primary cilia and coordination of signaling pathways

F A C U L T Y O F S C I E N C E U N I V E R S I T Y O F C O P E N H A G E N

PRIMARY CILIA AND COORDINATION OF SIGNALING PATHWAYS IN HEART DEVELOPMENT AND TISSUE HOMEOSTASIS

PHD THESIS BY

CHRISTIAN ALEXANDRO CLEMENT DEPARTMENT OF BIOLOGY SECTION OF CELL AND DEVELOPMENTAL BIOLOGY NOVEMBER, 2009

Christian Alexandro Clement, PhD thesis 2009 1/45

PREFACE

The work presented in this thesis was performed at the Department of Biology (DB), Section of cell and developmental Biology in the August Krogh building at the University of Copenhagen, Denmark. The thesis spanned over a three year period from 1‐11‐2006 to 1‐11‐2009 under the supervision of Associate professor, Søren Tvorup Christensen, PhD, and in close collaboration with Associate professor, Centre vice‐director Lars Allan Larsen, MSc, PhD at Wilhelm Johannsen Centre for Functional Genome Research, Department of Cellular and Molecular Medicine, University of Copenhagen. Furthermore, important collaborations were made with Professor Bradley K. Yoder, from the University of Alabama at Birmingham and Professor Kjeld Møllgård, at the Department of Cellular and Molecular Medicine, the Panum institute, University of Copenhagen. Professor Yoder contributed with the Ift88‐/‐ embryos and chimera mouse data, and Professor Møllgård contributed with the sectioning and H&E staining of the Ift88‐/‐ mice.

This study was carried out with financial support from the PhD School at the Faculty of Science, University of Copenhagen (C.A.C.), the Lundbeck Foundation, the Danish Science Research Council no. 272‐07‐0530 (S.T.C.), the Danish Heart Association (L.A.L.), NIH RO1 HD056030 (B.K.Y.) and GM60992 (G.J.P.). The Wilhelm Johannsen Centre for Functional Genome Research is established by the Danish National Research Foundation.

The papers included in this thesis are:

• [1] Expression and localization of the progesterone receptor in mouse and human reproductive organs. Teilmann SC, Clement CA, Thorup J, Byskov AG, Christensen ST. Journal of Endocrinology December (2006),191:525–535

• [2] Characterization of primary cilia and Hedgehog signaling during development of the human pancreas and in human pancreatic duct cancer cell lines. Nielsen SK, Møllgård K, Clement CA, Veland IR, Awan A, Yoder BK, Novak I, Christensen ST. Developmental Dynamics August, (2008), 237:2039–2052

• [3] Human embryonic stem cells in culture possess primary cilia with Hedgehog signaling machinery. Kiprilov EN, Awan A, Desprat R, Velho M, Clement CA, Byskov AG, Andersen CY, Satir P, Bouhassira EE, Christensen ST, Hirsch RE. The Journal of Cell Biology, March, (2008), Vol. 180, No. 5:897–904

• [4] The primary cilium coordinates early cardiogenesis and Hedgehog signaling in cardiomyocyte differentiation. Clement CA, Kristensen SG, Møllgård K, Pazour GJ, Yoder BK, Larsen LA, Christensen ST. September, (2009) Journal of Cell Science, 122:3070‐82

• [5] Using nucleofection of siRNA constructs for knockdown of primary cilia in P19.CL6 cancer stem cells differentiation into cardiomyocytes. Clement CA, Christensen ST, Larsen L.A, (2009) Methods in Cell Biology. December issue. In press.

In the thesis, these articles are referred to by their number in brackets, the rest by the first author and year.


ACKNOWLEDGEMENTS

First, I would like to thank my supervisor Dr. Søren T. Christensen for professional guidance, fresh ideas and an amazing collaboration throughout the three‐year period of this thesis. Second, I would like to thank Dr. Lars A. Larsen for a close and productive collaboration on the P19.CL6 EC stem cell and heart development project. I would also like to thank Professor Møllgård and Professor Yoder for a very fruitful collaboration and guidance on the heart development project. A special thanks to Stine G. Kristensen for the much‐appreciated contribution on the P19.CL6 work.

I wish to thank my family and Ditte Lynge Hansen for the support while home and during my three months stay in Brad´s lab in Alabama. A special thanks to my uncle Ian H. Lambert for moral support and in the general guidance in the field of biology on a daily basis.


ABBREVIATIONS

AVC: Atriumventricular canal

BCC: Basal cell carcinoma

BMP: Bone morphogenetic protein

CHD: Congenital heart disease

CNC: Cardiac neural crest

DAPI: 4’‐6’‐diamidino‐2‐phenylindole

Dhh: Desert hedgehog

DMSO: Dimethyl sulfoxide

E: Embryonic day

EC: Embryonal carcinoma

EM: Electron microscopy

EMT: Endocardial epithelial‐mesenchymal

transformation

FGF: Fibroblast growth factor

FHF: First heart field (also known as

primary heart field)

Gata4: GATA binding protein 4

Gli: Glioma transcription factor

H&E: Hematoxylin and eosin staining

hESC: Human embryonic stem cell

Hh: Hedgehog

ICM: Inner cell mass

IFM: Immunofluorescence microscopy

IFT: Intraflagellar transport

IGF1: Insulin‐like Growth Factor 1

Ihh: Indian hedgehog

IVS: Interventricular septum

JNK: Jun aminoterminal kinase

Kif: Kinesin superfamily protein

LIF: Leukemia inhibitory factor

LR: Left/Right

LV: Left ventricle

Mchr1: Melanin‐concentrating hormone receptor 1

mEC: Mouse embryonal carcinoma

Mef2c: Myocyte enhancer factor 2C

mESC: Mouse embryonic stem cells

MT: Microtubuli

Nkx25: NK2 transcription factor related, locus 5

OFT: Outflow tract

PanIN: Pancreatic intraepithelial neoplasia

PCM: Pericentriolar material

PCP: Planar cell polarity pathway

PDGFRα: Platelet‐derived growth factor receptor alpha

PKD: Polycystic kidney disease

Pkd1: Polycystin 1

Pkd2: Polycystin 2

PR: Progesterone receptor

Ptc: Patched

QRTPCR: Quantitative real‐time PCR

RA: Retionic acid

RNA: Ribo‐nucleic acid

RNAi: RNA interference

RTK: Receptor tyrosine kinases

RV: Right ventricle

SA: Sinuatrial

SEM: Scanning electron microscopy

SHF: Second heart field

Shh: Sonic hedgehog

siRNA: Small interfering RNA

Smo: Smoothened

Sst3R: Somatostatin Sst 3 receptor

Sufu: Suppressor of fused

TEM: Transmission electron microscopy

Tg737ORPK: Oak Ridge polecystic kidney mouse with

defect in gene Tg737 (encoding Polaris)

(TGF)Beta: Transforming growth factor beta

TRP: Transient receptor potential

WB: Western blot

Wnt: Wingless/INT

Wt: Wild type


CONTENTS

Abstract………………………………………………………………………………………………………………………………………………………………………………………………. 5 Abstract in Danish – dansk resumé…………………………………………………………………………………………………………………………………………………….. 6

Chapter 1 Thesis objectives………………………………………………………………………………………………………………………………………………………………. 7 1.1 Introductory remarks……………………………………………………………………………………………………………………………………………………………. 7 1.2 Thesis primary objective……………………………………………………………………………………………………………………………………………………….. 7 1.3 Thesis secondary objective……………………………………………………………………………………………………………………………………………………. 8 Chapter 2 Introduction and Background…………………………………………………………………………………………………………………………….................... 9 2.1 Ciliary structures and functions………………………………………………………………………………………………………………………………..……………. 9 2.1.1 Ciliary assembly and maintenance……………………………………………………………………………………………………………………….………… 10 2.1.2 Introduction to ciliary signaling pathways and ciliopathies.…………………………………………………………………..………………………... 11 2.1.3 Hedgehog signaling and primary cilia in developmental processes…………………………………………………………………………...…….. 13 2.2 Stem cells……………………………………………………………………………………………………………………………………………………………………………… 15 2.2.1 Embryonic stem cells…………………………………………………………………………………………………………………………………………………….. 17 2.2.2 Embryonal carcinoma (EC) cells…………………………………………………………………………………………………………………………………….. 17 2.3 Heart development……………………………………………………………………………………………………………………………………………………………….. 18 2.3.1 Heart fields and developmental stages…………………………………………………………………………………………………………………………… 20 2.4 Signaling pathways in heart development………………………………………………………………………………………………………………………………. 21 Chapter 3 – Primary cilia in stem cell differentiation and cardiogenesis………………………………………………………………………………………….. 23 3.1 Introductory remarks……………………………………………………………………………………………………………………………………………………………. 23 3.2 Primary cilia with functional Hh signaling in human embryonic stem cells………………………………………………………………………………. 23 3.3 Primary cilia and Hh signaling in stem cell differentiation and cardiogenesis………………………………………………………………………....... 23 3.4 Heart development studied in Chimera mice………………………………………………………………………………………………………………………….. 25 3.5 Primary objective conclusions and perspectives…………………………………………………………………………………………………………………….. 27 Chapter 4 – Sensory cilia in the pancreas and reproductive organs………………………………………………………………………………………………….. 30 4.1 Introductory remarks……………………………………………………………………………………………………………………………………………………………. 30 4.2 Hedgehog signaling in pancreatic development and cancer…………………………………………………………………………………………………….. 30 4.2.1 Primary cilia and Hedgehog signaling in pancreatic development …………………………………………………………………………………... 30 4.2.2 Primary cilia and Hedgehog signaling in pancreatic cancer……………………………………………………………………………………………... 31 4.3 Sensory motile cilia in the oviduct………………………………………………………………………………………………………………………………………….. 32 4.4 Secondary objective conclusions and perspectives…………………………………………………………………………………………………………………. 33 Chapter 5 – Thesis conclusions…………………………………………………………………………………………………………………………………………………………… 34 5.1 Thesis conclusions……………………………………………………………………………………………………………………………………………………………....... 34

Chapter 6 – References…………………………………………………………………………………………………………………………………………........................................... 35 Chapter 7 – Articles [15]……………………………………………………………………………………………………………………………………............................................. 45


ABSTRACT

This thesis focuses on cilia and their sensory function in the mammalian organism. In particular, the Hedgehog (Hh) signaling pathway functions via the primary cilium and plays a unique role in development, differentiation, cancer and possibly in stem cell fate. Defects in primary cilia assembly or function are tightly coupled to developmental disorders and diseases in mammals termed “ciliopathies”.

