on the duration of treatment (Fig. S5B). Similarly, when SuTOP-CFP cells were assayed at day 5, we found that Dkk1 treatment at days 3–4 or 4–5 inhibited reporter activity (Fig. S5C). Thus, treatment with Dkk1 for as little as 1 to 2 days at the concentration used appeared quite effective in suppressing SuTOP reporter activity. We then analyzed the number of T-GFP+ cells developing in response to activin when Wnt signaling was experimentally perturbed. T-GFP+ cell numbers were augmented by Wnt3a with the largest effect reached when Wnt3a was added only at day 3. Treatment with Dkk1 on the other hand reduced the number of TGFP+ cells most effectively when included in that last part of the three day period (Fig. 5A). Neither stimulating nor inhibiting Wnt signaling in GscGfp/+ cells, by treatment with Wnt3a and Dkk1, respectively, resulted in a significant change in the number of activininduced GFP+ cells irrespective of the treatment period (Fig. 5B). However, treatment of Sox17Gfp/+ cells with Wnt3a prior to appearance of GFP+ cells reduced the number of Sox17-GFPHi cells, while later Wnt3a treatment had no effect. Conversely, treatment with Dkk1 reduced the number of Sox17-GFPHi cells only if present after the first appearance of these (Fig. 5C).

We next used siRNA mediated knock-down of β-catenin (encoded by Ctnnb1) to confirm that inhibition of canonical Wnt signaling at the latter part of the 5 day activin stimulation period could suppress the appearance of Sox17-GFPHi cells and to further test the GscGfp/+ cell line which appeared refractory to Wnt inhibition in previous experiments. We transfected GscGfp/+ and Sox17Gfp/+ cells with control and two different Ctnnb1 siRNAs at day 2 of differentiation and assayed β-catenin expression by western blotting at days 3 and 5. In GscGfp/+ cells we found a reduction of β-catenin expression at day 3 which was normalized at day 5. siRNA treatment appeared more effective in Sox17Gfp/+ cells with strong inhibition of β-catenin expression at day 3 which was only partly recovered by day 5 (Fig. 6A). FACS analysis at day 5 showed a reduction in the number of Gsc-GFP+ cells after β-catenin knock-down, although this only reached significance in Ctnnb1 siRNA2 treated samples (p b 0.05, Fig. 6B). Furthermore, Q-RT-PCR analysis of GscGfp/+ cells after βcatenin knock-down did not reveal significant changes in expression of Lhx1 and Chrd (Fig. S6). However, in agreement with the results obtained with Dkk1 treatment, we found a prominent reduction in the number of Sox17GFPHi cells at day 5, in both siRNA1 and siRNA2 treated samples (p b 0.05, Fig. 6B). Activin dose-dependently induces an anterior gene expression pattern To determine whether the DE formed in the presence of high concentrations of activin was anterior or posterior in character we analyzed the expression of a number of markers displaying differential expression depending on the anterior–posterior position of the cells. RT-PCR analyses at day 5 showed that activin, regardless of concentration, could induce expression of genes associated with anterior cell fates, including Lefty1, Hex, (Martinez-Barbera et al., 2000) and Otx2, (Rhinn et al., 1998). However, robust expression of

Fig. 6. Inhibition of canonical Wnt signaling inhibits activin-induced Sox17-GFPHi definitive endoderm formation, but has little effect on Gsc expression. (A) Western blot analysis of β-catenin expression after siRNA mediated knock-down. Two different siRNAs targeting β-catenin (siRNA1 and siRNA2) or a scrambled control siRNA were introduced to GscGfp/+ and Sox17Gfp/+ cell lines on day 2 and β-catenin levels assayed at day 3 and 5. An anti-β-actin western blot served as loading control. (B) siRNA treated GscGfp/+ and Sox17Gfp/+ cells were analyzed for GFP expression by flow cytometry after 5 days of serum-free culture supplemented with 0, 3 or 100 ng/ml activin. The mean % GFP+ cells ± standard deviation of three independent experiments is presented. Untransfected and mock transfected controls are also shown.

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Cer1 as well as repression of the posterior marker Cdx2 was only seen in the presence of the high activin concentration (Fig. 7A). In contrast, Cdx2 (Beck et al., 1995) was expressed in samples treated with 10 ng/ ml BMP4 or 100 ng/ml Wnt3a, as well as cells treated with the low dose of activin (Fig. 7A). We next isolated Gsc-GFP+ and Gsc-GFP− cells as well as Sox17-GFPHi, Sox17-GFPLo and Sox17-GFP− cells from activin-stimulated cultures at day 5 by FACS and prepared RNA for gene expression analysis. Message for Otx2 and Cer1 was detected in both Gsc-GFP+ and Gsc-GFP− fractions, but with an enrichment in the GFP+ fraction (Fig. 7B). Cer1 was also enriched in Sox17-GFPHi cells compared to Sox17-GFP− and Sox17-GFPLo cells (Fig. 7B). Furthermore, when analyzing the expression of the respective DE and VE markers, Pyy and Tdh (Hou et al., 2007; Sherwood et al., 2007) we only found Pyy in the Sox17-GFPHi cells while Tdh was found in the Sox17-GFP− and, to a lesser degree, Sox17-GFPLo populations (Fig. 7B). However, it should be noted that FACS sorted populations are probably not completely pure. For example, the Sox17Hi and Sox17Lo populations are likely displaying “cross-contamination” to some extent. Finally, as Lhx1 is expressed in the anterior part of the PS in the latestreak embryo and expected to co-localize with Gsc (Shawlot and

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Behringer, 1995; Tada et al., 2005) we performed GFP-Lhx1 double immunofluorescent stainings on Gsc Gfp/+ cells after 5 days of differentiation. As expected, we found Gsc-GFP+Lhx1+ double positive cells in activin-treated cultures and the formation of these cells was inhibited by the addition of BMP4 but not Wnt3a (Fig. 7C). FGF receptor signaling is required for mesendoderm and endoderm differentiation If FGF signaling is compromised during early development as in FGF8, FGFR1, or UDP-glucose dehydrogenase mutants, the embryos' arrest at gastrulation and no embryonic mesoderm- or endodermderived tissues develop (Ciruna et al., 1997; Deng et al., 1994; GarciaGarcia and Anderson, 2003; Sun et al., 1999; Yamaguchi et al., 1994). The phenotype is associated with a failure of migration and it is unclear to what degree FGF signaling regulates allocation of mesodermal and endodermal fates. However, studies in zebrafish indicate that FGF signaling promotes mesodermal development at the expense of endodermal development (Mathieu et al., 2004; Mizoguchi et al., 2006; Poulain et al., 2006). We first examined the

Fig. 7. Activin, BMP and Wnt signaling influence anterior–posterior patterning during differentiation of ES cells. ES cells were grown for 5 days in serum-free medium supplemented with indicated growth factors. (A) The expression of Lefty1, Hex, Otx2, Cer1 and Cdx2 was analyzed in TGfp/+ cells by RT-PCR. The expression of TATA binding protein (Tbp) was used as internal standard. (B) ES cells were cultured with 100 ng/ml activin and FACS sorted based on native GFP fluorescence after 5 days of culture. The expression of Otx2 and Cer1 was analyzed in the GFP+ and GFP− fraction of GscGfp/+ cells, and Otx2, Cer1, Pyy and Tdh expression was analyzedin the GFPHi, GFPLo and GFP− fraction in Sox17 Gfp/+ cells. Glucose-6phosphate dehydrogenase (G6pd) was used as internal standard. (C) The expression of Lhx1 and GFP was examined in GscGfp/+ cells by immunofluorescence after 5 days in the presence of the indicated factors.

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expression of members of the FGF family in differentiating ES cell cultures treated with activin and BMP4 (alone or in combination). We found that FGF4 and FGF8 were expressed in ES cell cultures treated with activin or BMP4 (Fig. S4). We therefore addressed the role of FGF signaling by culturing the cells in activin (100 ng/ml) or BMP4 (10 ng/ml) with or without the FGF receptor inhibitor SU5402 (Mohammadi et al., 1997) or in the absence or presence of exogenously added FGF2. Consistent with a recent report (Kunath et al., 2007), flow cytometric analyses of undifferentiated cells (day 0) and at days 3 and 5 of differentiation revealed that both BMP4induced T and Mixl1 expression were strongly inhibited by 10 μM SU5402 (Fig. 8A), a concentration of SU5402 that did not compromise cell viability (data not shown). As BMP4-induced T expression has been shown to be insensitive to SU5402 in EB culture (Willems and Leyns, 2008), we repeated this experiment to determine if choice of culture system could explain this difference. Indeed we also find that BMP4-induced T expression is unaffected by addition of SU5402, and even by itself SU5402 is able to induce T expression (Fig. 8B). Nevertheless, addition of soluble FGF receptors (sFGFRs, b and c splice forms) failed to inhibit BMP4-induced T or Mixl1 expression at day 3 of adherent culture (Fig. 8A). Conversely, addition of FGF2 augmented the number of T-GFP+ and Mixl1-GFP+ cells formed in response to BMP4 treatment (from 39 ± 23% to 68 ± 6% and from 41 ± 8% to 53 ± 8%, respectively). However, this induction was only significant for Mixl1-GFP (p b 0.01). From studies in zebrafish we would expect that FGF signaling would inhibit endoderm development and redirect cells towards mesoderm (Mizoguchi et al., 2006; Poulain et al., 2006). When GscGfp/+ cells were cultured in the presence of activin with addition of FGF2, SU5402, PD173074, or soluble FGF receptors we observed a strong dependence on FGF signaling for the development of Gsc-GFP+ cells (Figs. 9A and B). The number of Gsc-GFP+ cells appearing at day 5 in the presence of activin was significantly inhibited by the addition of SU5402 (p b 0.05), PD173074 (p b 0.001) or sFGFRs

(p b 0.05). Furthermore, both the b and c splice forms of soluble FGF receptors reduced the number of Gsc-GFP+ cells (p b 0.05 using Student's one-tailed t-test). Moreover, the addition of FGF2 significantly enhanced the amount of Gsc-GFP+ cells seen at day 5 in activin-treated cultures (p b 0.01). These results demonstrate that activin-induced Gsc expression depends on FGF signaling and suggest that anterior PS fate is augmented by FGF signaling. However, as Gscexpressing cells develop further into both meso- and endoderm we next asked whether development of definitive endoderm is also FGF dependent. Hence we examined if activin-induced formation of Sox17GFPHi cells was influenced by FGF signaling. Culture of Sox17Gfp/+ cells in activin together with SU5402 or PD173074 resulted in a ∼ 50% and ∼90% reduction in the number of Sox17-GFPHi cells compared to activin alone (p b 0.001; Fig. 9B), demonstrating that FGFR signaling is required for normal formation of definitive endoderm. However, in contrast to results obtained with the Gsc reporter, addition of sFGFR did not affect Sox17 expression. Moreover, where the addition of FGF2 boosted the formation of Gsc-GFP+ cells it resulted in an almost 50% reduction in the number of Sox17-GFPHi cells (p b 0.005; Fig. 9B), suggesting that precise control of FGF levels is important for regulation of Sox17 promoter activity. To better define the time where FGF signaling acted during induction of Gsc- and Sox17 expression we cultured Gsc- and Sox17GFP cells with activin for 5 days and added either FGF2, SU5402, or PD173074 for shorter periods. The (modest) stimulatory effect of FGF2 upon Gsc-GFP expression required that FGF2 was present early in the 5 day culture period, prior to the initial appearance of GFP+ cells (Fig. 9C). In contrast, SU5402 and PD173074 reduced the number of Gsc-GFP+ cells both when added early and late in the culture period (Fig. 9C). In contrast, the number of Sox17-GFPHi cells was reduced when FGF2 and FGFR inhibitors were present after the initial appearance of GFPHi cells (Fig. 9C). We used RT-PCR to analyze whether the expression of additional markers, informative in relation to anterior–posterior positional

Fig. 8. FGF receptor signaling is required for BMP4-induced mesendoderm formation in adherent, but not in aggregate culture. (A) TGfp/+ and Mixl1Gfp/+ cells were cultured in medium containing 10 ng/ml BMP4, with or without 100 ng/ml FGF2, 10 μM SU5402, or different concentrations of soluble forms of FGF receptors (b or c splice forms, or a combination of both). Cells were analyzed for GFP expression by flow cytometry on days 0, 3 and 5. (B) TGfp/+ were induced to form embryoid bodies with the indicated growth factors and analyzed for GFP expression by flow cytometry on day 3. The mean % GFP+ cells ± standard deviation of three independent experiments is presented.

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Fig. 9. FGF receptor signaling is required throughout the five day period for activin-induced Gsc expression but only required late for activin induction of Sox17-expressing definitive endoderm. GscGfp/+ and Sox17Gfp/+ cells were cultured in medium containing 100 ng/ml activin, in combination with 100 ng/ml FGF2, 10 μM SU5402, 100 nM PD173074, or soluble forms of FGF receptors as indicated. (A) Time course analysis of appearance of Gsc-GFP+ cells by flow cytometry at time 0 and after 3 and 5 days in culture. (B) GscGfp/+ and Sox17Gfp/+ cells were analyzed for GFP expression at day 5 after culture with the indicated factors. (C) GscGfp/+ and Sox17Gfp/+ cells were analyzed for GFP expression at day 5 after culture with activin and either FGF2, SU5402, or PD173074 for shorted periods as indicated. The mean % GFP+ cells ± standard deviation of three independent experiments is presented.

identity, were affected by blocking FGF receptor signaling in activintreated cells. Expression of Otx2, Chrd, and Cer1, genes that are repressed by BMP4, is not affected by the addition of SU5402 after 3 days (Fig. 10). Among the markers analyzed only Bmp2, which is first expressed in the embryo proper at E7.5 just lateral to the anterior neural folds and in precardiac mesoderm and slightly later in foregut endoderm (Winnier et al., 1995), is repressed both by BMP4 and SU5402 at day 3. Chrd expression appears at day 3 in vehicle-treated cultures and at day 5 in BMP4-treated cultures but in both cases it is sensitive to SU5402. Also BMP4-induced expression of mesoderm markers Chrd, and Nog at day 5, as well as the posterior markers Bmp4 and Cdx2, is sensitive to SU5402 (Fig. 10 and Fig. S4). Foregut and pancreatic competence of ES cell-derived endoderm To test if the DE-like cells had potential to functionally integrate into developing embryonic endoderm, we implanted approximately 50 cells labeled with a fluorescent dye into 6 to 10 somite stage chicken

embryos at the level of the prospective pancreatic endoderm and incubated for 48 h. When ES cell progeny from activin-treated cultures were grafted, 15 out of 21 transplanted embryos contained dyelabeled, Foxa2+ cells incorporated into the endoderm. In contrast, only 3 out of 20 embryos receiving cells cultured without activin and 0 of the 8 embryos receiving activin- and BMP4-treated cells contained dye-labeled cell in the endoderm (Figs. 11A, D, G). Frequently, the activin-treated cells incorporated in the Nkx6-1+Pdx1+ pancreatic endoderm (Figs. 11E, F) (Pedersen et al., 2005). In a second series of grafting experiments we tested if the Sox17-GFPHi cells obtained after activin treatment for 5 days with or without additional Dkk1 treatment were capable of incorporating into chick foregut endoderm. Such cells were equally capable of contributing to the foregut endoderm (Figs. 11H, I, K–M), although the number of Sox17-GFPHi cells were reduced in cultures containing Dkk1. Orthogonal projections of the confocal stacks obtained from grafted embryos suggested expression of Nkx6-1 in some of the grafted cells (Fig. 11J). Considering the anterior markers expressed by activin-treated cells, and the absence of pancreatic or

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Fig. 10. TGF-β, Wnt and FGF signaling control the dynamic gene expression in differentiating ES cells. TGfp/+ cells cultured for 3 or 5 days in serum-free medium supplemented with 100 ng/ml activin, 10 ng/ml BMP4 and/or 10 μM SU5402, as indicated, were analyzed by RT-PCR. The expression of Otx2, Chrd, Cer1, Bmp2, Bmp4 and Cdx2 was analyzed. Tbp was used as internal standard.

other regional foregut markers at day 5 (Fig. 12), the grafting experiments suggest that DE formed in our cultures can respond to posteriorizing cues from the embryonic environment and progress further in differentiation. We therefore tested if the cells were able to respond to suspected posteriorizing cues in vitro. After 5 days of culture in activin, the cells were shifted to conditions where activin was replaced by candidate posteriorizing factors Wnt3a, FGF4, and retinoic acid (RA) (Grapin-Botton and Constam, 2007) and analyzed for expression of Sox2, Pdx1, and Cdx2, markers of fore-, mid-, and hindgut, respectively (Grapin-Botton and Melton, 2000). Remarkably, we found large numbers of Sox2+Foxa2+ cells in cultures treated with posteriorizing factors (Fig. 12). We confirmed Sox2 expression using the OS25 cell line which carries a β-geo reporter gene in the Sox2 locus (Li et al., 1998). β-galactosidase activity was visualized with X-gal staining (Fig. 12). The appearance of Sox2+ cells was depending on neither FGF4 nor RA. Scattered Pdx1+ cells also appeared but in contrast to Sox2+ cells these only appeared in the presence of RA. Cdx2 positive cells were not detected under any of the conditions tested. Together, our data demonstrate that DE formed from mES cells in adherent monoculture is capable of differentiation toward foregut endoderm in vivo and in vitro but that only a limited number of these cells appear to respond to posteriorizing cues in vitro. Discussion A recent work from Smith et al. has demonstrated that mES cells can be kept pluripotent under defined serum- and feeder-free conditions (Ying et al., 2003a), and be efficiently converted to Sox1+ neuroectodermal progenitors when kept in adherent monoculture in the absence of LIF and BMP4 (Ying et al., 2003a,b). In the present work, we extend this defined system to demonstrate that mES cells kept in serum- and feeder-free adherent monoculture respond dose-dependently to inducers of primitive streak formation by developing mesendoderm with capacity to differentiate further into cells resembling foregut endoderm, in vivo and in vitro. Although several recent studies have established that the TGF-β family members BMP4 and activin induce mesodermal and endodermal gene expression in differentiating mES cells (Gadue et al., 2006; Kubo et al., 2004; Mossman et al., 2005; Ng et al., 2005; Tada et al., 2005; Yasunaga et al., 2005), the varied conditions under which differentiation was induced; e.g. adherent vs. suspension culture, differences in basal cell culture media and supplements, cell density, absence or presence of serum; as well as the different reporter lines

utilized make a direct comparison of these studies difficult. Here we use a series of GFP-based reporters to study the dynamic expression of T, Mixl1, Gsc, and Sox17 after manipulating one or more of the FGF, TGF-β, and Wnt signaling pathways. Remarkably, we find that the lowest concentration of activin used (3 ng/ml) induced the highest number of T-GFP+ cells, whereas higher concentrations (10–100 ng/ml) resulted in progressively fewer T-GFP+ cells. These data apparently conflict with results obtained by Keller et al. who observed increasing numbers of TGFP+ cells with increasing activin concentration until a plateau of about 50–60% T-GFP+ cells was reached at 30 ng/ml of activin (Kubo et al., 2004). It is not entirely clear why a direct correlation between activin concentration and the number of T-GFP+ cells is observed by Kubo et al. while we observe an inverse correlation. One possible explanation relates to the embryoid body formation used by Kubo et al. in which only cells located at the periphery, directly exposed to the cell culture medium, may experience the full concentration of added growth factor as recently demonstrated (Sachlos and Auguste, 2008). Cells located in the interior of the embryoid body are most likely experiencing a lower concentration and the overall dose–response curve may therefore appear shifted towards higher concentrations. It is also possible that the environment of the embryoid body is more conducive to secondary signaling events that may influence the number of T-GFP+ cells. A recent study did however report that T mRNA levels in differentiating EBs were inversely correlated with activin concentration within the 5–50 ng/ml range (Willems and Leyns, 2008). Moreover, our results are strikingly similar to data from Xenopus animal cap explants, where activin dosedependently controls cell fate specification. The expression of the Xenopus T homolog Xbra is induced by low concentrations of activin while higher concentrations induce the expression of the Gsc homolog Xgsc (Green et al., 1992; Gurdon et al., 1994; Latinkic et al., 1997). However, even Xgsc-expressing cells induced by high doses of activin have undergone a transient Xbra-expressing phase, but direct repression of Xbra transcription by binding of Xgsc to the Xbra promoter limits the duration of Xbra expression (Latinkic et al.,1997). In this regard, analyses of the duration of T expression by time-lapse microscopy under different conditions would be highly informative. The notion of activin as a dosedependent inducer of anterior fate in ES cell progeny is indicated by several observations: the differential effect of 3 and 100 ng/ml activin on gene expression such that only the high dose induces the anterior marker Cer1 and represses the posterior marker Cdx2. Also, extensive co-localization of E-cadherin, Gsc-GFP and Sox17 was only observed after treatment with 100 ng/ml of activin. Additional marker analyses

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Fig. 11. mES cell-derived endoderm grafts contribute to the developing chicken endoderm. (A–F) ES cells were cultured for 5 days in serum-free medium containing vehicle, 100 ng/ml activin or 100 ng/ml activin and 10 ng/ml BMP4 as indicated. Small clusters of cells were labeled with the red fluorescent cell tracker dye CMTMR and grafted between the endoderm and mesoderm of explanted chicken embryos and allowed to develop for 2 days before being processed for confocal immunohistochemistry. (H–M) Sox17Gfp/+ cells were cultured for 5 days in serum-free medium containing either 100 ng/ml activin or 100 ng/ml activin plus 320 ng/ml Dkk1. Small clumps of GFP+ cells were labeled with the red fluorescent cell tracker dye CMTMR and grafted between the endoderm and mesoderm of explanted chicken embryos and allowed to develop in ovo for 2 days before being processed for confocal immunohistochemistry. (A–F, H, I, K, L) Optical sections of chicken embryos whole-mount stained with the indicated antibodies and transplanted with ES cell progeny developing after 5 days with the indicated growth factors. (J) Orthogonal view of a Z-stack revealing an Nkx6-1-expressing CMTMR-labeled cell. (G, M) Tabulated results of the grafting experiments. Arrowheads in A–F and H–L indicate implanted cells. dp: dorsal pancreas, e: endoderm, fp: floor plate, n: notochord, nt: neural tube, vp: ventral pancreas.

revealed that such cells were also Pyy+, Foxa2+ and CXCR4+ but Sox7− and Tdh−, suggesting embryonic rather than extraembryonic endoderm had formed (Kanai-Azuma et al., 2002; Tada et al., 2005; Yasunaga et al., 2005). Conversely, BMP4- and Wnt3a-treated cells expressed low levels of anterior markers and instead expressed the posterior markers Bmp4 and Cdx2. Furthermore, BMP4 prevented activin-induced gene expression and stimulated the expression of T and Mixl1. These observations are consistent with previous in vivo data showing that BMP4 is necessary for T expression (Winnier et al., 1995) and that activin-

induced Gsc expression can be counteracted by BMP4 in vivo (Imai et al., 2001; Jones et al., 1996; Sander et al., 2007; Shapira et al., 1999; Steinbeisser et al., 1995). Additionally, Keller et al. also noted a posteriorizing effect of BMP4 upon mouse ES cells derived, activininduced PS populations (Nostro et al., 2008) and Suemori et al. found that inhibition of BMP signaling redirected human ES cell-derived mesodermal cells (induced by forced expression of stabilized β-catenin) towards an anterior PS/DE lineage, in a process dependent on activin/ nodal signaling (Sumi et al., 2008).

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Fig. 12. ES cell-derived endoderm acquire foregut cell fates. (A–J′ and P–Z′) OS25 (Sox2βgeo/+) or (K–O) Pdx1LacZ/+ cells were cultured for 5 days in 100 ng/ml activin and subsequently treated with 5 ng/ml Wnt3a, 10 ng/ml FGF4 and/or 0,1 μM RA for 3 days as indicated. The co-expression of Foxa2 and Sox2 (foregut), Pdx1 (midgut) or Cdx2 (hindgut) was analyzed by immunofluorescence. The expression of Sox2 (A–E) and Pdx1 (K–O) was confirmed by analyzing β-galactosidase activity using X-gal staining.

Further analysis of BMP4- (and Wnt3a-) treated cells revealed the presence of Flk1+ cells demonstrating that mesodermal differentiation had occurred. Interestingly, Smith et al. recently reported that BMP4 treatment of mES cells, under conditions nearly identical to ours, resulted in cells of unknown identity that were unlikely to be mesodermal based on presence of E-cad immunoreactivity (Kunath et al., 2007). While we occasionally detect a faint plasma membrane staining in BMP4-treated cells (see for example Fig. 3), the intensity of the staining is strongly reduced compared to the signal seen in activintreated cells, and most cells appear E-cad− in our experiments. Furthermore, the presence of Flk1 expressing cells would appear to conclusively demonstrate mesodermal differentiation. This is also consistent with a number of studies where embryoid bodies are stimulated with BMP4 (Czyz and Wobus, 2001; Finley et al., 1999; Johansson and Wiles, 1995; Lengerke et al., 2008; Ng et al., 2005; Willems and Leyns, 2008).

Our examination of the role of Wnt signaling during mesendoderm differentiation seems to indicate a major difference between differentiation in embryoid bodies and in adherent monoculture. Apparently, the extent to which Wnt signaling is required to induce anterior PS formation is different in the two systems. Gadue et al. found that 100 ng/ml Wnt could induce formation of Foxa2-CD4+T-GFP+ anterior PS-like cells, and that this was dependent on endogenous ALK4/5/7 signaling. Moreover, the ability of activin to induce the same fate was dependent on Wnt signaling since Dkk1 could prevent the effect of activin (Gadue et al., 2006). At first glance these findings may appear to contrast with our results, namely that the same concentration of Wnt3a could not induce significant numbers of GscGFP+ in adherent culture and that Dkk1 could not prevent induction of Gsc-GFP+ cells by activin. However, the Gsc-GFP+ cells in this work do not express Brachyury and very likely represent a mixed population and are thus difficult to compare to the Foxa2/T double positive cells

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studied by Gadue et al. Furthermore, although siRNA mediated knockdown of β-catenin significantly reduced the number of Gsc-GFP+ cells developing in response to activin the effect never exceeded 50%. In contrast, the formation of Sox17-GFPHi DE cells was more sensitive to inhibition of canonical Wnt signaling, but only when signaling was inhibited at a relatively late stage of development, suggesting that the requirement for Wnt signaling is not during the formation of PS cells but may rather represent a Wnt-mediated lineage choice by a common mesendodermal precursor or alternatively, a requirement for Wnt signaling in the maintenance of Sox17-expressing DE cells. Indeed, our observation that Dkk1 only reduces the number of Sox17Hi cells if present at day 4, i.e. after Sox17 expression is initiated, is consistent with a requirement for Wnt signaling in the maintenance of DE. Also, our observations are consistent with the requirement for βcatenin function in DE to acquire or maintain DE fate (Lickert et al., 2002) as well as the observation of synergy between Sox17 and βcatenin in the activation of DE gene expression (Sinner et al., 2004). We also speculate that the proposed action of Wnt3 during gastrulation might offer at least a partial explanation for the difference in sensitivity towards Dkk1 treatment observed between activin and nodal as well as between adherent and aggregate culture. Wnt3 is required in vivo for nodal to initiate its auto-stimulatory cascade. In the absence of Wnt3, nodal expression is initiated but then regresses rather than expands (Barrow et al., 2007; Liu et al., 1999). Wnt3 is thought to induce expression of Cfc1 which encodes the obligate nodal co-receptor Cripto (Morkel et al., 2003). It was recently reported that the three dimensional environment in embryoid bodies prevent exogenously added growth factors from diffusing more than a limited distance into the embryoid body (Sachlos and Auguste, 2008) possibly making these more reliant on endogenous relay signaling via nodal which subsequently would require Wnt activity to induce Cripto expression. This notion is supported by our observation that nodalinduced Sox17-GFPHi cells appear more sensitive to Dkk1-mediated suppression than activin induced Sox17-GFPHi cells, and by our observation that activin-induced Sox17-GFPHi cells are more sensitive to Dkk1 treatment in aggregate culture compared to adherent culture. The apparent absence of such an autocrine nodal/Wnt signaling loop in adherent culture is not surprising given the remarkable instability of nodal when secreted by cultured cells (Le Good et al., 2005). Nodal may simply be degraded before it can reach other cells in the dish whereas in the confined environment of an embryoid body it may well signal to nearby cells and such signaling might propagate in a relay fashion. Thus, Wnt3a treatment would not be sufficient to induce this loop in adherent culture. Conversely, since all the cells in adherent culture are exposed evenly to exogenous activin there may be no requirement for Wnt signaling to facilitate ALK4 signaling. Nevertheless, Gsc-GFP+ cells appear largely refractory to the inhibitory effects of Dkk1 treatment. Since Wnt induced expression of Xgsc is mediated by two homeodomain proteins (Siamois and Twin which have no apparent homologs in the mouse) binding to the PE-element, this finding raises the question of whether the conserved Wntresponsive element in the mouse Gsc promoter (Watabe et al., 1995) is functional in all contexts. It is possible that Gsc is not always induced by Wnt as Gsc expression is reduced rather than increased in E8.5 Dkk1 mutant embryos (Lewis et al., 2008). Lastly, we cannot rule out a more trivial explanation related to the particular Gsc-GFP cell line, which may potentially harbor a defect in its responsiveness to Dkk1-mediated Wnt inhibition. It is possible that more efficient knock-down of β-catenin than we achieved might further reduce or even prevent the formation of Gsc-GFP+ cells. In contrast, we do observe a requirement for Wnt signaling for induction of Mixl1 expression. Our experiments revealed the simultaneous requirement of nodal/activin and Wnt signaling to induce expression of Mixl1 in ES cell progeny. Stimulation of one pathway while inhibiting the other reduced Mixl1 induction, which corroborates previous findings (Gadue et al., 2006). The Mixl1 promoter has

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previously been shown to be nodal/activin-responsive via the presence of Foxh1 and Smad binding sites in the promoter (Hart et al., 2005) and inspection of the published sequence reveals the presence of consensus TCF/LEF binding sites in the promoter as well. In vivo, Wnt/β-catenin signaling is upstream of two distinct gene expression programs acting during anteroposterior axis and mesoderm formation, respectively (Morkel et al., 2003). We suspect that we observe a requirement for Wnt signaling in mesoderm formation but did not pursue this aspect further. The two differentiation systems, adherent and aggregate culture, also differ strikingly in the requirement for FGF signaling. In agreement with our results a recent study using embryoid body formation found that FGF signaling was not required for BMP4-induced T expression (Willems and Leyns, 2008), which is unlike the situation in adherent culture where BMP4-induced T expression is completely prevented by SU5402. Moreover, consistent with our results Kunath et al., also using adherent culture, recently reported that FGF4 deficiency or treatment with PD173074 prevented the switch from BMP4-mediated support of pluripotency to BMP4-induced differentiation (Kunath et al., 2007). The requirement for FGF signaling in induction of mesodermal gene expression is also consistent with the in vivo requirement for FGF signaling in Xenopus and zebrafish mesodermal induction (Cornell et al., 1995; Mathieu et al., 2004). In zebrafish, FGF signaling is required downstream of Nodal for induction of One-eyed-pinhead, the zebrafish homolog of Cripto (Mathieu et al., 2004). However, as discussed above activin does not require Cripto to activate its receptors, thus activin induced Sox17 expression should not necessarily be inhibited by FGFR inhibitors. This is indeed what we find when inhibiting FGF signaling prior to appearance of Sox17-GFPHi cells, the time where one would expect to find a requirement for Cripto function if Nodal was the inducer. It might be illuminating to determine if Nodal induced Sox17 expression would be sensitive to these inhibitors. Nevertheless, we do observe that Sox17 expression depends on FGF signaling after the initial appearance of Sox17-GFPHi cells at the same time where we observe a dependency for Wnt signaling. It thus appears that maintenance of Sox17 expression and/or propagation of Sox17 expressing cells depend on both FGF and Wnt signaling. Consistent with our results, Brickman et al. also observed an absolute requirement for FGF signaling at days 3–7 for the formation of Hex+CXCR4+ADE cells when differentiating ES cells in monolayer culture under defined conditions (Morrison et al., 2008). In some cases we noticed what appeared to be conflicting results when inhibiting FGF and FGFR signaling by soluble FGF receptors and SU5402, respectively. It is possible that this is a reflection of FGF independent FGFR activation. FGF receptors are known to form complexes with N-cadherin and N CAM in neurons and in pancreatic β-cells, resulting in ligand independent receptor activation (Cavallaro et al., 2001; Saffell et al., 1997; Williams et al., 1994), and this would not be expected to be sensitive to the addition of soluble FGF receptors. More trivial explanations are also possible. We cannot rule out that SU5402 may have unknown non-FGFR-mediated effects or be certain that our soluble FGFR preparations are capable of inhibiting all FGF family members. Additional experiments with dominant negative receptors and cell lines mutated in genes coding for FGF signaling components will likely shed more light on these questions. The expression of anterior markers such as Otx2 and Cer1 in Sox17GFPHi cells suggest that these could be anterior definitive endoderm. This notion is supported by the selective presence of Pyy transcripts in this population, but the lack of good ADE specific markers prevents us from categorically making this conclusion. However, in vivo ADE forms under conditions of high nodal signaling (Ben-Haim et al., 2006; Lu and Robertson, 2004; Vincent et al., 2003) which we believe we mimic with culture conditions containing 30–100 ng/ml activin or 1 μg/ml nodal. Thus, it is likely that the Sox17-GFPHi cells formed in our cultures represent ADE. However, it is also likely that mesoderm is formed to some extent in cultures treated with high doses of activin or