The primary objective of this thesis was to investigate the role the primary cilium in coordinating Hh signaling in stem cell differentiation and heart development in the mouse. We show that human embryonic stem cells (hESC) and mouse embryonal carcinoma stem cells (P19.CL6 EC cells) have primary cilia that display ciliary localization of the essential Hh proteins; Gli2, Ptc1 and Smo. Inhibition of the Hh pathway by KAAD‐cyclopamine in P19.CL6 cells hinder formation of synchronously beating clusters of cardiomyocytes. Knockdown of the primary cilium in P19.CL6 EC cells by nucleofection with plasmids expressing Ift20 and Ift88 siRNA significantly reduced the appearance of beating cardiomyocyte clusters thereby mimicking the effect of cyclopamine treatment. In vivo experiments revealed that mouse E11.5 Ift88‐/‐ null mutants (which have no primary cilia) have severe endocardial cushion defects, decreased trabeculation and increased pericardial space along with shortened and malformed cardiac outflow tract. These observations suggest that primary cilia coordinate Hh signaling in stem cell differentiation and cardiogenesis. In support of this, preliminary chimera mouse studies showed that primary cilia are important for heart development. This was judged by the distribution of enzymatically tagged wt and Ift88‐/‐ ES cells in the developing heart at E8.5, where only the wt cells are localized to the heart chambers. This signifies that primary cilia are needed for the formation of the heart chambers.

The secondary thesis objective was to investigate the role of progesterone signaling in the female reproduction organs in addition to the role of primary cilia in human pancreatic development and cancer. The findings of the progesterone receptor in the lower half of the motile cilia in the oviduct, suggest a sensory role of motile cilia in progesterone signaling where they might coordinate post ovulatory events. In tissue sections of the developing human pancreas we found up to 20µm long primary cilia projecting into the duct lumen of the exocrine duct, which have increased ciliary localization of Gli2 and Smo after initiation of fetal development, i.e., at weeks 14 and 18. In contrast, ciliary localization of these Hh components was absent at the embryonic stage of development, i.e., at week 7.5. This suggests a role of primary cilia in coordinating Hh signaling in human pancreatic development and postnatal tissue homeostasis. In cultures of human pancreatic duct adenocarcinoma cell lines PANC‐1 and CFPAC‐1, Ptc in addition to Gli2 and Smo localize to primary cilia. These findings are consistent with the idea that the primary cilium continues to coordinate Hh signaling in cells derived from the mature pancreas. The fact that the Hh signaling pathway is active in the CFPAC‐1 and PANC‐1 cell lines without Hh stimulation suggests that ciliary Hh signaling plays a potential role in tumorigenesis.

In conclusion, this thesis supports the idea that both motile and primary cilia are critical organelles in the coordination of developmental processes and tissue function.


ABSTRACT IN DANISH – DANSK RESUME

Denne PhD afhandling har fokus på cilier og deres sensoriske funktion hos pattedyr. Mere specifikt spiller Hedgehog (Hh) signaleringsvejen, der virker via det primære cilium, en afgørende rolle for udvikling, differentiation, cancer og muligvis også stamcelle vedligeholdelse. Defekt ciliedannelse eller ciliefunktion er tæt knyttet til udviklingsdefekter og sygdomme der betegnes ”ciliopatier”.

Hovedopgaven i denne afhandling var at undersøge rollen af det primære cilium i koordineringen af Hh signaling under stamcelledifferentiering og hjerteudviklingen hos mus. Vi viser her, at humane embryonale stamceller (hESC) samt museembryonale carcinoma‐stamceller (P19.CL6) danner primære cilier, hvortil essentielle Hh proteiner, Gli2, Ptc1 og Smo lokaliserer. Inhibering af Hh signalvejen med KAAD‐cyclopamine i P19.CL6 celler, hæmmer dannelsen af synkront bankende minihjerter. Knockdown af det primære cilium i P19.CL6 celler ved nukleofektion med plasmider, der udtrykker Ift20 og Ift88 siRNA, reducerer signifikant dannelsen af bankende minihjerter og efterligner effekten af cyclopaminbehandlingen. In vivo forsøg viste at E11.5 Ift88‐/‐ mutant mus (der ikke har primære cilier) har alvorlige endokardiale pudedefekter, underudviklet trabekulering, udvidet perikardial hulrum samt forkortet og misdannet hjerteudløbstrakt. Disse resultater foreslår en mulig rolle for det primære cilie i koordineringen af Hh‐signaleringsvejen under stamcelledifferentiering og hjerteudviklingen. Ydermere støtter præliminære forsøg med chimeramus hypotesen, at det primære cilium er vigtigt for hjerteudviklingen. Dette er bedømt ud fra distribueringen af enzymatisk mærkede vildtype og Ift88‐/‐ ES celler, hvor vildtypecellerne lokaliserer til hjertekamrene, som stort set er fri for mutant ES celler. Dette betyder at det primære cilium er nødvendig for dannelsen af hjertekamrene.

Den sekundære opgave i denne afhandling var at undersøge rollen af progestosteronsignalering i de kvindelige reproduktionssorganer samt rollen af det primære cilium i udviklingen af human pankreas og cancer. Tilstedeværelsen af progestosteronreceptoren i den nedre halvdel af de bevægelige cilier i æggelederne indikerer en mulig sensorisk rolle for de bevægelige cilier i progestosteronsignaleringen som koordinator for postovulatoriske begivenheder. I undersøgelserne med den humane pankreas, fandt vi, at den udviklende eksokrine dukt danner primære cilier på op til 20µm, der projicerer ud i duktlumen fra duktepithelet både under embryonal (uge 7.5) og føtal udvikling (uger 14 og 18). Analyserne med vævssnittene viste endvidere, at niveauet af Hh‐komponenterne, Gli2 og Smo, kraftigt stiger under den føtale udvikling og er fraværende under den embryonale udvikling. Denne forøgelse i ciliær Hh signalering foreslår en rolle for det primære cilie i koordineringen af Hh signalering i modning af det eksokrine duktsystem. Ydermere ses der i kulturer af humane pankreasdukt adenocarcinoma cellelinier, PANC‐1 og CFPAC‐1, at både Ptc, Smo og Gli2 lokaliserer kraftigt til de primære cilier og at Hh signalering er kraftigt opreguleret i cancercellerne i fravær af Hh‐stimulering. Disse resultater tyder på, at ciliær Hh‐signalering kan spille en rolle i tumordannelse.

Afhandlingens resultater støtter konklusionen, at både motile og primære cilier spiller en afgørende rolle som et sensorisk organel under udvikling og i vævsfunktion.


CHAPTER 1 ‐ THESIS OBJECTIVES

1.1 INTRODUCTORY REMARKS

Primary cilia are solitary organelles which are organized in a 9+0 microtubule axonemal ultra structure that project from the centrosomal mother centriole at the surface of stem cells and most growth‐arrested cells in our body (Satir & Christensen, 2008). Primary cilia are sensory organelles that coordinate a series of signal transduction pathways to control developmental processes, tissue homeostasis and behavioral responses (Singla & Reiter, 2006; Satir & Christensen., 2008; Berbari et al., 2009). The physiological importance of primary cilia is underscored by an ever‐growing list of diseases and developmental disorders (‘ciliopathies’) associated with defective primary cilia, e.g. cystic kidney and liver diseases, retinal degeneration, abnormalities in neural tube closure and patterning, heart defects, skeletal and left‐right patterning defects, hydrocephalus, obesity and cancer (Kuehn et al., 2007; Kennedy et al., 2007; Mans et al., 2008; Michaud & Yoder, 2006; Plotnikova et al., 2008; Wong et al., 2009; Han et al. 2009; Slough et al., 2008; reviewed in; Davenport and Yoder, 2005; Christensen et al., 2008; Pan J, 2008; Lehman et al., 2008; Berbari et al., 2009; Veland et al., 2009). Although some of the overall pathways are known, our understanding of the detailed mechanisms by which the cilium controls cell organization and function is still rudimentary.

This thesis gives novel insights into the function of primary cilia in stem cell differentiation and in coordinating the complex events taking place in the early heart development. The work was carried out in part by investigating the role of the primary cilium in coordinating Hedgehog (Hh) signaling and in promoting the differentiation of mouse embryonal carcinoma (EC) stem cells (P19.CL6) into beating cardiomyocytes, and in part by investigating defects in heart development in Ift88‐/‐ mouse embryos, which have defects in ciliary assembly. Further, preliminary data obtained with chimera mice studies with Ift88‐/‐ stem cells support the thesis hypothesis that primary cilia are critical in heart development. As a second objective, this thesis also presents novel findings on the role of primary cilia in coordinating Hh signaling in human pancreatic development and postnatal tissue homeostasis as well as the potential role in progesterone signaling in motile cilia of the human and mouse oviduct in coordinating post ovulatory events. A more detailed description of my thesis objectives is listed in chapters 1.2 and 1.3.

Chapter 2 of this thesis is an introduction to primary cilia, ciliary signaling mechanisms in health and disease, the murine heart physiology, heart development, stem cells, female reproductive organs and the pancreas, which serve as background for the chapters 3 and 4 that discuss the primary and secondary objectives in the thesis respectively. The introduction contains unpublished data and observations made during the work period.

1.2 THESIS PRIMARY OBJECTIVE

Heart development, which includes the formation of the cardiac crescent, linear heart tube, heart looping, chamber formation and septation/maturation of the young heart, is regulated by a series of various signaling pathways that mediate or interact with progenitor cells to expand, migrate, differentiate and ultimately integrate into the forming heart. The signaling pathways in heart development include Hedgehog (Hh), Wingless/INT (Wnt), fibroblast growth factor (FGF), transforming growth factor (TGF)‐beta, platelet‐derived growth factor (PDGF) and bone morphogenic protein (BMP) signaling.


The main objective of this thesis was to investigate the role the primary cilium in coordinating Hh signaling in stem cell differentiation and heart development in the mouse. Initially, primary cilia were characterized by immunofluorescence microscopy (IFM) and electron microscopy (EM) analysis in cultures of human embryonic stem cells (hESC), and we show that primary cilia are associated with regulation of Hh signaling in these cells [3]. The focus of my thesis was then to investigate the function of the primary cilium in coordinating Hh signaling and differentiation of P19.CL6 EC stem cells into beating clusters of cardiomyocytes. In this work, a full characterization of how P19.CL6 EC stem cells in cultures differentiate under normal conditions was carried out before experiments on ciliary knockdown and inhibitory chemicals on signaling pathways could be tested. This included studies on heart transcription factors, stem cell markers and morphological analysis on the clustering cardiomyocytes using western blot (WB), quantitative real time‐PCR (Q‐RT‐PCR) and immunofluorescence microscopy (IFM) analysis. Electroporation with plasmids expressing siRNAs against Ift88 and Ift20 was used to knock down key proteins in ciliogenesis in order to analyze the significance of the primary cilium in early heart development in vitro. Furthermore, the role of primary cilia in heart development was analyzed in vivo. This was carried out partly by investigating heart defects in Ift88‐/‐ mice that lack or have severely stunted primary cilia, and partly by studying the development of the heart in chimera mice with injected wild type (wt) and Ift88‐/‐ mouse embryonic stem cells (mESC) in order to clarify whether primary cilia are required for heart development. In parallel to the P19.CL6 cardiomyocyte differentiation experiments, a few preliminary studies on Ift88‐/‐ and wt ES cell differentiation into cardiomyocytes were conducted to get a broader perspective on the role of the primary cilium in cardiomyogenesis.