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nodal. The presence of Otx2 transcripts in Sox17-GFPLo and -GFP− populations suggest that these also contain anterior cell types including perhaps small numbers of AVE cells. The latter is indicated by the presence of the VE marker Tdh and the presence of Otx2 and Cer1 is consistent with this idea. Lastly, we cannot exclude that Sox17-negative endoderm is present in our cultures. Chick embryo xenografting has previously been used to investigate the developmental potential of embryonic mouse cells (FontainePerus, 2000; Fontaine-Perus et al., 1996, 1997, 1995) as well as mouse and human ES cell derivatives (Lee et al., 2007; Wichterle et al., 2002) but to our knowledge we report the first functional assay for mES cellderived endoderm. Previous studies have relied almost exclusively on expression of cell-specific markers for the characterization of in vitro generated endoderm. Although the ES cell-derived endoderm did not express transcription factors associated with the regionalization of the primitive gut tube after 5 days of activin treatment, the cells were capable of turning on these genes when integrating into endodermal epithelium in vivo. Moreover, some embryos contained grafted cells in the Nkx6-1+ and Pdx1+ pancreatic endoderm. Overall, our data suggest that ES cell-derived DE, formed under defined conditions, can contribute to the developing gut tube and further suggests that such cells are, at least partly, capable of responding to patterning cues from the in vivo environment. The latter notion is corroborated by the induction of pancreas markers in a limited number of cells when such cells are cultured further in the presence of known and suspected posteriorizing factors. It is remarkable how the requirement for certain signaling events are strikingly different when comparing aggregate culture (i.e. embryoid body formation) and adherent culture. However, we do not think that one system is superior to the other but rather that comparison of directed differentiation under adherent and aggregate culture conditions will prove valuable when attempting to decipher the extent of secondary signaling events and their role in lineage selection. In this regard it is also noteworthy that endogenous signaling likely plays a prominent role also in adherent culture. This notion is based on several of our observations, including that many SuTOP-CFP+ cells formed in cultures that received activin as the only exogenous factor, the interdependence between activin and Wnt signaling for generation of Mixl1-GFP+ cells, and on the dependence on Wnt and FGF signaling for efficient activin-induced formation of Sox17-GFP+ cells. Acknowledgments We are indebted to Drs. G. Keller, A.G. Elefanty, S. Nishikawa, A. Smith, S.J. Morrison, J. Rossant, and C. Wright for the T-GFP, Mixl1-GFP, Gsc-GFP, Sox2-LacZ, Sox17-GFP, Flk1-LacZ and Pdx1-LacZ reporter ES cell lines and anti-Pdx1 antibody. We thank Ragna Jørgensen, Søren Refsgaard Lindskog, Gurmeet Kaur Singh, Heidi Ingemann Jensen, and Rodrigo Garcia for excellent technical assistance. This work was made possible by funding from the Juvenile Diabetes Research Foundation, EU Integrated Project No. 512145, PS; NIDDK U19-DK04-017, PS; and the Danish Stem Cell Research Doctoral School (PS and PM-H). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.ydbio.2009.03.026. References Ahnfelt-Ronne, J., Jorgensen, M.C., Hald, J., Madsen, O.D., Serup, P., Hecksher-Sorensen, J., 2007. An improved method for 3D reconstruction of protein expression patterns in intact mouse and chicken embryos and organs. J. Histochem. Cytochem. 55, 925–930. Ang, S.L., Wierda, A., Wong, D., Stevens, K.A., Cascio, S., Rossant, J., Zaret, K.S., 1993. The formation and maintenance of the definitive endoderm lineage in the mouse: involvement of HNF3/forkhead proteins. Development 119, 1301–1315.

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Supplementary data, Paper I ‘A late requirement for Wnt and FGF signaling ’during activin-inducde formation of foregut endoderm from mouse embryonic stem cells’ Developmental Biology, 2009, 330, p. 286 – 304. Mattias Hansson, Dorthe R- Olesen, Janny M.L. Peterslund, Nina Engberg, Morten Kahn, Maria Winzi, Tino Klein, Poul Maddox-Hyttel and Palle Serup

Figure S1

Figure S1: BMP4 and Activin induction of GFP expression in TGfp/+, Mixl1-GscGfp/+, GscGfp/+ and Sox17Gfp/+ cells. Primary flow cytometry histograms of GFP expression in TGfp/+ and Mixl1Gfp/+ cells cultured for 3 days with or without 10 ng/ml BMP4 and GscGfp/+ and Sox17Gfp/+ cells cultured for 5 days in the presence or absence of 100 ng/ml activin

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Figure S2

Figure S2: A subpopulation of T+ cells expresses Mixl1, whereas Gsc+ cells express no or low levels of T. Immunofluorescent analyses of T expression in TGfp/+ (A), Mixl1Gfp/+ (B) or GscGfp/+ cells (C) grown in adherent culture for 3 or 5 days in the presence of the indicated growth factors. The arrows in (C) indicate areas with high T expression and low/no Gsc (GFP) expression. (D) Immunofluorescent analyses of Foxa2 and E-cad expression in TGfp/+ (a–d), Mixl1Gfp/+ (e–h), or GscGfp/+ cells (i–l) after activin treatment.

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Figure S3

Figure S3: ES cell progeny treated with activin express markers of definitive endoderm and not visceral endoderm. (A) Flow cytometric analysis of CXCR4 expression in Mixl1Gfp/+ cells grown for 5 days with or without 100 ng/ml activin. Mouse embryonic day (E)11 head tissue was used as control. (B, C) RT-PCR analysis for Sox7 expression in TGfp/+ cells (B) and Sox17, Pyy, Sox7 and Tdh expression in Flk1LacZ/+ cells (C). Cells were cultured in 0, 3 or 100 ng/ml activin, 10 ng/ml BMP4, 100 ng/ml Wnt3a or 100 ng/ml activin + 100 ng/ml Wnt3a for 5 days as indicated. E7 mouse embryo cDNA was used as positive control and expression of Tbp and G6pd was analyzed as reference genes.

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Figure S4

Figure S4: Expression of TGF-β, Wnt and FGF signaling components during directed differentiation of ES cells. RT-PCR analysis of TGfp/+ cells cultured for 3 or 5 days in medium containing growth factors and/or inhibitor, as indicated. The expression of components of the nodal/activin, BMP, Wnt and FGF signaling pathway was analyzed. The expression of Tbp was analyzed as reference gene.

Opposite, Figure S5: Wnt signaling is inhibited by addition of Dkk1 to activin-stimulated SuTOP-CFP cells cultured for 3 and 5 days. (A) Chimeric E10.5 embryo recovered after blastocyst injection of SuTOP-CFP ES cells. Note CFP expression at sites known to harbor active Wnt signaling at this stage including the otic vesicle (white arrow), optic vesicle (white arrowhead), tail bud (red arrow), and somites and dorsal neural tube (red arrowhead). (B) SuTOP-CFP cells were cultured in the presence of 100 ng/ml activin for 3 days and supplemented with 100 ng/ml Wnt3a or 320 ng/ml Dkk1 for a variable number of days as indicated. Scale bar is 80 µm. (C) SuTOP-CFP cells were cultured in the presence of 100 ng/ml activin for 5 days and supplemented with 100 ng/ml Wnt3a or 320 ng/ml Dkk1 for a variable number of days as indicated in the figure. “–” indicates treatment with activin alone. Scale bar is 200 µm.

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Figure S5

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Figure S6

Figure S6: qRT-PCR analyses of activin-stimulated ES cells after inhibition of Wnt signaling. GscGfp/+ cells were cultured in 100 ng/ml activin for 5 days and (A) transfected with β-catenin siRNA as indicated or (B) treated with Dkk1 as indicated. At the end of day 5 of the culture period RNA was prepared and qRT-PCR was performed to measure the relative abundance of Lhx1 and Chrd message.

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4. Pxd1 induction Background The work shown in Figure 12 of paper I was performed by Nina Engberg and me in collaboration. We worked together with other members of the lab on patterning the naïve DE induced by high concentrations of activin. In this chapter, I will show additional data on this patterning performed by myself, and refer to data performed by other members of the lab. The motivation behind this project departs from work by Mattias Hansson and Dorthe R. Olesen showing that they could differentiate mES cells to DE after 5 days of culture in 100 ng/ml activin, referred to as ‘Step 1’ of the differentiation protocol. This DE was believed to be of a naïve type, as it did not stain positive for regional markers of the gut tube, such as SOX2, PDX1, NKX6.1 or CDX2 (Figure 4-1). We proposed that the DE could be patterned into cells resembling posterior foregut endoderm, the region from which the pancreatic outgrowth occurs, when presented to the correct inducing signal.

Figure 4-1: A high concentration of activin in Step 1 induces naïve DE. Flk1-LacZ cells were differentiated for 5 days in 100 ng/ml activin and both differentiated cells mouse and tissue was stained for whole endoderm and regional markers: FOXA2, SOX2, PDX1, NKX6.1, CDX2 and the nuclear stain DAPI. Mattias Hansson, unpublished data.

To pattern this naïve DE into posterior foregut endoderm, referred to as ‘Step 2’ of the differentiation protocol, we relied on data from mouse and chicken development. We chose specific genes regionally expressed along the anterior-posterior (A-P) axis of the gut tube as markers of cell fate. These were Sox2, expressed anterior to the pancreatic region; Pdx1, expressed in the duodenum and pancreatic regions; Nkx6.1, expressed in the pancreatic

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epithelium; Cdx2, expressed posterior to the pancreatic region. Foxa2, expressed throughout the gut tube, was used as a marker for DE-derived cell types (see also Figure 1-3; (Grapin-Botton and Melton 2000; Jorgensen et al. 2007)). In the embryo, endoderm cells are patterned by signalling events within the endoderm or from the mesoderm. Activin has an anteriorising effect on both endoderm formation and patterning whereas WNT3a has a posteriorising effect on PS-formation and cell specification during gastrulation (Grapin-Botton and Melton 2000). Dessimoz and co-workers showed that in chicken embryos, FGF4 also has a posteriorising effect starting during gastrulation and persisting through the early somite stages (Dessimoz et al. 2006). FGF-signalling is necessary for restricting anterior expressed genes and for establishing midgut gene expression. Retinoic acid (RA) is implicated in the development of tissues in all three germ layers. In mES cell differentiation, it is often used to induce cells of a neural type (Okada et al. 2004). In ES cell aggregates, RA has been shown to induce Pdx1-expressing cells (Micallef et al. 2005), and in endoderm patterning towards pancreatic cell types it is used in both mouse and human ES cell protocols (D'Amour et al. 2006; Micallef et al. 2007; Borowiak and Melton 2009).

Materials and Methods We cultured and analysed cells as described in Hansson et al., 2009. Below are additions to the methods and materials described. Cell culture and differentiation We used the following mES cell lines: Sox2-LacZ (Li et al. 1998) and Pdx1-GFP (Holland et al. 2006). We added KAAD-cyclopamine (Toronto Research Chemicals) and FGF10 (R&D Systems) to the differentiation medium. RT-PCR RT-PCR was performed using the primer sequences: Pdx1 F_AAATTGAAACAAGTGCAGGT, R_GACAGTTCTCCACTGCTCTC; GAPDH F_ CGGTGCTGAGTATGTCGTGGA, R_ GGCAGAA GGGGCGGAGATGA.

Results We applied a 2-step protocol for the patterning of our DE: ‘Step 1’ – DE induction: 5 days in 30 – 100 ng/ ml activin, seeding density of 2.000 cells/ cm2 ‘Step 2’ – Pdx1 induction: 3 – 5 days in a combination of growth factors and inhibitors

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Figure 4-2: WNT3a and FGF4 in Step 2 induce anterior gut tube cells. Sox2-LacZ cells differentiated for 5 days in 100 ng/ml activin (Step 1) and 3 days in 5 ng/ml WNT3a + 10 ng/ml FGF4 (Step 2) were stained for β– Galactosidase and imaged under 10× objective.

Figure 4-3: Increasing the FGF4-concentration does not result in posterior foregut-type cells. Flk1-LacZ cells differentiated for 5 days in 100 ng/ml activin (Step 1) and 3 days in 25 ng/ml WNT3a ± 10 or 100 ng/ml FGF4 (Step 2) were stained for FOXA2, SOX2, and PDX1. 20× objective.

FGF4 and WNT3a alone induce cells resembling anterior or posterior gut tube The first attempts to pattern our DE into cells of a pancreatic type was done by addition of 0, 10 or 100 ng/ml FGF4 and/ or 0, 5 or 25 ng/ml WNT3a to the basic medium for 3 days.

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Figure 4-4: A high FGF4-concentration induces CDX2+ posterior gut tube. Flk1-LacZ cells differentiated for 5 days in 100 ng/ml activin (Step 1) and 3 days in 25 ng/ml Wnt3a + 0, 10 or 100 ng/ml FGF4 (Step 2) were stained for FOXA2 and CDX2. 20× objective, scale bar indicates 50 µm.

This resulted in numerous Sox2-LacZ+ and SOX2+ cells as seen by β-Galactosidase stain and immune-cytochemistry, respectively (Figures 4-2 and 4-3). Variation in the concentrations of FGF4 and WNT3a did not seem to influence the numbers of Sox2-LacZ+ or SOX2+ cells, as long as both factors were present. We saw vast numbers of FOXA2+ cells throughout the conditions applied (Figure 4-3). FGF4 and WNT3a did not induce PDX1+ cells under any conditions (Figure 4-3), but high concentrations of FGF4 in combination with WNT3a (5 or 25 ng/ml) induced CDX2+ clusters in the culture (Figure 4-4). To be sure that we did not miss a weak expression of Pdx1 in the samples, we performed RT-PCR and saw only the housekeeping gene Glyceraldehyde 3-phosphate dehydrogenase (Gapdh) expressed (Figure 4-5). Thus, we concluded that FGF4 and WNT3a either alone or in combination were not enough to drive patterning of the DE into Pdx1-expressing posterior foregut cell types. We saw vast numbers of Sox2-LacZ+ and SOX2+ cells, indicative of an anterior DE-type. CDX2 was induced by 100 ng/ml FGF4, suggesting that this concentration is too high when aiming at PDX1-induction.

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Figure 4-5: Pdx1 cannot be detected by RT-PCR. RT-PCR for Pdx1 on E18,5 mouse pancreas and β-TC3, i.e. positive controls and Flk1-LacZ cells differentiated for 5 days in 100 ng/ml activin (Step 1) and 3 days in 25 ng/ml WNT3a ± 10 or 100 ng/ml FGF4 (Step 2). Gapdh is used as a house-keeping gene control. Loading in the individual wells is described next to the agarose gel images.