List of the cell types and animals used in the primary thesis objective:

• Human embryonic stem cells (hESC) (Chapter 7: [3]).

• Mouse embryonal carcinoma (mEC) stem cells (P19.CL6) (Chapter 7: [45]).

• Ift88‐/‐ and wt mouse embryonic stem cells (mESC) (preliminary data, not shown).

• E11.5 wt and Ift88‐/‐ mouse embryos (Chapter 7: [4]).

• Mouse Chimera with Ift88‐/‐ and wt ES injects (preliminary data shown in Chapter 3: section 3.4).

1.3 THESIS SECONDARY OBJECTIVE

The secondary objective of my thesis emerged because of collaboration with two students, Sonja K. Brorsen and Stefan C. Teilmann who worked on the sensory function of cilia in the pancreas and in the female reproductive organs in the laboratories of Drs. Søren T. Christensen, Kjeld Møllgård and Anne Grete Byskov. The work with Sonja K. Brorsen focused on the function of primary cilia in Hh signaling during development of the exocrine duct of the human pancreas and how aberrant Hh signaling may be associated with primary cilia in pancreatic cancer. This work was carried out partly by performing IFM analysis on tissue sections of the developing human pancreas and partly by IFM and WB analysis of human pancreatic duct adenocarcinoma cell lines. The work with Stefan C. Teilmann included IFM and WB analysis on the expression and localization of progesterone receptors in human and mouse female reproductive organs with special focus on changes in the localization of the receptors to motile cilia of the oviduct upon ovulation in the mouse. This work would help understand how post ovulatory responses are coordinated in the oviduct and how motile cilia could play a part of this regulation.

List of the cell types and tissues used in the thesis secondary objectives and related articles:

• Human pancreatic duct adenocarcinoma PANC‐1 and CFPAC‐1 cell lines and NIH3T3 cells (Chapter 7: [2]).

• Tissue sections from 7.5‐week‐old human embryos and 14‐ and 18‐week‐old human fetuses [2].

• Tissue sections from human and mouse oviduct and ovary (Chapter 7: [1]).


Figure 1. Structure and localization of motile and primary cilia. A: Schematic illustration of motile 9+2 cilia and primary 9+0 cilia. Motile cilia have inner and outer dynein arms, radial spokes, nexin links and a central microtubule (MT) pair surrounded by the central sheath. Primary cilia only have the 9 outer microtubule doublets with the nexin links to stabilize the structure. The MT doublets are composed of A and B sub fibers. B: Images of 9+2 motile cilia in: Tetrahymena thermophila (arrows: cilia, acetylated tubulin, tb: green, C. A. Clement, unpubl.), section of human oviduct (arrows: cilia, tb: red, progesterone receptor: green, DAPI: blue, C. A. Clement, unpubl.), rat brain section of the ventricles tissue (arrows: cilia, tb: red, DAPI: blue, C. A. Clement, unpubl.), DIC image of an isolated mouse oviduct cell (arrows: cilia, Teilmann et al.,2006) and DIC/IFM image of human adult oviduct (arrows: cilia, progesterone receptor: green, propidium iodide: red , [1]). C: Images of 9+0 primary cilia in: mouse NIH3t3 fibroblasts DIC/IFM (arrows: primary cilia, tb: green, C. A. Clement, unpubl.), hESC SEM image (arrow: primary cilia, [2]), mESC (arrows: primary cilia, tb: red, ARL13b: green, DAPI: blue (insert shifted overlay), C. A. Clement, unpubl.) and mouse embryonal carcinoma cells (arrow: primary cilia, Ift88: green, tb: red, DAPI: blue, C. A. Clement, unpubl.).

CHAPTER 2 ‐ INTRODUCTION AND BACKGROUND

2.1 CILIARY STRUCTURES AND FUNCTIONS

Cilia are membrane‐bounded, centriole‐derived projections from the cell surface that contain a microtubule (MT) cytoskeleton, the ciliary axoneme, surrounded by a ciliary membrane (Satir and Christensen, 2007) (Figure 1). The microtubule cytoskeleton of the cilium, the axoneme, grows from and continues the nine fold symmetry of the centriole that becomes a ciliary basal body. Ciliary axonemes are formed with two major patterns: 9+2, in which the nine doublet microtubules surround a central pair of singlet microtubules, and 9+0 cilia, in which the central pair is missing. Most often, 9+2 cilia are motile; motility being regulated by axonemal inner and outer arm dyneins that coordinate ciliary beat frequency and form, respectively (Brokaw & Kamiya, 1987; Satir 1998; Christensen et al., 2001). In contrast, 9+0 cilia usually lack axonemal dynein arms and are consequently non‐

motile (Satir and Christensen, 2008). Nodal cilia, like primary cilia also possess 9+0 axonemes, but nodal cilia have dynein arms with LR (left/right) dynein (Supp et al., 1999) and are motile, generating a leftward flow across the node required for establishment of the left–right asymmetry axis (Hirokawa et al., 2006). Motile 9+2 cilia are found in a wide range of organisms spanning from single celled organisms to humans, in which they are found lining the airway epithelium (Jeffery & Reid, 1975), the brain ventricles (Cathcart & Worthington, 1964), the ependyma/choroid plexus (Wodarczyk et al., 2009) and the oviduct epithelium (Boisvieux‐Ulrich et al., 1989). In ciliates, e.g. Tetrahymena termophila, the motile cilia are primarily used for swimming and to collect food particles from the surrounding environment. Mammalian 9+2 motile cilia may also function as a sensory organelles (Christensen et al., 2007) such as in the oviduct (Teilmann & Christensen, 2005; [1]) and in the airway epithelium,


where motile cilia possess sensory bitter taste receptors (Shah et al., 2009). In contrast to motile 9+2 cilia, primary cilia are solitary organelles that project from the centrosomal mother centriole at the surface of stem cells and most growth‐arrested cells in our body (Satir and Christensen, 2007) (Figure 2). Further, primary cilia lack radial spokes and the central sheath surrounding the central microtubule pair in 9+2 motile cilia (Satir and Christensen, 2008). As will be discussed in the below, primary cilia are thought to function as unique mechano‐, osmo‐ and chemosensory organelles, which enable the cells to interact and communicate with the surrounding environment via the primary cilium to control cell cycle entry, migration and differentiation during development and in tissue homeostasis.

Figure 2. The figure shows examples of mammalian tissues and cell types that have a primary cilium. For a more thorough list see the reference. http://www.bowserlab.org/primarycilia/cilialist.html

2.1.1 CILIARY ASSEMBLY AND MAINTENANCE Both primary and motile cilia are assembled and maintained via a highly conserved process called intraflagellar transport (IFT) (reviewed in; Rosenbaum & Witman, 2002; Pedersen & Rosenbaum, 2008) first discovered in the green algae Chlamydomonas reinhardtii by Kozminski and co‐workers in 1993 (Kozminski et al., 1993). Transmembrane proteins as well as axonemal components are transported in vesicles via the Golgi‐complex along microtubules using the cytoplasmic dynein 1 motor to the base of the cilium (see figure 3). It is proposed that Ift20 (a complex B particle) and GMAP210 (a golgin anchor protein) function at the Golgi‐complex to sort proteins into vesicles destined for the cilium, where Ift20 reside in the vesicles and GMAP210 stays in the Golgi‐complex. At the base of the cilium, Ift20 on the vesicles interacts with the Ift54 (a subunit of IFT complex B) to form the complete IFT complex (Follit et al., 2009) however knockdown of the Ift20 gene reduces ciliary assembly without affecting Golgi structure (Follit et al., 2006). Ift20 was shown to coordinate Wnt signaling and cell proliferation required for proper positioning of the centrosome in non‐dividing cells and for correct orientation of the mitotic spindle in kidney collecting duct epithelium cells (Jonassen et al., 2008).


The complete IFT complexes with the cargo destined for the cilium rendezvous at the base of the cilium where they connect with ciliary motor proteins. At the “ciliary necklace”, only proteins (or protein complexes) with a ciliary targeting motif can enter the zone or “pore complex” created by the transition fibers (Gilula & Satir, 1972; Rosenbaum & Witman, 2002). The anterograde transport of protein/IFT complexes is mediated by kinesin‐II along the ciliary axoneme to the ciliary tip along with inactive cytoplasmic dynein 2. In addition to kinesin‐2, motor proteins belonging to other kinesin families may contribute to ciliary structural and functional diversity (reviewed in; Scholey, 2008). At the ciliary tip, turnover products are switched over to cytoplasmic dynein 2 for retrograde IFT transport back to the basal body region to re‐enter the cytosol. Ift88, a subunit of the IFT particle complex B, is required for both anterograde and retrograde IFT (Pazour et al., 2000; Murcia et al., 2000; Haycraft et al., 2001; Taulman et al., 2001; Yoder et al., 2002b; Lucker et al., 2005). Kif3 motors (comprising of Kif3a and Kif3b subunits) are a functionally diverse subgroup of the kinesin super family, characterized by an NH2‐terminal motor domain (N‐IV class) and forms a complex with the non‐motor protein KAP3. Together they are responsible for MT‐based anterograde transport to membranous organelles including cilia (Yamazaki et al., 1995; Hirokawa, 1998; Haraquchi et al., 2006). A way to stop ciliogenesis and thereby ciliary functions is by using knockout or knockdown of IFT particles. Two well‐known IFT particles that have been used to disrupt ciliary assembly, includes Ift88 (Pazour et al., 2000; Murcia et al., 2000; Haycraft et al., 2001; Taulman et al., 2001; Yoder et al., 2002b; Lucker et al., 2005; Schneider et al., 2005) and Ift20 (Follit et al., 2006; 2008; 2009) that leave no or severely stunted cilia.

2.1.2 INTRODUCTION TO CILIARY SIGNALING PATHWAYS AND CILIOPATHIES The ciliary membrane consists of a bilayer lipid membrane that is continuous with the plasma membrane of the cell body, but which contains a different complement of membrane receptors and ion channels. As outlined in the above it is now evident that primary cilia play a major role in coordinating a series of signal transduction pathways in cell cycle entry, migratory responses and differential processes. These pathways include Hedgehog (Hh), Wingless/INT (Wnt), neuronal and purinergic receptors as well as signaling through the transient receptor potential (TRP) ion channels, receptor tyrosine kinases (RTK) and extracellular matrix communication ([2];

Figure 3. The mechanism of intraflagellar transport (IFT). Proteins and ciliary components destined for the ciliary compartment are transported in Golgi‐derived vesicles (containing both transmembrane and axonemal proteins) along microtubules to the base of the cilium with Ift20 particle interactions and cytoplasmic dynein 1, where the vesicles are exocytosed and the ciliary proteins associate with ciliary IFT particles (e.g. Ift88). Proteins are sorted at the transition zone by the transition fibers and transported anterogradely along the axoneme by kinesin‐II‐mediated IFT. At the ciliary tip IFT particles are remodeled, kinesin‐II is inactivated and cytoplasmic dynein 2 takes over the retrograde transport back to the basal body region. Abbreviations: MT, microtubule; PCM, pericentriolar material. Figure based on references (Rosenbaum & Witman, 2002; Pedersen & Rosenbaum, 2008, Chapter two in “Ciliary function in mammalian development”).