RA in combination with WNT3a and FGF4 induces posterior foregut cells As shown in paper I, figure 12, we could induce PDX1-expression by addition of 0.1 µM RA and intermediate levels of WNT3a (5 ng/ ml) and FGF4 (10 ng/ml) to Step 2 of the differentiation protocol. Under these conditions, we saw a vast number of FOXA2+/SOX2+ cells and no CDX2-expressing cells. This indicates that RA, in combination with WNT3a and FGF4, induces cells of an anterior-intermediate gut fate expressing FOXA2 and SOX2 or PDX1, but no CDX2. Prolonged differentiation does not increase+ Pdx1-induction To be able to quantify the numbers of PDX1 cells, we used a Pdx1-GFP cell line (Holland et al. 2006) and applied our 2-step protocol to this. We included KAAD-cyclopamine (cyclopamine; an inhibitor of SHH-signalling) in this protocol, as inhibition of SHH has proven crucial for Pdx1-induction in the developing gut tube (Hebrok et al. 1998). We generally saw induction of 2-4% Pdx1-GFP+ cells when applying our standard protocol of 0.1 µM RA, 5 ng/ml WNT3a and 10 ng/ml FGF4 (data not shown). This was not satisfactory for continued differentiation towards posterior foregut endoderm, and we speculated whether our DE was still to immature to be patterned after 5 days of DE-induction. Thus, we introduced an extra step between endoderm induction and patterning: two days of culture with or without patterning factors FGF10 and cyclopamine followed by combinations of RA, FGF10 and cyclopamine. In general, we saw a very low induction of Pdx1-GFP+ cells in all conditions, ranging from 0,5-4% (Figure 4-6). There was a tendency for more Pdx1-GFP+ cells with an increase in factors applied, the combination of all three being the most potent. Also, the cells grown in FGF10 and cyclopamine in step 2 showed higher numbers of Pdx1-GFP+ cells in general. There seems to be little endogenous SHH-signalling in the DE cell population, as inhibition thereof did not improve our protocol. However, with a total amount of 2-4% Pdx1GFP+ cells, this protocol needs further optimization.

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Figure 4-6: A 3-step protocol does not enhance the numbers of Pdx1-GFP+ cells. Pdx1-GFP cells differentiated for 5 days in 30 ng/ml activin (Step 1), 2 days in Step 2A-media and 3 days in Step 2B-media (see schematic for details). Concentrations used: 0,1 µM RA; 25 ng/ml FGF10; 0,25 µM KAAD-Cyclopamine (Cyc). n = 3 ± S.E.M. is shown.

Figure 4-7: Addition of the posteriorizing factor BMP4 in Step 1 does not increase Pdx1-GFP induction. Pdx1GFP cells differentiated for 5 days in 30 ng/ml activin (Step 1) and 3 days in Step 2-media (see schematic for details). Concentrations used: 0,1 or 1 ng/ml BMP4; 0,1 µM RA; 25 ng/ml FGF7; 1 µM SB431542; 50 mg/ml Noggin. n = 3 ± S.E.M. is shown.

Posteriorising DE-induction does not increase the posterior foregut cell population We decided to look into whether the DE we generated by a high concentration of activin could be posteriorized already in Step 1, thus leading to higher numbers of posterior foregut endoderm in Step 2. Based on DE-induction protocols used for mES or hES cells, we added BMP4 at different concentrations in Step 1 and inhibited this signalling-pathway in Step 2 (Candy H.-H. Cho, personal communication; (Morrison et al. 2008; Touboul et al. 2009)). In Step 1, we found that BMP4-concentrations of 0,1 ng/ml on days 1-2 or 1-5, or 1 ng/ml on days 1-2 had no inhibitory effect on the percentage of Sox17-GFP+ cells formed, whereas higher BMP4-concentrations did (Maria Winzi, unpublished data). Next, we used these concentrations in Step 1 and added FGF7 and RA ± the BMP4-inhibitor noggin and the ALK4/5/7 inhibitor SB431542. We saw a tendency for increased numbers of Pdx1-GFP+ cells when adding the low concentration of BMP4 for all 5 days or the high concentration for 2 days (Figure 4-7). However, the percentage of Pdx1-GFP+ cells was not significantly higher than when inducing

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DE in medium containing activin only. We therefore conclude that addition of BMP4 has no posteriorising effect on our DE in this system.

Discussion Our protocol for the generation of posterior foregut from an activin-induced DE cell population is successful as a ‘proof of principle’. We show that addition of posteriorising factors such as RA and FGF induce PDX1-expression in further differentiation of apparently naïve DE cells. The protocol is reproducible between E14, Pdx1-LacZ and Pdx1-GFP cell lines, in which we obtain 2-4% PDX1+ cells on average. However, attempts to increase the efficiency of Pdx1induction proved difficult and the protocol is therefore not satisfactory for further differentiation into pancreatic endoderm. It seems that there is a fundamental problem in our protocol setup, as we have had little success in improving the number of PDX1+ cells. One source of problems could be connected to our culture conditions. We use B27 which contains glucocorticoids that have been shown to inhibit Pdx1-expression (Tanimizu et al. 2004). Another issue is timing; we see the highest induction of Pdx1-GFP+ cells after 3 days of culture in Step 2 media, and introducing an extra 2 days of differentiation does not increase the numbers of Pdx1-GFP+ cells. However, 3 days seem to be a rather short induction period compared to other protocols. D’Amour and co-workers grew hES cell-derived DE for 4-8 days to differentiate them to posterior foregut (D'Amour et al. 2006). The rationales on which we have built our inducing factor-combinations seem reasonable, as studies performed in hES cells using FGF4 and RA showed an induction of 32% PDX1+ cells from DE-cultures (Johannesson et al. 2009). Also, intermediate concentrations of FGF2 were shown to induce pancreatic foregut specification, whereas higher concentrations induced a more posterior gut type (Ameri et al. 2010). In conclusion, we show that by addition of RA and FGF we can successfully induce PDX1+ cells, albeit in low numbers. This protocol lays the ground for further optimization, although this may prove difficult in the current culture conditions.

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5. Paper II FGFR(IIIc)-activation induces mesendoderm but is dispensable for definitive endoderm formation in mouse embryonic stem cells Manuscript to be submitted. Janny Marie L. Peterslund and Palle Serup* Department of Stem Cell Biology, Hagedorn Research Institute, Niels Steensens Vej 6, DK2820 Gentofte, Denmark. * Corresponding author. Author contributions Janny Marie L. Peterslund performed the experiments, analyzed and interpreted the data, made the figures and wrote the paper draft. Palle Serup conceived ideas, commented on the paper and was the principal investigator and supervisor on the research. Both authors designed the research and discussed the results. For co-authorship declaration, see Appendix B.

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FGFR(IIIc)-activation induces mesendoderm but is dispensable for definitive endoderm formation in mouse embryonic stem cells

Janny Marie L. Peterslund and Palle Serup* Department of Stem Cell Biology, Hagedorn Research Institute, Niels Steensens Vej 6, DK2820 Gentofte, Denmark. Running title: FGF(R)s and mesendoderm differentiation in mESCs Keywords: mouse embryonic stem cell, fibroblast growth factor (FGF), FGF receptors, mesendoderm, definitive endoderm, FGF4–/– cell line

*Corresponding author: Dr. Palle Serup Hagedorn Research Institute Niels Steensens Vej 6 DK-2820 Gentofte Denmark E-mail: [email protected] Phone: +45 44439822 Fax: +45 44438000

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Abstract Progress in embryonic stem (ES) cell research over the last decade has outlined a possible cell based intervention strategy in diabetes based on the hierarchical differentiation of embryonic stem cells into insulin producing beta cells. The definitive endoderm (DE) cell type constitutes an essential milestone in this differentiation pathway and fibroblast growth factor (FGF)signaling drives the formation of DE in the mouse ES cell model system. More specifically, it has been shown that FGF4 is crucial for cells to leave the pluripotent state and differentiate to ectoderm and mesoderm cell lineages. The FGF system counts several receptor isoforms with a possible functional redundant role in conducting the differentiation signal. Here, we investigate the spatio-temporal dynamics of FGF receptor (FGFR) distribution in the forming DE and find that FGFR(III)c-isoforms are highly represented in the whole culture, whereas FGFR2(III)b and FGFR4 are found in the DE fraction, specifically. The FGFR(III)c isoform-activating FGFs induce mesendoderm markers T and Gsc, but reduce the DE marker Sox17 whereas FGFs activating FGF(III)b-isoforms have no effect on either cell type. Notably, FGFR(III)c isoformactivating FGFs exhibit strong mitogenic effects on ES cells early in the differentiation period where FGFs activating FGFR(III)b-isoforms have only a moderate mitogenic potential confined to the late differentiation period. Interestingly, when applying our DE induction protocol onto an FGF4–/– cell line, we find that cells readily differentiate into endoderm cells without ectopic administration of FGF4. By antibody staining and qPCR analyses for definitive and visceral endoderm markers, we show that this endoderm is definitive rather than visceral. We conclude that FGFR(III)c-isoform activation selectively drives the differentiation of mES cells towards mesendoderm and that FGF4 is dispensable for the final differentiation step into DE.

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Introduction Based on knowledge obtained in developmental biology, mouse embryonic stem (mES) cells can be directed towards differentiation into specific germ layers and more mature tissues. Such a differentiation into glucose-responsive β cell-like insulin-secreting cells, serves in theory as a cure for type I diabetes mellitus (McCall et al. 2010). For this purpose, the first step is to generate definitive endoderm (DE) with the potential to further differentiate into cells resembling the primitive gut tube (reviewed by (Van Hoof et al. 2009)). Understanding the role of each component used in this directed differentiation is crucial for obtaining the optimal progenitor cell population in each step. In the late blastocyst stage of the developing mouse embryo (E4.5), the inner cell mass (ICM) is divided into the epiblast and the primitive, later visceral endoderm. The visceral endoderm (VE) is involved in the asymmetric anterior-posterior patterning of the epiblast, resulting in the onset of gastrulation (reviewed by (Rossant 2004)). In the gastrulating mouse embryo, epiblast cells migrate through the primitive streak (PS), and in this process become determined towards either mesoderm or DE germ layers (Lawson et al. 1991; Tam et al. 1993; Carey et al. 1995). The transforming growth factor-β family member nodal, an activator of SMAD2/3 signalling, is the main initiator of epiblast patterning and PS formation (Conlon et al. 1994; Waldrip et al. 1998). At high levels, nodal induces anterior PS structures and DE and at low doses it induces more posterior streak fates (Ben-Haim et al. 2006). At the posterior-most end of the PS, bone morphogenetic protein 4 (BMP4) is produced by the extra-embryonic ectoderm and establishes a signal gradient. BMP4 is critical for formation of the PS and induces mesoderm formation (reviewed by (Gadue et al. 2005)). It is currently accepted that cells in the mesoderm and endoderm tissues arise from a common progenitor cell population, the mesendoderm (Lawson et al. 1991; Kinder et al. 2001). The PS marker Brachyury (T) is expressed in the nascent and migrating mesoderm of the primitive streak during gastrulation in the mouse embryo (Kispert and Herrmann 1994). Goosecoid (Gsc) is located in the progressing primitive streak at E6.5, and later localizes to the anterior streak, from which the DE arises (Blum et al. 1992). It is induced by high concentrations of activin in animal cap explants from Xenopus and in mES cells, it is used as a marker for the mesendoderm cell population (Kubo et al. 2004; Gadue et al. 2006). Sry-related HMG box gene 17 (Sox17) is an early marker specifically expressed in the definitive endoderm of the gastrula, and later expands to the endoderm underlying the neural plate of the early-bud-stage embryo (Kanai-Azuma et al. 2002). Sox17 is also expressed in the extra-embryonic, but not in the embryonic visceral endoderm. In mES cell cultures, cells take on a mesendoderm-type fate before being committed to either the mesoderm or DE lineages (Tada et al. 2005). ActivinA (activin hereafter) is used as a surrogate for nodal as they both activate SMAD2/3 signalling by binding to the ALK4 receptor, thus functioning in the same manner (Schier 2003). In the mesendoderm population, high concentrations of nodal/ activin-signalling induce anterior streak and DE cells while BMP4 or low concentrations of nodal/ activin induce posterior streak or mesoderm (Kubo et al. 2004; Willems and Leyns 2008; Hansson et al. 2009). The PS genes T, Mix-like 1 (Mixl1) and Gsc are expressed in this population in response to increasing concentrations of activin. High activinlevels further induce the DE markers Sox17, E-cadherin and Forkhead box A2 (Foxa2). BMP4 induces T, Mixl1 and the mesodermal marker Fetal like kinase 1 (Flk1; VEGFR2/ Kdr; (Gadue et al. 2005)). During mES cell differentiation, T-expressing cells give rise to endoderm and mesoderm derivatives (Kubo et al. 2004) and we have previously shown that a T-GFP reporter cell line (TGfp/ +; (Fehling et al. 2003)) can be activated by BMP4 and a low concentration of activin (Hansson et al. 2009). For the differentiation of mesendoderm and DE to occur properly in mES cells, fibroblast growth factor (FGF)-signalling is required (Funa et al. 2008; Morrison et al. 2008; Willems and Leyns 2008; Hansson et al. 2009). The FGF family of proteins consists of 22 members named FGF1-23 (FGF15 is the mouse ortholog of human FGF19). They activate one or more of four receptor tyrosine kinases, the FGF receptors (FGFRs)1-4. FGFRs1-3 have two secreted splice

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variants in their Ig-like domain III, the FGFR(III)b or FGFR(III)c isoforms (hereafter FGFRb or FGFRc, respectively; (Ornitz and Itoh 2001; Itoh and Ornitz 2004)). FGFs are involved in many functions such as germ layer formation, limb development, cell proliferation and cell migration in the developing embryo (Ornitz and Itoh 2001). In early mouse development, FGFsignalling is necessary for the migration of epiblast cells through the PS (Ciruna et al. 1997; Guo and Li 2007). The loss of FGF4 is lethal at E4-5, due to the inability of epiblast cells to undergo epithelial-to-mesenchymal transition and migrate through the PS (Feldman et al. 1995). FGFR1–/– mice also die at gastrulation and both FGF4 and FGFR1 are expressed in the ICM and PS (Deng et al. 1994; Yamaguchi et al. 1994). FGF4 is expressed in pluripotent mES cells and has been shown to be necessary for differentiation into ectoderm and mesoderm lineages, suggesting a crucial role of FGF4 in the initiation of differentiation (Kunath et al. 2007). However, Wilder et al. showed that FGF4–/– cells could differentiate in vitro, albeit at a low frequency, and gave rise to tumours consisting of a wide range of differentiated cell types in vivo (Wilder et al. 1997). In this study, we expand on our previous finding that active FGF-signalling is necessary for DE formation (Hansson et al. 2009), and investigate the effects of different FGFR-isoforms on mesendoderm and DE differentiation. We first analyse the expression patterns of FGFRisoforms in the forming DE and find that FGFRc-isoforms are up-regulated in the bulk culture whereas FGFR2b and 4 are up-regulated specifically in the DE-fraction. By means of reporter cell lines and antibody staining, we find that FGFs activating primarily FGFRc-isoforms induce PS and mesendoderm markers T and Gsc but reduce the DE marker Sox17. FGFs activating FGFRb-isoforms have no effect on either cell type. The FGFRc isoform-activating FGFs show the highest mitogenic effects early in the differentiation period, and suggestively speeds up differentiation, as proliferation rates in the presence of these FGFs are reduced later in the culture period. Remarkably, an ES cell line carrying a knockout for one such FGFRc isoformactivating FGF, FGF4–/– was able to differentiate to endoderm cells at levels comparable to wt and FGF4+/– situations. The absence of FGF4 gave rise to DE differentiation, and although a few cells stained positive for the VE marker Sry-related HMG box 7 (SOX7), qPCR analyses confirmed the DE fate of the culture. Thus, we conclude that FGFRc-isoforms specify the mesendoderm but not DE cell population and that FGF4-signalling is dispensable for induction of DE cells.