Christensen et al., 2007; Christensen et al., 2008; Eggenschwiler & Anderson, 2007; Gerdes et al., 2009; Knight et al., 2009; Masyuk et al., 2008; Praetorius & Leipziger, 2009; Wong & Reiter, 2008; Jensen et al., 2004; Veland et al., 2009) (see also Figure 4). As examples on RTK signaling, the primary cilium in fibroblasts uniquely coordinates PDGF‐Rαα signaling to regulate cell cycle entry, wound healing events and regeneration (Schneider et al., 2005; 2009a; 2009b), and Insulin‐like Growth Factor 1 (IGF‐1) receptors localize to the primary cilium and its basal body in 3T3‐L1 preadipocytes to regulate adipocyte differentiation (Zhu et al., 2009). In the adult, primary cilia may also control behavioral responses. Hormone receptors like somatostatin Sst 3 receptor (Sst3R) localize to primary in the hypothalamus (Handel et al., 1999) and melanin‐concentrating hormone receptor 1 (Mchr1) localize to neuronal primary cilia (Berbari et al., 2008). The Sst3R and Mchr1 receptors are mal‐localized in mice with mutations in proteins that correspond to those from patients that suffer from the obesity condition Bardet‐Biedl syndrome which also involve leptin receptor signaling (Berbari et al., 2008; Seo et al., 2009). These observations link primary cilia signaling to feeding behavior and the way we sense hunger.

Since primary cilia are critical in regulation of signaling pathways in behavioral responses and cellular processes during development and in tissue homeostasis, lack of normal functioning primary cilia causes various diseases now commonly known as ciliopathies. These include cystic kidney and liver diseases, retinal degeneration, abnormalities in neural tube closure and patterning, heart defects, skeletal and LR patterning defects, hydrocephalus, obesity and cancer (Kuehn et al., 2007; Kennedy et al., 2007; Mans et al., 2008; Michaud & Yoder, 2006; Plotnikova et al., 2008; Wong et al., 2009; Han et al., 2009; Slough et al., 2008; reviewed in; Davenport & Yoder, 2005; Christensen et al., 2008; Pan, 2008; Lehman et al., 2008; Berbari et al., 2009; Veland et al., 2009).

One of the first diseases to be related to dysfunctional primary cilia, was polycystic kidney disease (PKD) found in mice mutated in the gene encoding the Ift88/Tg737/Polaris protein in the Oak Ridge Polycystic Kidney mouse (ORPK mouse, or currently designated Ift88Tg737Rpw), (Moyer et al., 1994; Pazour et al., 2002; Yoder et al., 2002a; Pazour et al., 2004; Lehman et al., 2008). In Chlamydomonas, Ift88 mutants showed defective ciliogenesis, and it was established that cilia of the mouse kidney were also abnormally short or missing, which suggested that PKD might be a ciliary disease (Pazour et al., 2000). The Tg737ORPK mouse was induced by insertional mutagenesis integrated into an intron near the 3´ end of the Tg737 gene thereby partially disrupting the expression and function of the Ift88 protein. The hypomorhpic allele of Ift88 in the ORPK mouse makes this mouse a good model to study the role of primary cilia since the animals are viable into young adulthood compared to the Ift88‐/‐ null

Figure 4. Listed examples and images of ciliary signaling pathways. Top list summarizes up different signaling pathways that are coordinated by primary cilia. Images from left to right: A merged IFM image on sections of the nephron in the toad Xenopus (arrows: primary cilia, acetylated tubulin: red, Polycystin‐2: green, DAPI: blue (insert shifted overlay), C. A. Clement, unpublished data). A merged IFM image of mouse NIH3T3 fibroblasts in culture (arrows: primary cilia, acetylated tubulin: red, Polycystin‐2: green, DAPI: blue, C. A. Clement, unpublished data). A merged IFM image of mouse embryonal carcinoma stem cells in culture (arrows: primary cilia, acetylated tubulin: red, Gli2: green, DAPI: blue, C. A. Clement, unpublished data). A merged IFM image of mouse NIH3T3 fibroblasts in culture (arrows: primary cilia, PDGFRα: red, acetylated tubulin: green, DAPI: blue, Schneider et al., 2005).


Figure 5. Listed examples of ciliopathies and syndromes associated with ciliary defects. List on the left summarizes up different diseases and syndromes that have been observed to associate with defects in primary cilia. Top image on the right show a healthy and a polycystic kidney. The polycystic kidney is ~five times the size of the normal kidney and is non‐functional due to the fluid filled cysts. Bottom image on the right show the polydactyly phenotype in a newborn human child. The infant’s foot has six toes.

mice (Ift88tm1Rpw, Ift88tm1.1Bky, and Ift88fxo), which are embryonic lethal around the beginning of organogenesis (Lehman et al., 2008). The core phenotypes of the Tg737ORPK mouse was originally described as a triad of the following; scruffy fur, severe growth retardation, and preaxial polydactyly on all limbs (Moyer et al., 1994). The Tg737ORPK mouse revealed another very significant phenotype which became the best known phenotype, the cystic renal phenotype which resembles that of human autosomal recessive polycystic kidney disease, which is characterized by extensive cystic enlargement of both kidneys that fail to concentrate the urine (see figure 5). This experimental mouse was also the first mammalian model to establish a connection between ciliary dysfunction and cystic kidney disease (Pazour et al., 2000; 2002; Taulman et al., 2001). Loss of cilia function in

the Tg737ORPK mice also revealed additional pheno‐types such as hepatic and pancreatic ductal abnor‐malities and cysts, retinal degeneration, skeletal de‐fects, cerebellar hypo‐plasia, hydrocephalus, respiratory defects, infertility, situs inversus and heart defects (Moyer et al., 1994; Pazour et al., 2002a; Cano et al., 2004; Banizs et al., 2005; Zhang et al., 2005; Chizhikov et al., 2007; Haycraft et al., 2007; Hildebrandt & Otto, 2005; Hildebrandt & Zhou, 2007).

Since primary cilia are involved in a wide range of signaling pathways controlling and coordinating cellular responses new ciliopathies are frequently added to the list. In embryogenesis, signaling through the primary cilium is necessary for normal development in e.g. PDGF‐R and Hh signaling pathways, probably because such signaling regulates the balance between cell division, polarity, migration, differentiation and apoptosis for many tissues (Schneider et al., 2005; Rohatgi et al., 2007; reviewed; Singla & Reiter, 2006; Michaud & Yoder, 2006; Christensen et al., 2007; Christensen & Ott, 2007). More specifically in cell migration, the primary cilium was proposed to function as a cellular GPS that orients towards the leading edge of the cell and in parallel to the migration path (Christensen et al., 2008). In terms of PDGF‐Rαα signaling, PDGF‐Rα is translocated to the cilium where activation of the receptor by homodimerization with its specific ligand, PDGF‐AA, induces the activation of the Mek1/2‐Erk1/2 and Akt pathways in the cilium or centrosomal region to control changes in cytoskeletal proteins partly via activation of the Na+/H+ exchanger, NHE1, at the leading edge (Schneider et al., 2005; 2009a; 2009b). In Ift88‐/‐ null fibroblasts without primary cilia, chemotaxis towards PDGF‐AA is blocked, leaving the cells blindfolded to coordinate their migration in early wound healing in vivo.

2.1.3 HEDGEHOG SIGNALING AND PRIMARY CILIA IN DEVELOPMENTAL PROCESSES In mammals, Hh signaling is induced by three different ligands, Indian (Ihh), Desert (Dhh) and Sonic hedgehog (Shh). The Hh signaling pathway controls and maintains many steps in development and several studies have revealed that dysfunctional Hh signaling results in a wide range of developmental disorders (reviewed in; Wong & Reiter, 2008; Simpson et al., 2009; Veland et al., 2009). Some examples where Hedgehog signaling is important for proper development are in LR asymmetry (Tsukui et al., 1999), skeletogenesis and digit patterning in the limbs (Johnson et al., 1994; Gouttenoire et al., 2007; Haycraft et al., 2007; Bastida et al., 2009), neural tube


formation (Gorivodsky et al., 2008), cerebellar development (Chizhikov et al., 2007; Spassky et al., 2008), mammary gland development, ovarian function (Johnson et al., 2008) and development of the lung (Bellusci et al., 1997; Rutter et al., 2009), the heart (Washington Smoak et al., 2005; [4]) and the pancreas ([3]; Bailey et al., 2009). Besides coordinating development, Hh signaling plays a pivotal role in cancer formation and generation of tumors. Indirect activation of Hh signaling in a subset of epithelial cancers; e.g. colon, pancreatic, and ovarian cancer can promote tumor growth by activating Hh signaling in the surrounding stroma, which then provides a more favorable environment for the developing tumors. This is why the Hh signaling pathways is a therapeutic target in cancer where manipulation of the Hh pathway potentially can delay or cure cancers. The Hh pathway is already being used in therapy and preclinical studies in addition to clinical trials, which are underway in a range of malignancies (reviewed in; Theunissen & Sauvage et al., 2009; O’Toole et al., 2009). The primary cilium is associated with regulation of Hh signaling and is also present on human tumors e.g. in basal cell carcinomas (BCCs) which are frequently ciliated. Removal of the primary cilium in these tumors strongly inhibited BCC‐like tumors induced by an activated form of Smoothened. On the other hand, removal of cilia accelerated tumors induced by activated Gli2. Somehow, there is a dual role for primary cilia controlling Hh signaling which can then either mediate or suppress tumorigenesis depending on the oncogenic initiating event (Wong et al., 2009).