Materials and Methods Cell culture and differentiation of mESCs We used the following mouse ES cell lines: E14 (Hooper et al. 1987), T-GFP (Fehling et al. 2003), Gsc-GFP (Tada et al. 2005), Sox17-GFP (Kim et al. 2007), FGF4+/– and FGF4–/– (Wilder et al. 1997). Cells were grown as previously described (Ying et al. 2003a; Hansson et al. 2009) on cell culture plastic ware (Nunc) coated with 0,1% gelatine (Sigma), using 0,05% Trypsin-EDTA (Invitrogen) for dissociation of cells during passage. Trypsin was inactivated by N2B27 medium: KO-DMEM supplemented with N2, B27, 0.1 mM non-essential amino acids, 2 mM L-glutamine, Penicillin/Streptomycin (all from Invitrogen), 0.1 mM 2-mercaptoethanol (Sigma-Aldrich). Cells were grown for at least 3 passages before onset of differentiation. For differentiation purposes, cells were dissociated into single cells and seeded at 2.000 cells/ cm2 in N2B27 medium containing one or more of the following growth factors: BMP4 (10 ng/ml), activinA (1 or 30 ng/ml; both from R&D Systems), FGF1 (100 ng/ml; Chemicon International), FGF2 (100 ng/ml; Invitrogen), FGF4, FGF5, FGF6, FGF7, FGF8b, FGF8c, FGF8e, FGF9, FGF10, FGF16 (5 or 100 ng/ml; all from R&D Systems). Media containing FGFs were supplemented with 10 µg/ml heparan sulfate (Sigma-Aldrich). Flow cytometry For GFP-analysis of reporter cell lines, live cells were dissociated into single cells by 0,05% Trypsin-EDTA (Invitrogen) and analysed by FACS Calibur flow cytometer (BD Biosciences). 61

For analysis of cells stained with antibodies, cells were fixed in LILLY’s fixative (Bie & Berntsen), and resuspended in 0,1% BSA in PBS. Cells were stained in 0,1% BSA in PBS for 2 hrs at 4°C with Flk1-PE (BD Pharmigen, # 555308) and EpCAM-PE-Cy7 (eBioscience, # 255791-80). Or cells were permeabilised in dilution buffer (0,3% Triton X-100 + 0,1% BSA in PBS), unspecific binding sites were blocked by 10% Normal Donkey Serum (Jackson Immunoresearch Laboratories) for 30 minutes at RT and stained for Brachyury (R&D Systems, # AF2085) for 2 hours at RT in dilution buffer, followed by a Cy3-conjugated secondary antibody (Jackson Immunoresearch Laboratories, # 705-165-147) for 1 hour at RT. Cells were analysed by FACS Aria flow cytometer (BD Biosciences). Cell sorting Cells were dissociated by 0,05% Trypsin-EDTA (Invitrogen), washed and resuspended in N2B27 medium before sorting by FACS Aria flow cytometer (BD Biosciences). Immunofluorescent staining Cells were grown in 9 cm2 slide flasks (Nunc) coated with 0,1% gelatine (Sigma) and fixed in LILLY’s fixative (Bie & Berntsen), permeabilised in dilution buffer (see above) and blocked for 30 minutes at RT in 10% Normal Donkey Serum (Jackson Immunoresearch Laboratories) in dilution buffer. They were stained ON at 4°C with primary antibodies: mouse anti-Oct3/4 (C-10), goat anti-Foxa2 (both Santa Cruz Biotechnology), goat anti-Brachyury (R&D Systems), rat anti-E-cadherin (Zymed/Invitrogen), goat anti-Sox17 (R&D Systems), and 1 hr with Cy2-, Cy3- or Cy5-conjugated species-specific secondary antibodies (Jackson ImmunoResearch Laboratories) and 4′,6-diamidino-2-phenylindole (DAPI, MP Biomedicals). Slides were mounted in Fluorescent mounting medium (KPL). Negative controls, where the primary antibodies were omitted, were included for all stainings and showed no unspecific staining of the secondary antibodies (data not shown). The slides were analyzed using an LSM 510 META laser scanning microscope (Carl Zeiss). qPCR Cells were harvested in Lysis solution (Invitek), supplemented with 10 mM dithiothreitol (DTT). Total RNA was isolated using the Invisorb Spin RNA kit (Invitek) with DNAse treatment (Promega) following the manufacturer’s protocol. cDNA was prepared from 250 ng RNA using MMLV Reverse Transcriptase (Invitrogen) with random oligos or oligo(dT)12-18 primers (both Invitrogen). qPCR was performed using the standard SYBR® Green program with dissociation curve on the Mx3005P (Stratagene). PCR reactions were run in duplicates using 10 µl Brilliant® SYBR® Green qPCR Master Mix (Stratagene), 1 µl cDNA, 1 µl 20 µM primer-mix and 8 µl dH2O. Quantified values for each gene were normalized against the housekeeping gene TATAbinding protein (TBP). Statistical analyses were performed using Student’s two-tailed, paired ttest. Primer sequences are: FGFR1c F_CCGTATGTCCAGATCCTGAAGA, R_GATAGAGTTACCCGCCAAGCA; FGFR2c F_GCCCTACCTCAAGGTTCTGAAAG R_GATAGAATTACCCGCCAAGCA; FGFR3c F_CCCTACGTCACTGTACTCAAGACTG R_GTGACATTGTGCAAGGACAGAAC; FGFR4 F_CGACGGTTTCCCCTACGTACA R_TGCCCGCCAGACAGGTATAC (all from (Woei Ng et al. 2007); FGFR1b F_CTTGACGTCGTGGAACGATCT, R_CACGCAGACTGGTTAGCTTCAC (Nakayama et al. 2007); FGFR2b F_AACGGGAAGGAGTTTAAGCAG, R_GGAGCTATTTATCCCCGAGTG (Yamanaka et al. 2000); Sox17 F_GGAGGGTCACCACTGCTTTA, R_TCAGATGTCTGGAGGTGCTG; Cxcr4 F_AGGTACATCTGTGACCGCCTTT, R_ AGACCCACCATTATATGCTGGAA (Kim et al. 2008); Sox7 F_GGCAGTGCAGAACCCGGACC, R_TGCAGAGGCGCTTGCCTTGT; Tdh F_ CCTGGAGGAGGAACAACTGACTA, R_ ACTCGAATGTGCCGTTCTTTG (Wang et al. 2009); TBP F_TCTGAGAGCTCTGGAATTGT, R_GAAGTGCAATGGTCTTTAGG.

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Cell count and proliferation assay Cells were fixed in LILLY’s fixative (Bie & Berntsen) and counted in a NucleoCassette read by the NucleoCounter (ChemoMetec A/S) according to the manufacturer’s protocol for counting non-living cells. As a proliferation assay, EdU-incorporation by the Click-iT® EdU HCS Assay (Invitrogen) was used (Salic and Mitchison 2008). Cells were incubated for 15 min in their respective media containing 10 µM EdU. Cells were washed, fixed and stained by the Click-iT reaction cocktail according to the manufacturer’s protocol, using the Alexa Fluor 488-conjugated antibody to detect incorporation. Stained cells were quantified using the FACS Aria flow cytometer (BD Biosciences). Statistics Mean % of the cells of interest ± standard deviation or standard error of the mean (S.D. or S.E.M.) was calculated and statistical analyses by Student’s paired, two-tailed t-test or Ratio ttest were performed.

Results Expression of FGF receptor-isotypes during definitive endoderm formation Recently, we and others have shown that fibroblast growth factor (FGF)-signalling in mouse embryonic stem (mES) cell cultures is necessary for the differentiation of definitive endoderm (DE; (Funa et al. 2008; Morrison et al. 2008; Willems and Leyns 2008; Hansson et al. 2009)). These studies are based on a general requirement for active FGF-signalling during early mouse development where FGF3, 4, 5, 8b and FGFR1 are expressed in the epiblast to post-gastrulation embryo (Wilkinson et al. 1988; Haub and Goldfarb 1991; Hebert et al. 1991; Niswander and Martin 1992; Ciruna et al. 1997; Guo and Li 2007). To elaborate on the observed dependence on FGF-signalling during DE formation, we made a thorough investigation of the expression of isoforms of FGFRs during the 5-day differentiation period by quantitative RT-PCR (qPCR). We used a Sox17Gfp/ + reporter cell line (Kim et al. 2007) and sorted cells into Sox17-GFPHi and Sox17-GFPLo fractions, in order to isolate RNA from the forming DE and the non-DE populations of cells, respectively (Figure 1A). In general, FGFR1c was expressed at high levels, FGFR2b and 2c at intermediate levels and FGFR1b, 3c and 4 at low levels. During the 5-day differentiation period, FGFR2b and 4 were up-regulated in the unsorted and Sox17-GFPHi fractions while their expression was either unchanged or down-regulated in the Sox17-GFPLo fractions (Figure 1B). FGFR1b was down-regulated in both the unsorted and Sox17-GFPLo fractions upon initiation of differentiation, although not significantly different from the undifferentiated culture. These data confirm findings by other groups, showing that FGFR2b and 4 are expressed in endodermal epithelia such as the definitive endoderm (Stark et al. 1991; Orr-Urtreger et al. 1993; Elghazi et al. 2002). FGFR1c was upregulated in the Sox17-GFPLo fraction alone, peaking on day 5, whereas FGFR2c and 3c were up-regulated in both Sox17GFPLo and Sox17-GFPHi fractions, upon differentiation by activin (Figure 1B). In summary, FGFRc-isoforms are highly up-regulated throughout the cell culture or in the Sox17-GFPLo fraction alone, whereas FGFR2b and 4 are up-regulated in the Sox17-GFPHi fraction specifically, suggesting a role for especially FGFRc isoform-activation during DE formation in mES cells. FGFs activating specific sub-populations of FGFRs differentially activate PS and DE markers Since FGFs activate specific FGFR-isofoms, we speculated that certain FGFs were likely to have a more potent effect on expression of PS and DE markers in a culture system aimed at inducing such cell types. We chose to focus on FGFs that are described to have a function during gastrulation and in the development of the DE and based on which FGFRs they activate

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they were divided into three categories (Figure 2A; (Ornitz et al. 1996; Bottcher and Niehrs 2005; Zhang et al. 2006)). FGF1, 2 and 9 activate a mixed population of both FGFRb- and FGFRc-isoforms, with a preference for the latter; FGF7 and 10 activate FGFRb-isoforms only; and FGF4, 5, 6, 8b, 8c, 8e and 16 activate one or more FGFRc-isoforms and/ or FGFR4 (MacArthur et al. 1995; Ornitz et al. 1996; Olsen et al. 2006; Zhang et al. 2006; Mason 2007). FGFR4 is grouped with the FGFRc-type of receptors, based on the fact that it structurally resembles this group of FGFRs (Vainikka et al. 1992). Importantly, most FGFs activating FGFRc-isoforms also activate the FGFR4 (Mason 2007), making it also functionally an FGFRc-type. To evaluate the effect of the different FGFs in mesendodermal differentiation, we monitored at the expression of PS and DE markers by means of reporter cell lines on days 3 and 5. Accordingly, we induced mesoderm by 10 ng/ml BMP4 and added different FGFs to evaluate their effect on the PS marker TGfp/ + cell line expression. FGF1, 2, 4, 6 and 9, binding a mixed population of FGFRs or FGFRc-isoforms only, increased the number of T-GFP+ cells on day 3 by up to 20% compared to BMP4-treatment alone, i.e. 79 – 83 ± 6 – 10% and 69 ± 6%, respectively (mean % ± S.D., n=3; Figure 2B). FGF7 and 10 had no significant effect on TGFP induction, nor did FGF8b, 8c, 8e or 16, but FGF5 slightly repressed T-induction (Figure 2B). Looking at the same marker in a posterior streak/ mesoderm-inducing protocol, using 1 ng/ml activin, we saw that FGF4 and 6 show a 31 – 42% increase in T-GFP induction on day 3 (Figure 2C), while FGF5 and 10 show a smaller increase. FGF1, 2, 4, 6 and 9 induced numbers of T-GFP+ cells by up to 34% on day 5. FGF7, 8b, 8c, 8e and 16 showed no effect on the numbers of T-GFP+ cells on either day 3 or 5. Thus, the largest effect was seen when adding FGFs binding FGFRc-isoforms or a mixed population of FGFRs, mediating an increase in TGFP+ cells in general and on day 5 in particular. The reason for this pronounced effect can be through either a delay in the response by some cells (in media containing BMP4 or activin only, it peaks on day 3), or maintaining the T-expression for a time period extending beyond day 3. Next, we looked at the effect of FGFs on anterior streak/ DE-induction by 30 ng/ml activin in a mES cell line containing a GFP knock-in allele of the anterior streak marker Gsc, GscGfp/ + (Tada et al. 2005). Addition of FGF1, 2, 4, 6, 8b and 9 increased the number of Gsc-GFP+ cells by 22 – 40% (Figure 2D). Activation of FGFRb-isoforms only, by FGF7 and 10, had no effect and nor did FGF5, 8c, 8e and 16. This finding was not due to the lack of receptors, as they were present in the cell population (Figure 1B). Looking at the DE marker Sox17, we saw up to a 50% decrease of the Sox17-GFPHi fraction, from 34 ± 4% to 17 ± 3% when adding FGFs activating FGFRc-isoforms (Figure 2E). FGFs activating FGFRb-isoforms only, slightly increased the number of Sox17-GFPHi cells or had no effect (FGF7 and FGF10, respectively). In summary, FGFs binding predominantly FGFR4/FGFRc-isoforms, i.e. FGF1, 2, 4, 6, 8b and 9, promote differentiation towards a mesendoderm cell population expressing primitive and anterior streak markers. The mesendoderm population responds to FGFRc-isoform activation only Next we investigated whether the FGFs inhibiting DE formation would instead promote PS and mesoderm marker expression by analysing Sox17-GFP cells stained with antibodies against T, FLK1 and EpCAM. For analytical purposes, cell populations were divided into Sox17-GFP–/Lo and Sox17-GFPHi fractions. We limited the number of FGFs introduced in these experiments, focusing only on those showing the largest effect on PS and DE marker induction/ repression, namely FGF1, 2 and 9 (mixed FGFR-activation); FGF7 and 10 (FGFRb-activation only); and FGF4, 6 and 8b (FGFRc-activation only). As a control, we included differentiation by BMP4 and saw that these cells expressed the most T in Sox17-GFP–/Lo cells as expected (Figures 3A and B). In the Sox17-GFPHi fraction, 12% of cells were T+ on day 3 when treated with activin alone (Figure 3A). This number was increased by addition of FGF2, 4, 7 and 10, and decreased in the presence of FGF6, 8b, 9 and especially FGF1. On day 5, numbers of T+ cells were rather low and an increase in the Sox17-GFPHi/T+ cell population was seen only when adding FGF2 (Figure 3A). We also investigated the Sox17-GFP–/Lo fraction and saw that FGF7 and 10 had no