The general mechanism by which Hh works in vertebrates, is by the binding of the Hh ligand to the transmembrane receptor patched (Ptc) which thereby abolishes the inhibitory effect of Ptc on Smoothened (Smo), a seven‐transmembrane receptor. Complete loss of Ptc activity turns the Hh pathway fully on even in the absence of Hh ligands (Ingham & McMahon, 2001). Following the loss of Ptc activity, Smo is able to transduce a signal via Gli transcription factors to the nucleus that initiate expression of Hh target genes. The activity of Smo is essential for any response either to Hh or to loss of Ptc activity, which indicate that Smo acts downstream of Ptc (reviewed by; Kalderon, 2005; Varjosalo & Taipale, 2008). There exist three Gli transcription factors, Gli1‐3, where Gli1 functions as a constitutive activator (Hynes et al., 1997; Ruiz I Altaba, 1999; Liu et al., 2005). In contrast, Gli2 and Gli3 have an N‐terminal transcriptional repressor domain and a C‐terminal transcription activator domain. The proteolytic events that switch between the activating and repressing form of Gli2 and Gli3 are controlled by Smo (Huangfu et al., 2006; Pan et al., 2006). Hh signaling plays a critical role in establishing the LR asymmetry axis and proper heart tube looping during gastrulation, as well as maintaining the adult coronary vasculature and survival of small coronary arteries and capillaries (Lavine et al., 2008). The LR axis is initiated at the Hensen´s node of the mouse at E7.75 where two populations of nodal cilia coexist (McGrath et al., 2003); 1) the first are motile cilia with a mixture of 9+2 and 9+0 cilia containing the outer arm dyneins, called left–right dynein (lrd), which generate a left‐ward fluid flow at the node (Supp et al., 1997; Caspary et al., 2007; review; Basu & Brueckner, 2008), 2) the second are non‐motile cilia with a 9+0 microtubule architecture that are located around the edge of the node, which functions as mechano sensors and/or chemo sensors via the cation channel polycystin‐2 in the ciliary membrane. The non‐motile cilia respond to the nodal flow generated by the motile cilia which initiate a Ca2+ response in the cells at the left border of the node (McGrath et al., 2003). Within the Hensen´s node, LR asymmetry is initiated by asymmetric expression of activinβB that inhibits Shh expression in the right portion of the node and thereby allowing its expression in the left. Here it diffuses into the adjacent lateral plate mesoderm and induces Nodal expression (Wagner & Siddiqui, 2007). Consequently, Shh mutants show severe effects on cell survival in the pharyngeal arch and neural crest, in addition to reduced size of the right ventricle (RV) and out flow tract (OFT) and delayed Nkx2.5 expression and heart development, thus suggesting direct effects of Shh on the second heart field (SHF) specification, proliferation or deployment (Zhang et al., 2001).

The primary cilium has been proposed to act as a key regulator of Hh signaling (Kovacs et al., 2008; for reviews; Eggenschwiler & Anderson, 2007; Christensen & Ott, 2007; Wong & Reiter, 2008). In many cell types the essential Hh signaling components Gli2, Gli3, and Smo localize to the primary cilium and transports actively together with the IFT complexes, e.g. in fibroblasts (Haycraft et al., 2005; Rohatgi et al., 2007; 2009), epithelial cells in renal tubules (Harris & Torres, 2008) and the exocrine duct of the pancreas [2] as well as in human embryonic stem cells ([3]; Breunig et al., 2008). In these cells the Smo and Ptc was found to enter and leave the


cilium upon stimulation with Hh ligand, see figure 6. The binding of ligands to Ptc in the cilium may activate the Hh pathway by removal of Ptc from the ciliary compartment and in that process, allowing Smo to enter the cilium and hereby coordinating the proteolytic events of the Gli2 and Gli3 transcription factors (Rohatgi et al., 2007; Wong & Reiter, 2008). In these events it has been proposed that the primary cilium may function as a switch by which the cells can regulate Hh signaling during development and tissue homeostasis (Corbit et al., 2005; Rohatgi et al., 2007). Suppressor of fused (Sufu) is a major negative regulator of Hh signaling in vertebrates (Cooper et al., 2005; Svard et al., 2006) and is taking part in the regulation of protein levels of full‐length Gli transcription factors. Sufu has been found to localize to the primary cilium and in the nucleus/cytosol (Haycraft et al., 2005), where it directly interact with the Gli transcription factors (Dunaeva et al., 2003). A possible hypothesis was that Sufu could regulate Gli proteolysis and generation of activator forms in the cilium in coordination with Smo, but recent data suggest that the regulation of Gli protein levels by Sufu is cilium‐

independent. The generation of Gli activator forms might still be a cilia dependent process that is regulated by a Smo mediated mechanism, but where Sufu controls ubiquitination of Gli proteins via the speckle‐type POZ protein, Spop. This is a new role of mammalian Sufu in controlling Gli protein stability that is important for understanding ciliary Hh signaling and how it is regulated (Jia et al., 2009; Chen et al., 2009; Ruel & Thérond, 2009).

2.2 STEM CELLS

Stem cell research is a very important field of study with the purpose of gathering information on how to use stem cells as a therapeutic tool in regenerative medicine and as a model of human development. Stem cell transplantations are seen as a possible cure for Alzheimer’s disease, cancer, neurodegenerative disorders and in regeneration of the heart in patients with myocardial infarction, which is characterized by irreversible loss of cardiomyocytes leading to heart failure (Guillaume & Zhang, 2008; Song et al., 2009). The use of hESCs in differentiation experiments in vitro will help identifying new gene targets for drugs in tissue regeneration therapies. However, many key elements in stem cell signaling and differentiation are still not known and will need to be clarified before a wide spread use of stem cells can be trusted in regenerative medicine. The Geron Corporation is the first company in the world given clearance (Jan. 23rd ‐2009) for clinical trials on humans with

Figure 6. Ciliary Hh signaling mechanisms. The binding of Hh to Ptc1 in the cilium abolishes inhibition of Smo. Smo enters the cilium in contrast to Ptc1 leaving the cilium for degradation in the cytoplasm. With Smo active in the cilium it has been proposed that it may coordinate the proteolytic events that favor the Gli2 and Gli3 full length activator forms. The Gli2‐3A then translocate to the nucleus and initiate transcription of Hh response genes (Ptc1 and Gli1). In mammals, Smo is thought to inhibit Suppressor of Fused (Sufu), a negative regulator of the Hh pathway, leading to activation of target‐genes through the Gli transcription factors.


hESC derived cells, where spinal cord injuries can be treated with oligodendrocyte progenitor cells injected into the lesion.

Stem cells are found in most multi‐cellular organisms and are characterized by the ability to self‐renew through mitotic cell division and differentiate into any cell type (Smith, 2001). Embryonic stem cells (ESC) are pluripotent, which mean that they have the capacity to generate derivatives of all the three embryonic germ layers: the ectoderm, mesoderm and endoderm. The ectoderm contribute to the central nervous system, the lens of the eye, the ganglia and nerves, pigment cells, head connective tissues, the epidermis, hair, and mammary glands. The mesoderm forms skeletal muscle, bone, connective tissue, the heart, blood, and the spleen. The endoderm forms the gut and lung tissue, the liver, pancreas, the urinary bladder, the thyroid and more (Chandros et al., 2001). Pluripotent stem cells occupy the inner cell mass of the early blastocyst during embryonic development (Lensch, 2009), see figure 7.

The internationally recognized gene markers to characterize hESCs for determining if the cells are in an undifferentiated state are: NANOG, TDGF, POU5F1, GABRB3, GDF3 and DNMT3B. No hESC lines reported of today have tested negative for these six markers (Adewumi et al., 2007), provided by the International Stem Cell Initiative, ISCI. In mESC three important transcription factors have been identified for regulating pluripotency, namely Oct4, Sox2 and Nanog. All of these transcription factors are highly expressed in the inner cell mass, epiblast and in undifferentiated mESC (Pesce & Scholer, 2001; Niwa, 2007). Null mutations of each of the three genes cause early embryonic lethality due to the inability to maintain cells in a pluripotent state (Nichols et al., 1998; Avilion et al., 2003; Mitsui et al., 2003). Oct4 by itself, induces differentiation of ES cells through Cdx2 and eomesodermin if the expression of Oct4 is reduced by 50% (Niwa et al., 2000; 2005), and Sox2 RNAi silencing results in ES cell differentiation into multiple linages (Ivanova et al., 2006). Sox2 can also interact synergistically with Oct4 in vitro to activate Oct–Sox enhancers, which in turn can regulate Nanog, Oct4 and Sox2 themselves (Masui et al., 2007). It is therefore important that Nanog, Oct4 and Sox2 are closely regulated since changes in their expression rates have dire consequences for controlling stem cell pluripotency and developmental processes (Niwa et al., 2000). Recent work has shown that stem cells possess primary cilia with signaling molecules and receptors that may be critical for stem cell renewal and differentiation ([3]; Awan et al., 2009). The following sections 2.2.1 and 2.2.2 describes in more detail the features of stem cells in developmental research.

Figure 7. Schematic of stem cell origin. After the fusion of the sperm and oocyte, a morula is formed. The cells in the morula are totipotent (meaning they are omnipotent and able to develop into a complete viable organism including a placenta). The morula develops into a blastocyst where the inner cell mass contain pluripotent embryonic stem cells. Pluripotent cells can differentiate into nearly all cell types e.g. cells derived from all the three germ layers. Unipotent cells can only produce their own cell type but have the ability to self‐renew, which distinguishes them from non‐stem cells (http://en.wikipedia.org/wiki/Stem_cell).


2.2.1 EMBRYONIC STEM CELLS It took 17 years after the first isolation of mouse ES cells before James Thomson and co‐workers derived hESC from donated blastocysts from couples undergoing treatment for infertility (Thomson et al., 1998). The method used was almost the same as was used for isolating mESC. The trophectoderm (trophoblast, group of cells that produces no embryonic structures) was removed by immunosurgery, where the inner cell mass (ICM) was plated onto mouse embryonic fibroblasts to act as a feeder layer. Several groups had tried this approach but the culture media from the mESC protocol resulted in differentiation and not the derivation of stable pluripotent cell lines (Bongso et al., 1994). Some experiments with ES cell lines from two non‐human primates: the rhesus monkey and the common marmoset (Thomson et al., 1995; 1996), gave the necessary experience to adjust the culture conditions to produce stable undifferentiated human pluripotent ES cells. mESC are different in many aspects compared to primate ES cells, particularly in their morphology and their ability to withstand dissociation into single cells (Pera et al., 1999). Human ESC are karyotypically normal and have the capability to maintain the developmental potential to contribute to all of the three germ layers even after extended undifferentiated proliferation (Amit et al., 2000). After the first successful attempt to isolate stable hESCs (Thomson et al., 1998), others derived them from the morula and the blastocyst stage embryos (Stojkovic et al., 2004; Strelchenko et al., 2004), followed later by isolation from single blastomeres (Klimanskaya et al., 2006), and parthenogenetic embryos (unfertilized human eggs), (Lin et al., 2007; Mai et al., 2007; Revazova et al., 2007). It is still not known whether pluripotent cell lines derived from these various sources have any consistent developmental differences or whether they have an equivalent potential (Yu & Thomson, 2008).

Pluripotent stem cells are not present at all times in the developing embryo, since they rapidly differentiate into more specialized somatic cells. The first mESC lines were extracted from the ICM of a mouse blastocyst and then cultured on a mitotically inactivated fibroblast feeder layer with serum. These culturing conditions were adapted from the mESC cultures in vitro (Evans & Kaufman, 1981; Martin, 1981). ES cell cultures that are clonally derived from a single ES cell could then differentiate into a wide variety of cell types in vitro and form teratocarcinomas when injected into mice (Martin, 1981). In contrast to mouse embryonal carcinoma cells (mEC), mESC can differentiate into a variety of tissues in chimeras at high frequency, including germ cells, which give the possibility of introducing modifications to the mouse germ line (Bradley et al., 1984; Robertson, 1986). To culture mESCs several methods have been used. One is to culture the ES cells on feeder layers as described above; another is to culture them in conditioned media that are able to sustain the ES cells without growing them on feeder cells. This led to the identification of leukemia inhibitory factor (LIF), a cytokine that is a key factor to sustain ES cells in an undifferentiated state (Smith et al., 1988; Williams et al., 1988).

2.2.2 EMBRYONAL CARCINOMA (EC) CELLS Embryonal carcinoma (EC) cells comprise a special class of tumor cells which have the ability to change phenotype from malignant into non‐malignant via cellular differentiation. EC cells are derived from teratocarcinomas which is where the field of pluripotent stem cells arose from in the 1950s. Teratocarcinomas are found in the testes of mice and humans that occur from defective germ cells (Stevens & Little, 1954; van der Heyden et al., 2003). In 1964, Kleinsmith and Pierce showed that single EC cells are capable of self‐renewal and differentiation into multiple cell lineages and hereby establishing the existence of pluripotent stem cells (Kleinsmith & Pierce, 1964). This provided the intellectual basis for more advanced studies of both mouse and human ES cells and was also the first experimental demonstration of a cancer stem cell (Yu & Thomson, 2008). In the 1970s, stable mouse EC cell lines could be cultured in vitro and used for studies in development that could not be carried out with intact mammalian embryos (Kahan & Ephrussi, 1970). On the other hand, most EC cell lines have limited developmental potential and contribute poorly to chimera mice studies (see section 3.4), properly due to the accumulation of genetic changes during teratocarcinomas formation and growth (Atkin et al., 1974). But still mouse EC cells, compared to human EC cells, are more useful because the human EC cells are highly aneuploid (have abnormal number of chromosomes), which might explain why they can’t differentiate


into a wide range of somatic cell types, which limits the use for studies of human development in vitro (Yu & Thomson, 2008; Kennedy et al., 2009).

The P19 cell line, a murine EC cell, is an undifferentiated stem cell that originates from a teratocarcinoma (Martin, 1980). As a stem cell, it is able to differentiate into all three germ layers by culturing the cells in suspension with several chemical inducers. With addition of high concentrations of retinoic acid (RA, 0.1‐1µM), the cells can differentiate into neurons and glia (McBurney et al., 1982) or with low concentrations of RA (1‐10nM) or dimethyl sulfoxide (DMSO) (0.5‐1%) the cells can differentiate into cardiac and skeletal myocytes (McBurney et al., 1982; Edwards et al., 1982). Because of the multipotential abilities of P19 cells, this cell line is an often used model system to study early heart differentiation in vitro. To improve on the P19 cells ability to differentiate into cardiomyocytes, a clonal line was isolated from the P19 cells, called CL6 (Habara‐Ohkubo, 1996). This P19.CL6 sub line efficiently differentiates into cardiac muscle with the addition of 1% DMSO in adherent culture (Habara‐Ohkubo, 1996). Unlike the P19 cells that depend on aggregate formation in suspension (Campione‐Piccardo, 1985), the CL6 cells can be cultured without aggregation and feeder cells. How the CL6 cells effectively differentiate into beating muscle in adherent rather than suspension culture is unclear, but aggregate structures that resemble embryoid bodies are observed during the multilayer sheet formation during the differentiation into cardiomyocytes (Habara‐Ohkubo, 1996). Although CL6 cells are thought to be morphologically similar to P19 cells, only CL6 cells express the mesodermal marker gene Brachyury but not the stage‐specific embryonic antigen‐1, SSEA‐1, which is a cell surface embryonic antigen whose loss of expression characterizes the differentiation of murine EC cells (Habara‐Ohkubo, 1996; Uchida et al., 2007). Moreover, CL6 cells differentiate into neurons at a much lower frequency than P19 cells, which is why it was suggested that the CL6 cells are not committed to the mesoderm but represent a developmental stage closer to differentiated cardiomyocytes than P19 cells (Habara‐Ohkubo, 1996). Further, P19.CL6 cells are quite sturdy and are easily electroporated in siRNA knockdown experiments. For these reasons, we used cultures of the P19.CL6 EC cell line to study the role of the primary cilium in cardiogenesis.

2.3 HEART DEVELOPMENT

The heart is among the most studied of all organs but also the one most susceptible to disease. Early heart development in vertebrates is a complex process initiated in embryos shortly after gastrulation, where cardiomyocyte progenitor cells aggregate and become allocated from the mesodermal population which migrate and organize into the cardiac crescent. Hereafter the cardiac crescent will develop into a beating linear heart tube, the first functional organ of the developing embryo, as a result of migration and fusion along the ventral midline of the precursor cells from the cardiac crescent (Sucov, 1998; Nemer, 2008). The heart is not only composed of muscle cells but also contain a wide range of cell types such as atrial/ventricular cardiac myocytes, conduction system cells, smooth muscle/endothelial cells of the coronary arteries and veins, valvular components, endocardial cells and connective tissue (Laugwitz et al., 2008). Three major sources of heart cell precursors have been identified in the mouse embryo: the cardiogenic mesoderm, the proepicardial organ and the cardiac neural crest. These three sources represent a distinct pool of progenitor cells where the cardiogenic mesoderm gives rise to the linear heart tube and the myocardium in the ventricular and atrial chambers, see figure 8. The proepicardial organ and the cardiac neural crest gives rise to the epicardial mantle, which later contributes to the coronary vasculature and to the vascular smooth muscle of the aortic arch, ductus arteriosus respectively (Laugwitz et al., 2008). It is critical that regulation of these different cell progenitors is under the strict control so that the correct cell lineages differentiate at the correct time and in the correct location (Bruneau, 2008). Many signal transduction systems are implicated as essential coordinators of early cardiogenesis, including Hh, Wnt, BMP, FGF, PDGF and (TGF)‐beta signaling pathways (Washington Smoak et al., 2005; Kwon et al., 2008; Hirata et al., 2007; Rochais et al., 2009; van Wijk et al., 2007). These signaling pathways


control multiple genes that are expressed throughout the cardiomyocyte population prior to the fusion of the linear heart tube and remain expressed hereafter.

Figure 8. Schematic diagram illustrating the origin and lineage relationships of cardiac cell types in mouse development. A: Three sources contribute in heart development with progenitor cells during cardiac morphogenesis in the mouse: the cardiogenic mesoderm (red), the cardiac neural crest (CNC, purple) and the proepicardial organ (yellow). Early in development at E7.5, progenitor cells of the cardiogenic mesoderm are recognizable under the head folds (HFs) of the embryo, which then move ventrally to the midline (ML) and form the linear heart tube and finally the four chambers of the heart. After the looping of the heart tube at E8.5, cardiac neural crest progenitors migrate from the dorsal neural tube at E10.5 to engulf the aortic arch (AA) arteries and further contribute to the vascular smooth muscle cells of the outflow tract (OFT). Simultaneously the proepicardial organ precursors contact the surface of the developing heart and give rise to the epicardial mantle (yellow arena around the heart at E10.5) and contribute later to the coronary vasculature. Abbreviations: PhA, pharyngeal arches; PLA, primitive left atrium; PRA, primitive right atrium; LV, left ventricle; RV, right ventricle. B: A display of cardiac cell types that arise through the lineage differentiation of the three embryonic precursor pools. The contribution of the proepicardium to the smooth muscle cells of the coronary system and to the mesenchymal cells of the heart is well accepted, the origin of the endothelial lineage in the coronary vasculature is still controversial. Modified from Laugwitz et al., 2008.

Congenital heart disease (CHD) is a common infant morbidity and arises from abnormal heart development during embryogenesis. CHD is reported to have 6 to 8 incidences per 1000 live births and approximately accounts for 3% of all infant deaths and 46% of deaths from congenital malformations. Further, cyanotic heart defects (a group‐type of CHD where the patient appears blue “cyanotic”, due to deoxygenated blood bypassing the lungs and entering the systemic circulation) occur in about 60 per 100,000 live births in the United States. Cyanotic heart defects can be caused by right‐to‐left or bidirectional shunting, or mal‐positioning of the great arteries. Also, the frequency of CHD in premature infants is 12.5 per 1000 live births (Sadowski, 2009). Stem cell regeneration of cardiac tissue may be a therapeutic tool in the future to save a large number of patients suffering from myocardial infarction. In order to transform stem cell therapy from idea to clinical use a lot of basic


knowledge is needed on how the heart signaling pathways interact, coordinate, initiate and maintain the developing/adult heart.

2.3.1 HEART FIELDS AND DEVELOPMENTAL STAGES The cardiac crescent originates from cells in the cardiogenic mesoderm and is one of the earliest steps in cardiogenesis. The cardiogenic mesoderm consists of two populations or heart fields of cardiac precursor cells that contribute to different parts of the heart. The first heart field (FHF, the earliest), is located in the anterior splanchnic mesoderm, which primarily gives rise to the cardiac crescent, as well as to the linear heart tube and to parts of the atrial chambers and the left ventricular region later in development. The second heart field (SHF, also known as the anterior heart field) lies anterior and dorsal to the linear heart tube and is derived from the pharyngeal mesoderm medial to the cardiac crescent. Cells from this second heart lineage are added to the developing heart tube and give rise to the outflow tract, the right ventricular region and the main parts of the atrial tissue, see figure 9 (reviewed by Buckingham et al., 2005; Laugwitz et al., 2008).

Figure 9. Schematic diagram illustrating the early steps in heart development and with key transcription factors activation points at the different stages. The diagrams of the heart development are shown in ventral views. At the earliest stages of heart formation (cardiac crescent), two pools of cardiac precursors exist. The first heart field (FHF, in pinkish colour) contributes to the LV, and the second heart field (SHF, in bluish colour) contributes to the right ventricle (RV) and later to the outflow tract (OT), sinus venosus (SV), and left and right atria (LA and RA, respectively). Abbreviations: V, ventricle; A, Atria; PA pulmonary artery; Ao, Aorta. Bottom half of the diagram show when the transcription factors are turned on. Figure modified from Bruneau, 2008 and Nemer, 2008.