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influence on the numbers of Sox17-GFP–/Lo/ T+ cells on day 3 (Figure 3B), but the other FGFs all reduced cell numbers. On day 5, cell numbers were in general very low, showing a slight induction of Sox17-GFP–/Lo/ T+ cells when adding FGF2, 4 or 6 (Figure 3B). These data demonstrate that the FGFs activating either a mixed population of FGFRs or FGFRc-isoforms can reduce the number of T+ cells in the emerging DE and in the rest of the cell culture, which has been suggested to consist of cells of an anterior streak type, not yet determined to express DE markers (Hansson et al. 2009). Tada and co-workers showed that the mesendoderm population can be distinguished by the PS marker Gsc, and later the DE can be separated from the mesoderm by means of the DE markers SOX17, E-cadherin and FOXA2 and mesoderm markers FLK1 and platelet-derived growth factor receptors (PDGFRs) α and β (Tada et al. 2005). Trying to define whether individual FGFs would switch the mesoderm vs. endoderm balance in the differentiating culture, we investigated markers able to distinguish between these cultures namely Epithelial cell adhesion molecule (EpCAM) and FLK1. EpCAM is expressed in pluripotent mES cells and in the DE epithelium during embryonic development (Balzar et al. 1999; Sherwood et al. 2007) whereas FLK1 is expressed in all mesoderm cells leaving the embryonic posterior streak. Subsequently, FLK1 is observed in developing mesodermal cardiac crescent cells and in most extraembryonic mesoderm and at E8.5 it is expressed in splanchnic mesoderm and endothelial cells of the dorsal aorta only (Ema et al. 2006). Focusing on day 5, we stained differentiated Sox17GFP cells with antibodies for Flk1 and EpCAM. Looking at the three markers separately, we saw a decrease in Sox17-GFPHi cells in the presence of FGF1, 2, 4 and 6 in concordance with previous data (Figures 3C and 2E). There were very few FLK1+ cells when treating with activin, regardless of FGFs added, but a high induction in the BMP4-treated positive control and BSA control samples (Figure 3D). The induction of EpCAM is 87 ± 2% in the presence of activin (Figure 3E) and FGF6, 7, 8b and 9 modestly but significantly increase EpCAM expression in the range of 88 ± 3% to 92 ± 3%. We see a high amount of cells that are FLK1+ or EpCAM+ in the N2B27+BSA negative control medium. This condition gives rise to 50-75% Sox1-expressing neural progenitors at day 5 of culture ((Ying et al. 2003b); Peterslund et al., unpublished data). The high amount of FLK1+ and EpCAM+ cells in this condition is most likely due to random and/ or neural differentiation. In the Sox17-GFPHi fraction, cells turned primarily into Sox17-GFPHi/ FLK1–/ EpCAM+ cells in the presence of activin, representative of definitive endoderm (Figure 3F; numbers shown are % of Sox17-GFPHi cells). The addition of FGFs did not alter this picture, and also did not change fates of the remaining cells, these being small fractions of Sox17Hi/ FLK1+/ EpCAM+ and Sox17Hi / FLK1–/ EpCAM–. The former are likely to represent cells in a transition phase expressing both EpCAM and FLK1 and were present mainly in the BSA control sample. We only found Sox17Hi/ FLK1+/ EpCAM– cells in the BMP4-condition, thus inducing what appears to be a Sox17-expressing mesodermal cell type. We also looked at cell fates in the Sox17-GFP–/Lo fraction. Here, the vast majority of cells were still Sox17–/Lo/ FLK1–/ EpCAM+ (Figure 3G), and FGF1, 2, 8b and 9 had a positive effect on this population, elevating numbers of this population by up to 10%. In summary, FGFs activating FGFRc-isoforms slightly decrease the number of Sox17-GFP+/T+ and Sox17–/Lo/T+ cells, indicating random and/ or neural differentiation rather than PS formation at the expense of DE. In the Sox17-GFP+ population, FGFs activating FGFRcisoforms increase the number of FLK1–/EpCAM+ and decrease the number of FLK1–/EpCAM– cells possibly indicating a pool of non-mesodermal cells at intermediary steps of differentiation. FGFRc-isotype activation induces cell growth during the first 3 days of culture FGFs were originally discovered as having a mitogenic effect in fibroblast cells, and were later found to have adverse effects in embryonic development, including endoderm formation (Gospodarowicz and Moran 1975; Ornitz et al. 1996; Bottcher and Niehrs 2005). We analysed the mitogenic effect of the FGFs in wt mES cells on days 3 and 5, and found that activintreatment alone gave a 3,3 times increase in cell numbers by day 3 (from 2.000 cells/ cm2 to 6.600 cells/ cm2; Figure 4A). All FGFs improved cell growth to varying degrees, FGF1, 2, 4

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and 9 being the most effective ones (up to 20.400 cells/ cm2 or a 3 times increase compared to the activin-treated cells). By means of 5-ethynyl-2’-deoxyuridine (EdU)-incorporation (Salic and Mitchison 2008) we could quantify the proliferation of cells (see Figure S1 for gating and controls). We found that the absolute number of proliferating cells was higher at day 3 than day 5, by an approximate 4 time increase for the activin-treated cells and a 4-7 times increase for the conditions supplemented with FGFs. At day 3, the relative number of proliferating cells did not differ much from the activin-treated sample (Figure 4A) except for a small reduction when adding FGF1 or not adding any growth factors at all, i.e. the BSA control. On day 5, there was app. 80.000 cells/cm2 in samples treated with activin alone. This number increased 1,6 times to app. 130.000 cells/ cm2 when adding FGF2 (Figure 4B). This indicates that the effect of FGFs on cell growth is decreasing over time probably because more cells are differentiated at day 5. At this stage, especially FGF1 and 2 positively affect cell numbers. These have been shown also in other cell systems to have the largest mitogenic effect (Ornitz et al. 1996; Zhang et al. 2006). By EdU-incorporation, we saw a 50% higher proliferation rate in the BSA control than with cells treated with activin, showing 25% and 15% total proliferation, respectively (Figure 4B). The FGFs most frequently reduced proliferation; FGF1, 2, 4 and 6 by 40 – 50% and FGF8b, 9 and 10 showing moderate reductions. These data indicate that the main effect seen by FGF1, 2, 4, 6 and 9 on proliferation occurs prior to day 3, and that most of the cells in these cultures have left the proliferative state by day 5. FGF4 is dispensable for the formation of endoderm Of the FGFs tested, FGF4 and 6 binding FGFRc-isotypes only, increase BMP4 and activininduced expression of T and Gsc the most. FGF4 is important during gastrulation where it is responsible for the cell movements through the PS (Bottcher and Niehrs 2005). We wanted to investigate whether an FGF4–/– cell line (Wilder et al. 1997) would be able to i) differentiate into the endoderm lineage; and ii) promote differentiation of DE cell types, in that the latter would not be inhibited by the FGFRc isoform-activating FGF4. When cells were maintained as pluripotent cells, the FGF4–/– cell line showed a different cell morphology than the E14 and FGF4+/– cell lines. Cells grew in small, very dense clusters indicative of pluripotent cells (Figure 5A) and growth rates were slower, confirming the mitogenic effect of FGF4. Cells stained positive for the pluripotency marker OCT4 and negative for the endoderm marker Sox17, similar to the wt and heterozygote cell lines. We subjected the wt E14, FGF4+/– and FGF4–/– cell lines to our DE induction-protocol. Through antibody staining of SOX17, E-cadherin (Ecad) and FOXA2 we identified cells of an endoderm origin. Foxa2 is expressed during embryonic development in the anterior primitive streak, the newly formed definitive endoderm and is maintained throughout most mature endoderm-derived tissues (Kaestner et al. 1994; Weinstein et al. 1994). FGF4+/– cells behaved much like E14 wt cells, showing vast numbers of OCT4–/SOX17+ and SOX17+/FOXA2+/Ecadherin+ cells by day 5 (Figure 5B). In each cell line, differentiation was not 100% and small clusters of tightly connected, undifferentiated OCT4+ cells persisted in the culture. Remarkably, FGF4–/– cells readily differentiated along the endoderm lineage, showing mainly OCT4– /SOX17+ cells and only a few more OCT4+ cells than the wt and heterozygote cell lines (Figure 5A). When administrating FGF4 protein ectopically, the number of OCT4+ cells was reduced to wt levels. There were comparable numbers of SOX17+/FOXA2+/E-cadherin+ in the knock-out cell line and wt or heterozygote cell lines, and these did not change by the addition of FGF4 to the medium (Figure 5B). Thus, we conclude that FGF4 is dispensable for differentiation of mES cells along the endoderm lineage and for cells to leave the pluripotent state when the differentiation protocol applied includes activin. Definitive endoderm is formed in the absence of FGF4 Looking for expression of Sox7 and Thermostable direct hemolysin gene (Tdh), markers of visceral endoderm (VE; (Sherwood et al. 2007)), we wanted to see whether the endoderm formed was definitive or visceral. Staining for endoderm and VE markers in the

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undifferentiated cells, we found that E-cadherin was abundantly expressed but not SOX17 or SOX7 expression (Figure 6A). When cells had undergone differentiation for five days, E14 and FGF4+/– cell lines were SOX17+/E-cadherin+/SOX7– indicative of a DE identity (Figure 6B). FGF4–/– cells mainly showed the same expression pattern but also showed small clusters of SOX17+/E-cadherin+/SOX7+ cells indicating formation of VE in these areas (Figures 6B and S2). When adding ectopic FGF4 to the growth medium, hardly any SOX7+ cells were seen in the culture (Figure 6B). By qPCR analyses, we confirmed the DE phenotype of all three cell lines but did not see evidence of a VE sub-population in the FGF4–/– cell culture. We saw a large induction of Sox17 and especially Cxcr4 transcription upon DE-induction (Figure 6C) indicative of the formation of DE rather than VE. The VE marker Sox7 showed similar levels of expression in the pluripotent and differentiated states for all three cell lines and the absolute amount of transcription was very low, i.e. similar to Sox17-expression levels in mES cells. Tdh was expressed at intermediary levels in mES cells but was down-regulated upon DE-induction. Interestingly, the FGF4+/– cell line showed a somewhat elevated expression of Sox17 and Cxcr4 both in the pluripotent and differentiated states whereas the FGF4–/– cell line expressed these genes at levels comparable to wt cells. This suggests that FGF4 acts as a morphogen and that an intermediary expression level most efficiently induces DE-formation whereas high or low levels in the wt and FGF4–/– cell lines, respectively, fail to do so. In summary, we conclude that FGF4-signalling is dispensable for induction of DE in FGF4–/– mES cells and that an intermediary FGF4-koncentration may be beneficial to DE formation.

Discussion During embryonic development, epithelial tissues express FGFRb-isoforms while mesenchymal tissues express mainly FGFRc-isoforms (Ornitz and Itoh 2001). FGFs specifically activating FGFRb-isoforms (i.e. FGF7 and 10) are mainly expressed in the mesenchyme and FGFs activating FGFRc-isoforms (i.e. FGF4, 8 and 9) are mainly expressed in the epithelium, resulting in specificity during reciprocal epithelial-mesenchymal signalling in developing organs such as the lung, cecum, salivary glands and pancreas (Stark et al. 1991; Orr-Urtreger et al. 1993; Colvin et al. 2001; Ornitz and Itoh 2001; Elghazi et al. 2002; Manfroid et al. 2007). Data from mouse embryos obtained in our group confirm findings by others that in the developing pancreas FGFR2b and 4 are expressed in the epithelium, whereas FGFR1c and 2c are expressed in the mesenchyme (Kathrine Beck Sylvestersen, personal communication; (Stark et al. 1991; Orr-Urtreger et al. 1993; Elghazi et al. 2002)). In the present report we show how FGFR2b and 4 are up-regulated in the Sox17-GFPHi or DE-fraction during DE formation in mES cells. FGFR1c, 2c and 3c were up-regulated in the Sox17-GFPLo fraction alone or in both fractions. The reason for this unexpected high expression of FGFRc-isoforms in both the Sox17-GFPLo and Sox17-GFPHi fractions may be that the Sox17-GFPLo fraction contains a large pool of cells not yet committed to an epithelial fate or cells that are undifferentiated. The fraction is characterised by high numbers of non-mesodermal Sox17-GFP–/Lo/ FLK1–/ EpCAM+ expressing cells. These cells may still undergo differentiation and maturation and thus have the potential to later become epithelial endoderm or they may contain mesenchymal cells to some degree. The Sox17-GFPLo expressing cells may have a function in the culture similar to that of the mesenchyme in pancreatic development, i.e. signalling to direct cell fate of DE or foregut progenitors. Accordingly, preliminary data in our lab show that if Sox17-GFP cells are sorted after 5 days of DE formation, then re-plated and cultured under conditions inducing pancreatic progenitors (Hansson et al. 2009), the Sox17-GFPHi fraction will loose GFP expression and fail to turn on foregut markers such as SOX2 and PDX1 (Maria Winzi, personal communication). On the contrary, cells of the Sox17-GFPLo fraction will turn on GFP expression and differentiate into SOX2+, NKX6.1+ and PDX1+ positive cells similar to the unsorted culture. This suggests that signals from the Sox17-GFPLo to the Sox17-GFPHi cells include FGFs and is crucial for their propagation and ability to differentiate further.

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Most of the remaining cells are Sox17–/Lo/ FLK1–/ EpCAM– and these may represent a mesoderm population which is no longer expressing FLK1, i.e. is not haematopoietic. Flk1 is necessary for hematopoietic and endothelial development, but not for other mesodermal lineages expressing the marker at an early stage (Ema et al. 2006). We have previously shown that FGF-signalling has a positive effect on differentiation towards PS-type cells peaking on day 3 (T-GFP+ and Gsc-GFP+), but inhibitory effect on DE cells (Sox17-GFP+; (Hansson et al. 2009)). In the present study we show that FGF activating several FGFRc-isotypes elicit this effect and that FGFs activating only FGFRb-isotypes have no effect. This suggests that activation of FGFRc-isoforms is beneficial for early differentiation towards an intermediary PS-type cell, primarily through increased proliferation of cells induced to differentiate by activin or BMP4. Later, FGFRc-activation must be removed in order to optimize differentiation into a DE cell, probably because FGFRc-activation drives mesendodermal cells towards the mesodermal lineage. The FGFR-expression seen during mesendoderm/ DE-induction supports this, as mainly FGFRc-isoforms are expressed in the culture. FGFR2b is expressed late during DE-induction suggesting that the forming epithelium can respond to FGFs activating this receptor. However, we failed to substantiate this hypothesis as the number of Sox17-GFPHi cells were unaffected by FGF7 and 10. Possibly, this FGFR2b expression renders the cells competent to respond to later signals during organogenesis. Prior to gastrulation, FGF4 and 5, activating FGFRc-isoforms only, are expressed in the embryonic ectoderm in the area of the later PS, and in the PS during gastrulation (Haub and Goldfarb 1991; Hebert et al. 1991; Niswander and Martin 1992). During gastrulation, FGF3, activating FGFRb-isoforms only, is expressed in the PS (Wilkinson et al. 1988). This supports our suggestion that FGFRc-activation is important early in differentiation, although we do not see a requirement for FGFRb-activation during either mesoderm or endoderm induction. A combination of factors, activating FGFRc-isoforms early during differentiation and FGFRbisoforms later may be beneficial for future DE-induction. The FGFs having the largest effect on PS-marker induction were FGF4 and 6. FGF4 is an important growth factor during gastrulation where it is responsible for the cell movements through the PS (Bottcher and Niehrs 2005) and FGF4 knock-out mice die during gastrulation at E4-5 (Feldman et al. 1995). It has also been shown to be necessary for mESCs when leaving the pluripotent state and differentiating into either ectoderm or mesoderm lineages (Kunath et al. 2007; Stavridis et al. 2007). Kunath and co-workers found that this knock-out cell line could not differentiate into either lineage, except when supplementing the growth medium with FGF4 protein. Remarkably, using culture conditions similar to Kunath and co-workers, we found that ectopic FGF4 was dispensable when differentiating the FGF4–/– cell line into SOX17+/Ecadherin+/FOXA2+/SOX7– DE cells expressing Sox17 and Cxcr4, but not Sox7 and Tdh. This supports our previous finding that induction of DE-formation is not dependent on early FGFsignalling, as cells readily become Sox17-GFP+ in the presence of the FGFR-inhibitor PD173074 at early stages (Hansson et al. 2009). In the FGF4–/– cell line, slightly more cells stay in a pluripotent state, i.e. more cells stain OCT4+, than what is seen for wt and heterozygote cell lines. Some of these cells can be induced to differentiate when FGF4 is added to the differentiation medium, but FGF4 supplement does not seem to alter the fate of the differentiated cells. We propose that ectopic administration of FGF4 is only necessary for differentiation into ectoderm and mesoderm lineages but not for leaving the pluripotent state (Kunath et al. 2007; Stavridis et al. 2007). Although FGF4 knockout mice have been shown to be embryonic lethal at the stage of gastrulation (Feldman et al. 1995; Wilder et al.), their dependence on FGF4 signalling may lie at an earlier time-point, namely in the area of embryonic ectoderm where later the PS forms. This would render FGF4 necessary for formation of the PS rather than it’s function (Niswander and Martin 1992; Tam et al. 1993). This also shows that FGF4 is not necessary for cells to leave the pluripotent state when the differentiation protocol applied includes activin.

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FGF4, 5, FGF8b and FGFR1, are expressed during PS-formation and gastrulation (Haub and Goldfarb 1991; Hebert et al. 1991; Deng et al. 1994; Yamaguchi et al. 1994; Sun et al. 1999). FGFR1–/– or FGF8–/– embryos are embryonic lethal at this time point, the latter failing to express Fgf4 in the streak. In mES cell cultures, Fgf5 is upregulated at the onset of differentiation (Kunath et al. 2007). Interestingly, Kunath and co-workers could not rescue neural differentiation in the FGF4–/– cell line when adding FGF5 (Kunath et al. 2007). However, we suggest a redundancy in FGF-signalling at the point of initiation of endoderm differentiation taking place, in such that FGF5 or 8b, binding FGFRc-isoforms as does FGF4, facilitate the activation of key differentiation genes. Most studies on endoderm formation from mES cells rely on culturing conditions using either embryoid bodies as starting material or high cell densities (Funa et al. 2008; Morrison et al. 2008; Willems and Leyns 2008). Compared to Kunath and co-workers, we seed cells at a lower density. Possibly, an excess of cells to some degree inhibits differentiation, a common phenomenon seen in many ES cell differentiation systems. Indeed, when applying the ectoderm differentiation protocol as described by Kunath and co-workers to cells at low density, we saw a significant increase in differentiation (data not shown). It has been shown that only mES cells grown at high density loose their renewal properties after ROCK-inhibition (Chang et al. 2010). Cells grown at high densities have more cell-cell interactions and their β-catenin pool is partly located at the plasma membrane, activating the Wnt signalling pathway implicated in maintaining the pluripotent state in both human and mouse ES cells. Thus, we speculate that high cell densities preferentially retain mES cells in the pluripotent state to a higher degree than cells at low density and that FGF4-signalling may be necessary for leaving the pluripotent state at high cell densities only.

Acknowledgements We are thankful to Drs. G. Keller, S. Nishikawa, S. J. Morrison and A. Rizzino/ T. Kunath for the T-GFP, Gsc-GFP, Sox17-GFP, FGF4+/– and FGF4–/– cell lines, respectively. We thank Søren Refsgaard Lindskog for excellent technical assistance and Mads Daugaard for critically reading of the manuscript.

Figure legends Figure 1: Screen for FGFR-isotypes during DE differentiation in sorted fractions of Sox17GFP cells The expression of each FGFR-isoform was analysed by qPCR in both sorted and unsorted fractions of Sox17-GFP cells, differentiated by our DE protocol (30 ng/ml activin for 5 days). A) A histogram showing sorting gates in GFP–, GFPLo and GFPHi fractions. B) The absolute expression of each FGFR-isoform was standardised to the house-keeping gene TATA-binding protein (Tbp). Sox17Hi fractions are shown only at day 4 and 5, when they appeared in the culture. The fraction of Sox17-GFP– cells was to low for RNA-extraction. We did not obtain functional primers for FGFR3b. The relative mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s paired, two-tailed t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the ESC condition for each fraction (Sox17-GFP+ fractions were compared to the unsorted ESC sample). Figure 2: Activation of FGFRc-isoforms boosts mesendoderm but inhibits DE marker expression Using GFP-reporter cell lines T-GFP, Gsc-GFP and Sox17-GFP, we differentiated cells for 3 (T-GFP cell line only) and 5 days in BMP4- or activin-containing media, adding different FGFs. Cells were analysed by a FACS. A) Table of FGF – FGFR binding of selected FGFs 69

used in this paper. Modified from (Ornitz et al. 1996; Olsen et al. 2006; Zhang et al. 2006; Mason 2007). B-C) Cells were differentiated in 10 ng/ml BMP4 (= mesoderm-induction) or 1 ng/ml activin (= posterior streak/ mesoderm-induction) w/wo FGFs, and expression of T-GFP was measured at days 3 and 5. D) Gsc-GFP cells were differentiated in 30 ng/ml activin w/wo FGFs, and expression of GFP was measured at day 5. E) Sox17-GFP cells were differentiated in 30 ng/ml activin w/wo FGFs, and expression of GFP was measured at day 5. The mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the BMP4 or activin conditions. Figure 3: Activation of FGFRb or FGFRc-isoforms differentially affects the expression of PS, DE and mesoderm markers Sox17-GFP cells were differentiated in media containing BMP4 or activin w/wo FGFs for 3-5 days before harvest. Cells were stained for markers of PS, DE or mesoderm and analysed using a FACS. A & B) Cells were stained for T and analysed on days 3 and 5. Shown here are the relative number of T+ cells in the Sox17-GFPHi fraction A), or the Sox17-GFP–/Lo fraction B). The mean expression ± S.E.M. of 2 independent experiments are shown, no statistical analysis performed. C-G) Cells were stained for FLK1 and EpCAM and analysed in either single channel for C) Sox17-GFP, D) FLK1, E) EpCAM; or in multichannel for the Sox17-GFPHi fraction F), or Sox17-GFP–/Lo fraction G) dividing data into four fractions: FLK1–/EpCAM+; FLK1+/EpCAM+; FLK1–/EpCAM–; FLK1+/EpCAM–. The mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s paired, two-tailed t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the activin conditions. Figure 4: FGFs activating FGFRc-isoforms affect early cell growth and proliferation A wt mES cell line (E14) was grown in media containing activin w/wo FGFs and harvested for analysis of total cell number and proliferation on days 3 &5. A count of total number of cells and relative proliferation of cells is shown for day 3 A) and day 5 B). The mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the activin conditions. Figure 5: FGF4-signalling is dispensable for endoderm differentiation E14, FGF4+/– and FGF4–/– cells were stained for markers of pluripotency and endoderm. A) Undifferentiated cells were stained for OCT4 and SOX17 and the nuclear stain DAPI. B) Cells were differentiated for 5 days in 30 ng/ml activin w/wo 5 ng/ml FGF4 and stained for OCT4 and endoderm markers SOX17, FOXA2, E-cadherin (Ecad) and DAPI. Representative images are shown for each condition. Scale bar: 100 µm. Figure 6: In the absence of FGF4, DE rather than VE is formed Antibody stain and qPCR-analyses of DE and VE markers in E14, FGF4+/– and FGF4–/– cells. A) Undifferentiated cells were stained for SOX17, E-cadherin, SOX7 and the nuclear stain DAPI. B) Cells were differentiated for 5 days in 30 ng/ml activin w/wo 5 ng/ml FGF4 and stained for the same markers. Representative images are shown for each condition. Scale bar: 100 µm. C) qPCR data showing the relative expression levels of Sox17, Cxcr4, Sox7 and Tdh mRNA present in undifferentiated and differentiated cells of each cell line compared to E14 ESCs, all standardized to the house-keeping gene Tbp. The mean expression ± S.E.M. of 3 independent experiments is shown (n=2 for FGF4–/– cells treated w/ activin + FGF4), using a Ratio t-test for the statistical analysis: # = P < 0,05; ## = P < 0,01 compared to the E14 mES cell condition. SUPPLEMENTARY DATA Figure S1: Negative controls of EdU-incorporation and sort gate A histogram showing the distribution of pluripotent E14 mES cells with/without EdUincorporation and without EdU-stain (red and black samples); without EdU-incorporation and

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with EdU-stain (blue sample); and w/ EdU-incorporation and with EdU-stain (stained with Alexa-488, i.e. positive control; green sample). Figure S2: A few areas of SOX7+ cells in the FGF4–/– cell line after DE-induction Cells were differentiated for 5 days in 30 ng/ml activin w/wo 5 ng/ml FGF4 and stained for SOX17, E-cadherin, SOX7 and the nuclear stain DAPI. The yellow frame indicates area blown up and shown to the right (red signal boosted). Representative images are shown for each condition, except for FGF4–/– cells in activin alone which shows a SOX7+ area in the culture. White scale bar: 100 µm; yellow scale bar: 50 µm.

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Figures

Figure 1: Screen for FGFR-isotypes during DE differentiation in sorted fractions of Sox17-GFP cells. The expression of each FGFR-isoform was analysed by qPCR in both sorted and unsorted fractions of Sox17-GFP cells, differentiated by our DE protocol (30 ng/ml activin for 5 days). A) A histogram showing sorting gates in GFP–, GFPLo and GFPHi fractions. B) The absolute expression of each FGFR-isoform was standardised to the housekeeping gene TATA-binding protein (Tbp). Sox17Hi fractions are shown only at day 4 and 5, when they appeared in the culture. The fraction of Sox17-GFP– cells was to low for RNA-extraction. We did not obtain functional primers for FGFR3b. The relative mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s paired, two-tailed t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the ESC condition for each fraction (Sox17-GFP+ fractions were compared to the unsorted ESC sample).

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Figure 2: Activation of FGFRc-isoforms boosts mesendoderm but inhibits DE marker expression. Using GFPreporter cell lines T-GFP, Gsc-GFP and Sox17-GFP, we differentiated cells for 3 (T-GFP cell line only) and 5 days in BMP4- or activin-containing media, adding different FGFs. Cells were analysed by a FACS. A) Table of FGF – FGFR binding of selected FGFs used in this paper. Modified from {Olsen, 2006 #25;Mason, 2007 #21;Ornitz, 1996 #20;Zhang, 2006 #19}. B-C) Cells were differentiated in 10 ng/ml BMP4 (= mesoderm-induction) or 1 ng/ml activin (= posterior streak/ mesoderm-induction) w/wo FGFs, and expression of T-GFP was measured at days 3 and 5. D) Gsc-GFP cells were differentiated in 30 ng/ml activin w/wo FGFs, and expression of GFP was measured at day 5. E) Sox17-GFP cells were differentiated in 30 ng/ml activin w/wo FGFs, and expression of GFP was measured at day 5. The mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the BMP4 or activin conditions.

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Figure 3: Activation of FGFRb or FGFRc-isoforms differentially affects the expression of PS, DE and mesoderm markers. Sox17-GFP cells were differentiated in media containing BMP4 or activin w/wo FGFs for 3-5 days before harvest. Cells were stained for markers of PS, DE or mesoderm and analysed using a FACS. A & B) Cells were stained for T and analysed on days 3 and 5. Shown here are the relative number of T+ cells in the Sox17GFPHi fraction A), or the Sox17-GFP–/Lo fraction B). The mean expression ± S.E.M. of 2 independent experiments are shown, no statistical analysis performed. C-G) Cells were stained for FLK1 and EpCAM and analysed in either single channel for C) Sox17-GFP, D) FLK1, E) EpCAM; or in multichannel for the Sox17-GFPHi fraction F), or Sox17-GFP–/Lo fraction G) dividing data into four fractions: FLK1–/EpCAM+; FLK1+/EpCAM+; FLK1–/EpCAM–; FLK1+/EpCAM–. The mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s paired, two-tailed t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the activin conditions.

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Figure 4: FGFs activating FGFRc-isoforms affect early cell growth and proliferation. A wt mES cell line (E14) was grown in media containing activin w/wo FGFs and harvested for analysis of total cell number and proliferation on days 3 &5. A count of total number of cells and relative proliferation of cells is shown for day 3 A) and day 5 B). The mean expression ± S.E.M. of 3 independent experiments is shown, using a Student’s t-test for the statistical analysis: * = P < 0,05; ** = P < 0,01 compared to the activin conditions.

Opposite, Figure 5: FGF4-signalling is dispensable for endoderm differentiation. E14, FGF4+/– and FGF4–/– cells were stained for markers of pluripotency and endoderm. A) Undifferentiated cells were stained for OCT4 and SOX17 and the nuclear stain DAPI. B) Cells were differentiated for 5 days in 30 ng/ml activin w/wo 5 ng/ml FGF4 and stained for OCT4 and endoderm markers SOX17, FOXA2, E-cadherin (Ecad) and DAPI. Representative images are shown for each condition. Scale bar: 100 µm.

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Opposite, Figure 6: In the absence of FGF4, DE rather than VE is formed. Antibody stain and qPCR-analyses of DE and VE markers in E14, FGF4+/– and FGF4–/– cells. A) Undifferentiated cells were stained for SOX17, Ecadherin, SOX7 and the nuclear stain DAPI. B) Cells were differentiated for 5 days in 30 ng/ml activin w/wo 5 ng/ml FGF4 and stained for the same markers. Representative images are shown for each condition. Scale bar: 100 µm. C) qPCR data showing the relative expression levels of Sox17, Cxcr4, Sox7 and Tdh mRNA present in undifferentiated and differentiated cells of each cell line compared to E14 ESCs, all standardized to the housekeeping gene Tbp. The mean expression ± S.E.M. of 3 independent experiments is shown (n=2 for FGF4–/– cells treated w/ activin + FGF4), using a Ratio t-test for the statistical analysis: # = P < 0,05; ## = P < 0,01 compared to the E14 mES cell condition.

SUPPLEMENTARY DATA

Figure S1: Negative controls of EdU-incorporation and sort gate. A histogram showing the distribution of pluripotent E14 mES cells with/without EdU-incorporation and without EdU-stain (red and black samples); without EdU-incorporation and with EdU-stain (blue sample); and w/ EdU-incorporation and with EdU-stain (stained with Alexa-488, i.e. positive control; green sample).

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Figure S2: A few areas of SOX7+ cells in the FGF4–/– cell line after DE-induction. Cells were differentiated for 5 days in 30 ng/ml activin w/wo 5 ng/ml FGF4 and stained for SOX17, E-cadherin, SOX7 and the nuclear stain DAPI. The yellow frame indicates area blown up and shown to the right (red signal boosted). Representative images are shown for each condition, except for FGF4–/– cells in activin alone which shows a SOX7+ area in the culture. White scale bar: 100 µm; yellow scale bar: 50 µm.

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6. General discussion Endoderm differentiation Basic culture conditions Our culture system is based on serum and feeder-free conditions by means of gelatine-coated culture flasks and the N2, B27, BMP4 and LIF supplements. Remnants of serum and feeders have been removed by growing cells for 3 passages, i.e. 6 days, under these conditions prior to experimental setup. In the presence of LIF and BMP4 cells do not differentiate, but it is hard to determine the effect of trace amounts of BMP4 on the very early differentiation in the culture. However, such differentiation would most likely be opposed by trace amounts of LIF and therefore not have any practical effect especially when cells are differentiated using high concentrations of growth factors. A recent study showed that inhibition of FGFRs, MEK and GSK3 through addition of the small molecule inhibitors SU5402, PD184352 and Chir99021 respectively, could maintain cells pluripotent over time (Ying et al. 2008). This protocol was refined by using the stronger MEKinhibitor PD0325901 in combination with Chir99021 (Nichols et al. 2009). Thus, LIF and BMP4 are not necessary for maintenance of pluripotency, as inhibition of the MAPK-pathway and GSK3 is sufficient. This protocol may be preferred in the future to avoid maintenance of the pluripotent state through addition of e.g. BMP4, a potent inducer of mesoderm differentiation. ‘Mesendoderm’: Does it exist? Nodal and BMP4, both members of the TGFβ superfamily are expressed in the PS and ExE, respectively, and act in opposite directions to induce DE and mesoderm. These two germ layers derive from the same population of epiblast cells migrating through the PS during gastrulation. Cells moving through the PS are bipotent and are collectively termed the mesendoderm before they are further differentiated into mesoderm and endoderm (Lawson et al. 1991; Kinder et al. 2001; Rodaway and Patient 2001). This mesendoderm population seems much conserved, as it is found from Caenorhabditis elegans (C. elegans) through Xenopus laevis (Xenopus) to zebrafish (Rodaway et al. 1999; Rodaway and Patient 2001). A bipotent mesendoderm cell population has also been shown in mES cell work where a pool of cells expressing T or GSC/FOXA2/PDGFRα, i.e. a mesendoderm population, had the potential to form both mesoderm and endoderm dependent on the growth factors presented (Kubo et al. 2004; Tada et al. 2005). Indeed, we found that by using GFP-reporter cell lines for the PS markers T, Gsc and Mixl1, we could analyse the effect of anteriorising and posteriorising TGFβ- and WNTsignalling in the early differentiation steps. Although the mesendoderm cell population is well-established from C. elegans to mouse development, it is challenged by a recent study. By use of single-cell lineage tracing, the authors claim that endoderm and surface ectoderm segregate during gastrulation, whereas a pool of bipotent neuromesoderm persists through all stages of axis elongation (Tzouanacou et al. 2009). If these observations hold true, it has implication for ES cell differentiation towards cells of the DE lineage, as it will then be better to obtain a pool of cells expressing DE markers only, and not a combination of DE and mesoderm markers at early stages. Other groups and we have shown derivation of DE cells from a population of cells expressing app. 40 – 70% mesendoderm markers (Tada et al. 2005). Whether these DE cells derive from a mesendoderm population or rather from an endoderm population directly, is difficult to establish without the use of lineage tracing during differentiation. Also, any contribution of signals from randomly differentiated cells to the forming DE may prove important, but are as yet not studied in depth. All in all this mesendoderm population may prove not to have a practical limitation to ES cell differentiation in vitro as long as the DE is properly established.

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The definitive problem of endoderm When initiating differentiation into endoderm there is a great concern as to whether it has become visceral or definitive endoderm. These cell types share many markers (Sox17, Foxa2, E-cadherin), and it is therefore necessary to test the obtained cell population for the diverging markers to make a solid conclusion. In pratice this has proven less important, as induction of DE is based on addition of activin, in which visceral endoderm (VE) does not form (Yasunaga et al. 2005). However, it is still necessary to show more than the shared markers to draw convincing conclusions. In the developing embryo, the forming DE cells have traditionally been believed to replace the outer layer of VE, thus forming a homogeneous cell layer. A recent study performing lineagetracing of VE cells in the mouse embryo suggests that DE cells insert into the VE cell population (Kwon et al. 2008). The VE cells are diluted by the faster proliferating DE cells, but VE cells are found in the gut tube lining as late as the 16-18 somite stages. This indicates that the VE does indeed contribute to the embryo proper and suggests that it may not be detrimental to have a small population of VE cells in the differentiating culture after all.