The myocardium was thought to be derived from a single source of cells until recently. The identification of a second source of myocardial cells that contribute to the cardiac chambers has modified the classical view of heart formation. The SHF was first discovered in the chick (Mjaatvedt et al., 2001; Waldo et al., 2001; reviewed in Buckingham et al., 2005) and then in mouse (Kelly et al., 2001). In the mouse it was shown that a second source of myocardial cells in the pharyngeal mesoderm contributes to the outflow‐tract myocardium at the arterial pole of the heart. These cells initially lie medially to the cardiac crescent before assuming a position that is dorsal and anterior to the heart tube (Kelly et al., 2001). Later by using linage‐tracing CRE‐LOXP recombination system experiments, results showed that the heart tube derived from the FHF may predominantly provide a scaffold upon which cells from the SHF migrate to and build the requisite cardiac chambers at the later stages in heart development (Meihac et al., 2004; Brown et al., 2004; Xu et al., 2004). The SHF is further subdivided into a number of lineage pools (Buckingham et al., 2005), which contribute either to anterior structures (such as the


OFT) or posterior components (such as the atria). These findings may explain how mutations associated with CHD, by only affecting specific cell lineages within the SHF result in defects in specific heart structures (Bruneau, 2008).

The initial steps to build a fully functional four chambered heart starts with ventral movement of cells from the cardiac crescent which combine into a linear heart tube (DeHaan, 1965) that consist of an interior layer of endocardial cells and an exterior layer of myocardial cells. At the linear heart tube stage, the heartbeat is initiated (Srivastava, 2006) and transcription factors initiate distinct segmental precursors of the OFT, atria, and ventricles (Srivastava & Olson, 2000). The heart tube continuously grows by division of myocardial cells and by the addition of cells to both poles of the heart (Buckingham et al., 2005). Around E9 in mouse development (heart looping stage), the outflow region swings to the right as the heart adopts a spiral form, which realigns the future ventricles into a left‐right juxtaposition. The inflow portion of the heart moves in an anterior and dorsal direction such that the inflow and outflow complexes converge. The crude heart then undergoes considerable remodelling where the most highly proliferative cardiomyocytes are located along the outer surface of the heart, also termed the compact myocardium, which then thickens and becomes the myocardial wall (Sedmera & McQuinn, 2008). On the inside the cardiomyocytes organizes into trabeculae, a sponge‐like layer of myocytes and finger like projections thought to enhance oxygen and nutrient exchange in the absence of a coronary circulation (Franco et al., 2006). Polarised growth of myocardial cells forms in a highly defined region, called the interventricular septum (IVS), which will divide the ventricles, encompassing the junction of future left and right ventricles. The sinuatrial (SA) and atrioventricular (AVC) nodes form within slow‐conducting myogenic tissue of the inflow tract where the SA node becomes the cardiac pacemaker (Nanot & Douarin, 1977). In the AVC, endocardial cushions are the precursors of the tricuspid and mitral valves, while in the OFT they form a scaffold for the aorticopulmonary septum which divides the OFT into the aorta and pulmonary artery and forms the aortic and pulmonary valves. During the growth process of the cardiac epithelium another distinct cell lineage, the migrating cardiac neural crest cells, populate the heart through the outflow channel and contribute to the septation of the OFT into distinct vessels of the aortic and pulmonary arteries (reviewed in; Hutson & Kirby, 2007; Bruneau, 2008).

2.4 SIGNALING PATHWAYS IN HEART DEVELOPMENT

Hedgehog, Wnt, BMP, FGF, PDGFR and (TGF)‐beta signaling coordinate heart development in part by activating essential early heart genes such as GATA binding protein 4 (Gata4), NK2 transcription factor related, locus 5 (Nkx25), myocyte enhancer factor 2C (Mef2C), cardiac actin, and desmin (Lyons, 1994). The Gata family transcription factors are zinc‐finger proteins that play important roles in heart formation, e.g. in cardiac muscle and heart tube development at the ventral midline (Grepin et al., 1994; Kuo et al., 1997). In vertebrates, three Gata genes exist (Gata46), which are expressed in the heart (Molkentin, 2000; Molkentin et al., 2000a). Gata4‐/‐ null mice embryos have bilateral heart tubes (cardia bifida) and a reduced number of cardiac myocytes (Kuo et al., 1997; Molkentin et al., 1997), where Gata5‐/‐ null mutants are viable (Molkentin et al., 2000b). Never the less, homozygous Gata4‐/‐ null embryonic stem cells are able to differentiate into contractile myocytes in chimeric embryos, which suggests that the cardia bifida phenotype is related to an endoderm deficiency (Narita et al., 1997). Heart development studies in vitro show that Gata4 expression precedes that of Nkx2‐5 which is also one of the earlier heart transcription factors, which are important for proper heart development and cardiomyocyte differentiation (Grepin et al., 1997). Nkx2‐5 is a transcription factor with a homeobox domain, which is highly expressed in cells of both the FHF and SHF (Stanley et al., 2002) and continuously during cardiac development throughout adulthood (Lints et al., 1993). In mice, Nkx2‐5 is required for terminal differentiation of cardiac myocytes and the expression is clearly crucial for the normal growth of the embryonic myocardium, which is visible in the poorly developed myocardium of mice lacking Nkx2‐5. These mice do not grow beyond the earliest


stages of heart looping (Lyons et al., 1995; Tanaka et al., 1999) and show left ventricular and conduction system defects (Yamagishi et al., 2002; Jay et al., 2004). In cardiomyocyte differentiation, the Myocyte enhancer factor 2c (Mef2c) act as a cofactor for Gata proteins (Morin et al., 2000) during the cardiac, skeletal, and smooth muscle development (Skerjanc et al., 1998).

Positive inducers of cardiogenesis are BMP, FGF, Shh and Wnt‐JNK (also known as the Wnt‐polarity pathway, Wnt11), which are expressed in the mesoderm, endoderm and ectoderm. Inhibitory signals include Wnt ligands expressed in dorsal neural tube (Wnt‐3a and Wnt‐8c) via β‐catenin, and anti‐BMPs expressed in the axial tissues e.g. Noggin in the notochord (reviewed in; Brand, 2003; Wagner & Siddiqui, 2007), see figure 10. Collectively these positive and negative signals drive mesodermal cells to the cardiogenic cell lineage, presumably by inducing the expression of cardiogenic transcription factor genes (Wagner & Siddiqui, 2007; Rochais et al., 2009).

Figure 10. Overview of the signaling pathways implicated in cardiogenic induction. Endoderm‐derived signals are labeled in green: BMP2, FGF8, Crescent, and Shh/Ihh. These molecules act as inducers of cardiac mesoderm formation (they induce activation of cardiogenic transcription factors, such as Nkx2‐5, Gata factors, myocardin, T‐box (Tbx2, ‐3, ‐20) and Mef2c). Mesoderm‐derived signals are labeled in red: Chordin, Noggin, Wnt1, ‐3, ‐8, and Serrate which all are inhibitory except Wnt11. Wnt11 is a potent cardiac inducer that is present in mesoderm. The neural ectoderm is a source of inhibitors of heart field formation. Labeled in white are the signal transducers of the particular signaling pathways that have a function in coordinating cardiogenesis (modified from; Brand, 2003).


CHAPTER 3 – PRIMARY CILIA IN STEM CELL DIFFERENTIATION AND CARDIOGENESIS


Without a doubt, intensive research in the last decade has revealed that the primary cilium plays a critical role in a wide range of developmental processes in mammals (reviewed in; Lehmann et al., 2008; Satir & Christensen, 2008; Veland et al., 2009, Berbari et al., 2009). This thesis presents new data that support the conclusion that primary cilia are critical organelles in heart development and stem cell fate. This chapter will discuss and summarize the novel data from the primary objective articles and will round up with some conclusions. In addition, some new preliminary data will be presented and taken into consideration in the Discussion. The articles for the primary objective are found in chapter 7 in the following sections: Collaborative work with Aashir Awan on hESCs and ciliary Hh signaling (Chapter 7: [3]) and Heart development in P19.CL6 cells and cardiomyocyte differentiation (Chapter 7: [45]).

3.2 PRIMARY CILIA WITH FUNCTIONAL HH SIGNALING IN HUMAN EMBRYONIC STEM CELLS

In our paper [3] we demonstrate for the first time that hESC in cultures form primary cilia with the characteristic 9+0 axoneme as evidenced by transmission electron microscopy (TEM), scanning electron microscopy (SEM) and IFM analysis [3]. Further, we show that key components in Hh signaling, including Smo, Ptc and Gli2 localize to hESC primary cilia, and that stimulation with the Smo agonist, SAG, promotes the concerted movement of patched out of, and smoothened into, the primary cilium, in accordance with the hypothesis that primary cilium functions as a cellular switch in turning the Hh signaling pathway on and off (Christensen & Ott, 2007). In addition, SAG promotes the increased expression of Ptc1 and Gli1, which are the two immediate response genes upon Hh pathway activation [3]. These results support the conclusion that primary cilia are involved in the regulation and coordination of the first steps of hESC differentiation, and/or the maintenance of the undifferentiated state/self‐renewal. Since hESCs hold promise for the treatment of many diseases and provide an excellent system for studying mechanisms involved in early human development, these findings provide the groundwork to determine specific aspects of early differentiation controlled by the machinery of primary cilia. This knowledge may ultimately reveal pathways for manipulation of hESC differentiation into specific cell and tissue lineages.

3.3 PRIMARY CILIA AND HH SIGNALING IN STEM CELL DIFFERENTIATION AND CARDIOGENESIS

Recently, Slough et al., (2008) showed that the embryo heart at E9.5 Kif3a‐/‐ mice have abnormal heart development, indicating that primary cilia could coordinate processes in cardiac morphogenesis. However, since Kif3 family proteins regulate cellular processes in mammalian cells that are not necessarily related to the primary cilium (Teng et al., 2005; Haraguchi et al., 2006; Corbit et al., 2008), there was a need for a more


Figure 11. Immunofluorescence microscopy images showing P19.CL6 cells on day 13 in their differentiation. The four images show the localization of anti‐Gata4 (green), anti‐Beta3‐tubulin (red), DAPI (blue). Inside the cluster of cardiomyocytes are two neurons visualized with the neuron‐specific class III beta‐tubulin, C. A. Clement, unpublished data.

thorough investigation on the role of the primary cilium in early cardiogenesis and Hh signaling, which is critical in cardiomyocyte differentiation. In our paper [4] we demonstrate that primary cilia play a critical role in coordinating Hh signaling and cardiomyogenesis in P19.CL6 EC cells. Further, we show that E11.5 old embryos of the Ift88tm1Rpw (Ift88‐/‐ null) mice, which form no cilia, have many different heart defects, supporting the conclusion that cardiac primary cilia are critical in early heart development, partly via coordination of Hh signaling.