FGFR isoform-specific activation in differentiation FGFs in evolution: why are there so many in Mammalia? The temporal and spatial expression of FGFs and FGFRs determine signalling at any given time. The large number of FGFs and FGFRs may seem excessive since a simpler system would be more ‘cost-efficient’ to the cell. FGFs are found only in multicellular organisms (metazoa), ranging from the nematode C. elegans to mouse and human (Itoh and Ornitz 2004). In C. elegans there are two FGFs and one FGFR and in Drosophila melanogaster there are three FGFs (Ornitz and Itoh 2001; Itoh and Ornitz 2004). The reason behind the large family of FGFs found in mouse and human lies in two phases of expansion during evolution. The first expansion occurred by genome duplication before chordate evolution. The second expansion occurred during early vertebrate evolution by two large-scale gene-duplications, possibly of the whole genome each time (Itoh and Ornitz 2004). This resulted in the 22 FGFs found in mice and humans today. FGFRs co-evolved with the FGFs and they have since evolved splicevariants, which add to the complexity of ligand-receptor specificity and downstream signalling (Itoh and Ornitz 2004). The temporal and spatial expression of FGFs and FGFRs determine signalling at any given time, and an additional layer of complexity is added by the evolution of HSs, which are also differentially expressed (Allen and Rapraeger 2003). The many different functions of FGFs during development and adult homeostasis is probably the basis for the abundance of FGFs and FGFRs found in mammals, as specific regulation of their function is important. Most null mutants show different phenotypes, arguing for distinct functions of each FGF during development. These specific functions are tissue- and time pointdependent: some FGFs are expressed at the same time in certain tissues, but have distinct functions in other tissues, as is the case for the FGF8 family, comprising FGF8, 17 and 18. These are co-expressed in the midbrain-hindbrain junction, but FGF8 and FGF18 show differential expression and effects in limb and bone development, respectively (Xu et al. 2000; Liu et al. 2002). This overlapping expression of FGFs belonging to the same sub-family, and their shared FGFR-binding properties, argue for a functional redundancy among FGFs (Itoh and Ornitz 2004). Redundancy between sub-families is also seen, for example by FGF3 and 8 in the induction of otic placode and forebrain development in zebrafish (Liu et al. 2003; Walshe and Mason 2003). In embryonic development, the specific activation of FGFRs is not only dependent on the concentration of FGFs secreted, but also on their diffusion through the extracellular matrix where HS retains them. The effective dose may therefore be different among cells, and could explain e.g. the differential expression of epiblast and primitive endoderm markers in the ICM, which was recently shown to be FGF concentration-dependent (Yamanaka et al. 2010). 84

Overall, the multitude of FGF-signalling is huge and has to be interpreted in the spatio-temporal context in which it is investigated. In in vitro ES cell differentiation, however, one can make use of redundant functions of the different FGFs as the effect upon differentiation is only dependent on the expression of FGFRs and not necessarily on the endogenous range of FGFs expressed. Still, the endogenously expressed FGFs must be taken into account when aiming for the optimal protocol to ensure no conflicting signalling is taking place in the cells. FGFR-isoforms in DE formation and patterning We have shown that FGFs activating FGFRc-isoforms increase the numbers of cells expressing mesendoderm markers but reduce the numbers of Sox17-GFP+ DE cells. On the other hand, FGFs activating only FGFRb-isoforms have no effect on these cells, most likely due to their lack of expression. This argues for a beneficial effect of FGFRc-activation during early stage DE-induction, namely in the generation of a mesendodermal population of cells, but an inhibitory effect of these same FGFs during the later DE-specification. Also, the concentrations of FGFs used in this study may prove to be important. DE-formation is dependent on FGFRsignalling, as inhibition by small molecules inhibits Sox17-GFP+ cells. Likewise, high concentrations of FGFs have an inhibitory effect on this same population, arguing for an optimal intermediate concentration. Such an intermediate concentration is possibly even below the endogenous FGF concentration, as the FGF4+/– cell line seems to differentiate into DE at a higher success that the wt or the FGF4–/– cell line, even when the latter is supplemented with a low concentration of FGF4. Meanwhile, experiments in ‘Step 2’ of the Pdx1-inducing protocol (Chapter 4) show that addition of FGFs increases the number of Pdx1-GFP+ cells, possibly through their mitogenic effect. Furthermore, FGF7 and 10 function even better than FGF4 in induction of Pdx1-GFP+ cells (Nina Engberg and Claude Rescan, unpublished data), arguing that FGFRb-isoforms are present and have an additional role to the mitogenic alone. These DE cells are epithelial and as such probably require activation of FGFRb-isoforms during DE patterning and/ or organogenesis. This would make sense from a developmental point of view as the development of epithelial components in many organs depends on FGF10 for epithelio-mesenchymal interactions (Min et al. 1998; Ohuchi et al. 2000). Also, FGFRb-isoforms are expressed in epithelial tissues during development (Kathrine Beck Sylvestersen, unpublished data; (Ornitz and Itoh 2001)). Overall, addition of FGFs activating FGFRc-isoforms during mesendoderm formation followed by absence of FGFs during DE-formation and finally FGFRb isoform-activation during induction of posterior foregut, i.e. Pdx1-expressing cells, could prove the most beneficial. Successful DE formation in an FGF4 null cell line At a low cell density, we see that the FGF4–/– cell line readily differentiates into DE, showing independence of FGF4-signalling in induction of differentiation. This was surprising, as a previous study showed dependence for FGF4 in the induction of ectoderm and mesoderm differentiation, thus suggesting that FGF4-signalling is needed for cells to leave the pluripotent state altogether (Kunath et al. 2007). This inhibition of differentiation could be reverted by addition of FGF4, but not by FGF5, which is expressed early during differentiation and activates FGFR1c as does FGF4 (Haub and Goldfarb 1991). However, FGF4 additionally activates FGFR2c, 3c and 4 and a redundancy by FGF6 or 8(b) may prove significant as these bind FGFR1c, 2c, 4 and FGFR2c, 3c, 4, respectively, similar to FGF4 (Ornitz et al. 1996; Zhang et al. 2006; Mason 2007). From in vitro studies of mammalian cell cultures, it is known that there is a positive correlation between cell density and cellular response, measured by receptor phosphorylation or gene expression (Polk et al. 1995; Bedrin et al. 1997; Batt and Roberts 1998; Mukhopadhyay et al. 1998). On the other hand, an inverse correlation between cell density and gene expression has been shown in other systems (Li and Goldstein 1996; Singh et al. 1996; Posern et al. 1998). In ES cell cultures little is known about cell density and gene expression responses. It was shown that cells grown at high densities have a pool of β-catenin located at the cell surface, where it is

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involved in WNT-signalling and maintenance of the pluripotent state (Chang et al. 2010). Thus, here it seems that a high cell density may inhibit differentiation, which has also been suggested by Smith and co-workers (Smith et al. 1992). Similarly, we propose inhibition by high cell densities in the FGF4–/– ES cell culture, and suggest low seeding densities for optimal differentiation.

DE patterning It seems rather symptomatic that we successfully reach 40-60% SOX17+ DE cells, but have limited success in further patterning this DE to PDX1-expressing posterior foregut. We saw an induction of PDX1-expressing cells when adding RA and FGF (and Cyclopamine), but never more than 2-4% on average. Competing groups have successfully obtained 32% PDX1+ cells in hES cell cultures by using some of the same posterior foregut-inducing factors, basing their scientific approach on the same hypotheses as we do (Johannesson et al. 2009; Ameri et al. 2010). Therefore the reason for our inefficient Pdx1-GFP induction shall possibly be sought elsewhere. The Pdx1-GFP cell line we use has shown a nice expression pattern within the posterior foregut region when injected into mouse blastocysts to form chimaeras (Tino Klein, unpublished data). The cell line works in vivo and is pluripotent, and we therefore expect it to also work in vitro. One point could be that we have optimized the DE-induction step in the Sox17-GFP and not the Pdx1-GFP cell line. Data from hES cells indicate that the outcome of a differentiation protocol varies much between cell lines (D'Amour et al. 2006; Mfopou et al. 2010). If this is also the case for mES cells, we will have to redo optimization of DE-induction in our Pdx1-GFP cell line to have the best starting material. However, our protocol induces 2-4% Pdx1+ cells in E14, Pdx1-LacZ and Pdx1-GFP cell lines, showing robustness of the protocol. A developmental-based explanation is that the DE we have is simply not the ‘correct’ one. Using our protocol, we get many SOX2+ cells, suggesting that the DE we have after 5 days in high concentrations of activin may be somehow pre-patterned to respond to patterning factors predominantly by induction of anterior foregut, marked by SOX2. Finally, there could be one or more components in our basic medium or medium supplements that inhibit differentiation. This problem could be overcome by changing the basic medium, medium supplements and the culture dish coating individually to decipher which may be inhibitory.

Cell replacement therapy as a future cure for TIDM ES cells are not the only source of β cells or β-like cells envisioned as material for future transplantation in the treatment or even cure for diabetes. Some perspectives of the various alternatives are discussed below. Xeno-transplantation Xeno-transplantation of islets of Langerhans from pig to primate is being investigated as a treatment for type I diabetes. The use of pigs is promising, as their vascular physiology is similar to that of humans and they are relatively cheap to breed. Furthermore, the so-called mini-pigs weighing app. 120 kg are similar to humans in organ and body sizes and can be inbred to homozygosity at e.g. the porcine major histocompatibility complex (MCH). The latter holds a great potential for manipulation to create customized donor organs/ cells and overcome some of the problems of immune-responses normally seen in transplantation. This could be done by introducing porcine MHC genes into the bone marrow of the recipient human inducing mixed chimaerism thereby introducing immunological tolerance to the xenograft (Hoerbelt and Madsen 2004). Alternatively, pigs could be genetically manipulated not to express gene products to which the recipient immune system reacts. Of major concern in xeno86

transplantation is the spread of animal diseases to humans. Some of the risk factors may be eliminated by breeding homozygous mini-pigs in controlled environments. EpiSCs, hESCs and iPSCs It has recently been shown that epiblast stem cells (epiSCs) derived from post-implantation mouse blastocysts show characteristics of hES cells in their need for pluripotency-maintaining factors activin and FGF2 (Brons et al. 2007; Tesar et al. 2007; Vallier et al. 2009). Also, their response to differentiation-inducing factors is more similar to hES cells than what is seen for mES cells (Vallier et al. 2009). This close resemblance to hES cells may make epiSCs a better model for studying differentiation, as extrapolation of knowledge to the hES cell field may prove easier and more valuable. One major advantage is that existing ES cell lines can be converted into epiSCs without new derivation from mouse embryos (Guo et al. 2009), making already established transgenic cell lines readily transferable by a low work load. This may hold great potential for better inter-species protocol transfer between epiSCs and hES cells. In 2006, the Yamanaka-group showed that mature somatic cells, i.e. skin fibroblasts, can be induced to achieve an ES cell-like phenotype, i.e. become pluripotent and are reported to behave in the same way as mES or hES cells upon differentiation. These induced pluripotent stem (iPS) cells were generated by introduction of four transcription factors Oct4, Sox2, Klf4, and C-myc (Takahashi and Yamanaka 2006). This was done in mice, and the protocol has since been modified in several ways and has been transferred to human cells (Takahashi et al. 2007; Yamanaka 2009). iPS cells hold the potential for development of patient-specific pluripotent stem cell lines, which can be differentiated into any cell type of choice. They therefore represent a source of transplantable cells, which eliminates the need for immune-suppressing agents to a large degree. Although this is a very positive future application, in reality it may prove much too expensive for actual treatment. iPS cells will likely be important tools for modelling of and investigating the aetiology of (inherited) human diseases, which are not discovered until the disease state is complete. For instance, type I diabetes is normally not discovered until patients suffer from high blood glucose levels at which time point their β cell mass is practically obsolete (Maehr et al. 2009). Whether iPS cells will serve as material for cell replacement-therapies is still to be seen. Of major concern is to ensure that the genomic reprogramming of the cells is complete, a trait believed necessary for the cells to adopt the correct fate upon exposure to differentiation-inducing conditions (Yamanaka 2009). Also, teratoma formation from fully reprogrammed iPS cells cannot be avoided so far, making them unsuited for treatment in humans at this point. In general, avoiding teratoma-formation from differentiated cell populations is a major concern in transplantation. It is not acceptable to cure for instance diabetes but at the same time induce a cancerous condition, and as long as this risk exists with cell therapy-protocols, they will not be approved for treatment. Generation of β cells from existing cell sources in the pancreas The presence of a pancreatic stem cell, which has clonogenic potential, is multipotent and can be induced to generate insulin-producing cells in vitro has been suggested in both mice and humans (Ramiya et al. 2000; Seaberg et al. 2004; Zhao et al. 2007). However, it is speculated that these cells only show such stem cell-like properties due to the in vitro culture conditions (Baeyens and Bouwens 2008). A more convincing in vivo study showed the presence of islet precursors that could be activated upon serious tissue injury by the so-called partial ductligation, in which facultative multipotent progenitor cells in the ductal lining differentiate and proliferate into functional β cells (Xu et al. 2008). An alternative approach is to generate β cells by reprogramming of existing endocrine or exocrine cells in the pancreas. Following pancreatectomy to a mild or severe degree (70% and 95% respectively), regeneration of β cell mass is achieved through either replication of existing β cells or through neogenesis of precursor cells in addition to replication (Dor et al. 2004; Bouwens and Rooman 2005). Exocrine ductal cells show convincing potential as they seem able to contribute to glucose-responsive β cells through reprogramming (Baeyens and Bouwens

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2008). Lineage tracing of these cells has not yet been performed but will elucidate whether the increases in β cell mass are truly by neogenesis of ductal cells. Acinar cells cultured in vitro show dedifferentiation, possibly to the stage of the common acinar and endocrine cell which is found during pancreas development. They can subsequently be re-specified into hepatocytes in human cultures and β cells in mouse cultures, showing a large amount of plasticity of these cells (Lardon et al. 2004; Okuno et al. 2007). Whether it is possible to circumvent the patient’s auto-immune response towards these β cells is still to be seen. For the moment, life-long treatment with immune-suppressors represents the only alternative and is not desirable due to inherent side effects.

Concluding remarks The reason for putting a large effort into differentiation in a 2-dimensional, serum and feederfree protocol is first of all to be able to better control the signalling events going on in the dish and thereby better direct differentiation into the cell types of desire. A second and not less important point is that cell culture with animal products is not desirable due to fear of spread of disease. The latter especially has major implications in xeno-transplantations. Recently, a publication showing derivation and culture of a xeno-free hES cell line was published (Ellerstrom et al. 2006). In the future, this could very well be the only material we are comfortable with using for transplantation and it will as such have major impact on the application of the various differentiation protocols applicable to clinical work.

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Acknowledgements The work presented here has been made possible first and foremost by Palle Serup and Helle V. Petersen offering me a position as a Ph.D.-student in their department and through the supervision by Palle Serup. The former Department of Developmental Biology (now ‘Dept.s of Stem Cell Biology’ and ‘β-Cell Regeneration’) is an inspiring and pleasant workplace, both during and outside work-hours. I thank all of you for 5 really good years at Hagedorn and the stem cell group in particular for primarily supplying the scientific input. Thanks to Søren R. Lindskog and Gurmeet K. Singh for help on practical lab work. A special thanks to Nina Engberg, my significant other in the lab, office, cell room and in achieving the Master’s degree and working through the Ph.D. I am grateful to Jan N. Jensen, Tino Klein and Mattias Hansson, whose constructive natures helped me at critical time points. Thanks to Anette Bjerregaard for being the social backbone of our department. Outside work, I have been lucky to have a very supportive family and friends bringing much fun, seriousness and good times into my life. I’m always amazed at how many people can be persuaded to come and party on Mors! I could not have done this without you, Mads: you believe in me like no one else! A special thanks to Hjalte, my dancing tiger for demanding focus on what’s important and always making me happy.

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