To sum up on the mouse P19.CL6 EC differentiation studies [4], we showed that P19.CL6 stem cells form primary cilia, which have ciliary Hh components such as Ptc1, Smo and Gli2. This is the first discovery of primary cilia in this cell line. The mouse P19.CL6 EC cell line is of pluripotent lineage as evidenced by expression of the stem cell markers Sox2 and Oct4. Moreover, inhibition of the Hh pathway by KAAD‐cyclopamine blocked DMSO‐induced differentiation of P19.CL6 cells into beating clusters of cardiomyocytes by restraining the expression and nuclear localization of the heart transcription factors Gata4 and Nkx2‐5. In addition, KAAD‐cyclopamine inhibited Hh signaling in P19.CL6 cells, confirmed by a failed up‐regulation of Gli1 and Ptc1 mRNA expression and nuclear localization of Gli1, Gli2 and Gli3. The Gli2‐repressor protein levels was increased in the KAAD‐cyclopamine treated cells in contrast to the DMSO‐induced control cells, which had higher levels of full length Gli2 that may function as the activator form. These results suggest that Hh signaling is required for differentiation of P19.CL6 cells into cardiomyocytes, and that Hh signaling may be associated with primary cilia in P19.CL6 EC cells. However, recent data showed that treatment with KAAD‐cyclopamine during aggregation in P19 EC cells does not inhibit the general up regulation of Gata4, BMP4, Brachyury T, Meox1 and Gli2 during cardiomyogenesis, but merely delays it (Gianakopoulos & Skerjanc, 2009). This interesting difference might be explained by the culturing conditions by aggregation in the P19 cells, which is supposed to initiate mesoderm induction. Somehow, P19 cells manage to initiate the early cardiomyogenesis in absence of Hh signaling, possibly by Wnt signaling pathways that may compensate and activate Gli2. This has been observed in P19 cells where Wnt3a induces skeletal myogenesis in aggregated P19 cells, which is then followed by up‐regulation of Gli2 (Petropoulos & Skerjanc, 2002). The Wnt/β‐catenin pathway is critical in regulation of cardiogenesis where precise timing is an important factor to coordinate specific cellular responses. Studies in mouse and zebrafish embryos, as well as in embryonic stem cells clearly demonstrate that the Wnt/b‐catenin pathway plays distinct,

even opposing, roles during various stages of cardiac development (Tzahor, 2007). Whether the P19.CL6 cells have already undergone mesoderm induction compared to the P19 cells is not known. However, the P19.CL6 cells are a sub clone from the original P19 cells and should be more prone to cardiomyocyte differentiation. Even though the P19.CL6 cells primarily differentiate into cardiomyocytes, they are not only a mesoderm driven cell line since they can also produce neurons, which are ectoderm derived cells, see figure 11.

To answer the question whether primary cilia are important in Hh signaling for driving P19.CL6 cells down the cardiogenic pathway, we used the nucleofector technique described in detail in chapter 7, article


[5]. The primary cilium was knocked down by Ift88 and Ift20 siRNA which both are IFT complex B proteins required for functional IFT and delivery of ciliary membrane proteins from the Golgi complex to the cilium respectively (Pazour et al., 2000; Lucker et al., 2005; Follit et al., 2006; 2009). Knockdown of Ift88 in P19.CL6 cells resulted in a reduced frequency of ciliated cells to about 30% and inhibited the expression levels of Gata4 and Nkx2‐5 to about 40% of mock controls. Consequently the number of beating cardiomyocytes was reduced as well as the nuclear localization of Gata4 at day 12. The Ift88 siRNA transfected cells were Sox2 positive, indicating that knock down of the cilium maintains cells in their undifferentiated state which do not undergo apoptosis or differentiate into other cell lineages. Using a combination of both Ift88 and Ift20 siRNA we further reduced the number of ciliated cells and resulted in an additional decrease in the number of beating clusters of cardiomyocytes along with a more pronounced decrease in mRNA expression levels of Gata4 and Nkx2‐5 and protein levels of Gata4, Nkx2‐5 and α‐actinin. The knockdown of Ift88 and Ift20 in P19.CL6 cells mimics the inhibitory response on Ptc1 and Gli1 expression levels as seen with KAAD‐cyclopamine treatment at day 5 of differentiation. These data lead to the hypothesis that differentiation of P19.CL6 cells into cardiomyocytes is coordinated by the primary cilium and partly by regulation of Hh signaling. To test if this hypothesis was correct we performed microscopy analysis on the Ift88‐/‐ mice. These mice display at E11.5 severe endocardial cushion defects, decreased trabeculation and increased pericardial space along with malformations of the OFT. Many of these phenotypes are observed in the Pkd2‐/‐ and Kif3a‐/‐ mice but not in the lrd‐/‐ embryos, which indicate that the malformations of the heart are not due to defective left‐right asymmetry, which is coordinated by the nodal cilia (Slough et al., 2008). The OFT malformations in our Ift88‐/‐ embryos was not observed in Slough et al., (2008), possibly due to the time difference in embryonic development since they sacked the mice at E9.5 compared to our E11.5, or the sheer difference of the Kif3a‐/‐ versus Ift88‐/‐ cilia derived mice. Gli2 localizes to both primary cilia in the developing heart of wt embryos and in P19.CL6 cells, as in contradiction to Ift88‐/‐ embryos where this localization is disrupted. Interestingly, the OFT phenotype of Ift88‐/‐ embryos resembles the phenotype of Shh‐/‐ mice embryos (Washington Smoak et al., 2005; Goddeeris et al., 2007), suggesting that primary cilia mediate Hh signaling responsible for correct OFT development and potentially in development of the other cardiac structures that are malformed in the Ift88‐/‐ embryos. These results support the hypothesis, that the primary cilium is critical for proper development of the mammalian heart.

3.4 HEART DEVELOPMENT STUDIED IN CHIMERA MICE

The first embryonic mouse chimera was generated by aggregating two eight‐celled embryos (Tarkowski, 1961). The result was a normal‐sized mouse with tissue that was a mix of cells from both embryos. Chimera mice can also be generated by injecting foreign pluripotent ES cells into a mouse blastocyst, which allows the ES cells to differentiate into all tissue types. Homologous recombination of ES cells (Doetschman et al., 1988; Thomas & Capecchi, 1989) is a powerful tool to engineer and generate designer chimera mice that are mutated in genes of interest, where the effect of genetic changes can be analyzed (Tam & Rossant, 2003). Today, chimera embryos can be generated by injecting enzymatically tagged wt and mutant ES cells into a wt blastocyst. In order to investigate the function of the primary cilium in heart development in more detail, we performed analysis on chimera mice in further collaboration with Professors Bradley K. Yoder and Robert Kesterson from the University of Alabama at Birmingham, USA. Embryos were harvested at the preferred time points in mouse development and stained to visualize where mutant or wt ES cells contribute in specific tissue/organ of the chimera embryo. Figure 13 shows an example of enzymatically tagged wt and mutant Ift88‐/‐ mouse ES cells that are cultured on feeder fibroblasts. Cultures like these can be trypsinized and separated into single cells followed by differential attachment culturing to separate the ES cells from the feeders. The single ES cells can then be collected and injected into mouse blastocysts. To assess the in vivo significance and relevance of the in vitro P19.CL6 data, we conducted chimera embryo studies generated by the method described above. The chimera mice were created by using the mouse ES cells shown in figure 12, which were injected into blastocysts. The mouse embryos were harvested at various time points in embryonic development and stained for wt and Ift88‐/‐


ES cell contribution. To distinguish the contribution of the wt from the mutant Ift88‐/‐ mESCs, the β‐galactosidase gene reporter was incorporated into the wt cells (which stains cells blue), while the bacterial alkaline phosphatase reporter was used for the Ift88‐/‐ mESC (which stain cilia mutant cells red). With this approach, tissues where primary cilia function is required should reveal no contribution of Ift88‐/‐ cilia mutant cells. The distribution of cells was determined using whole mount embryos in addition to paraffin and cryofreeze sections of the embryos.

Figure 12. Light microscope images of enzymatically tagged mouse ES cells growing on fibroblast feeders and of E8.5 chimera mouse embryos. Top row, from left to right: wt mouse ES (f/f‐zap)colonies that contain a LacZ gene reporter that express the β‐galactosidase enzyme that is thereby able to cleave X‐gal substrate and consequently stain wt cells blue (inserted image: IFM with DAPI and anti‐hnn showing a primary cilia: arrow). Hnn is an allele of Arl13b, a small GTPase of the Arf/Arl family, and the Arl13b protein predominantly localizes to cilia (Caspary et al., 2007). Embryo showing only wt ES cell contribution after β‐galactosidase and alkaline phosphatase assay with 8‐10 injected cells of both wt (blue) and Ift88‐/‐ (red) mESC. The arrows points at the ventricle region of the early heart. Bottom row from left to right: mutant Ift88‐/‐ mESC colonies that contain a bacterial alkaline phosphatase gene reporter that stain Ift88‐/‐ cells red in response to Ift88 deletion (inserted image: Immunofluorescence microscopy (IFM) with DAPI and anti‐Hennin (hnn) showing missing primary cilia). Embryo showing only mutant Ift88‐/‐ ES cell contribution after β‐galactosidase and alkaline phosphatase assay with 8‐10 injected cells of wt and Ift88‐/‐ mESC. The arrows points at the ventricle region of the early heart. C. A. Clement, unpublished data made in collaboration with Bradley K, Yoder, Nickolas Berbari and Robert Kesterson).

In these preliminary studies, Ift88‐/‐ ESCs (red) fail to contribute to the development of the heart chambers, in contrast to differentiated wt ESCs (blue), which clearly localize to the heart region (figure 13); wt contribution is additionally seen in other parts of the developing embryo such as the brain. On the other hand, the mutant Ift88‐/‐ ESC contribution localize to the tissue surrounding the heart chambers (body wall), whereas the internal regions such as the out flow tract and atria are practically free of mutant Ift88‐/‐ ESC. Although conducted on a small number of embryos at this point, the results support our hypothesis that primary cilia are important for cardiogenesis.


Figure 13. Chimera embryo A: Light microscope image of E8.5 chimera mouse embryo after β‐galactosidase and alkaline phosphatase assay with 8‐10 injected cells of both wt (blue) and Ift88‐/‐ (red) mESCs, (dotted line: heart region). B: Light microscope image of a frontal cut 10µm cryofreeze section of the mouse embryo in viewed in (A): Abbreviations; A: atrium, OFT: outflow tract, B: future brain, BW: body wall, closed arrow: alk. phos. staining of the tissue surrounding the heart, open arrow: LacZ staining of the heart chambers. C. A. Clement, unpublished data.

3.5 PRIMARY OBJECTIVE CONCLUSIONS AND PERSPECTIVES

At several points in cardiomyogenic development multiple signaling pathways and their downstream effecter molecules crosstalk and overlap. This complicates the matter in understanding the specific mechanisms that determines when, where and how stem cells initiate differentiation, migration and proliferation in early heart development. In figure 14, I have given a brief overview of the signaling pathways in cardiogenesis to describe where there could be possible crosstalk between the individual pathways in heart development. Some examples of signaling pathways that overlap are the Hh and Wnt pathway, which were proposed to be coordinated by the primary cilium. The essential Hh signaling components Gli2, Gli3, and Smo localize to the primary cilium, in various cell types including fibroblasts (Haycraft et al., 2005; Rohatgi et al., 2007), where Wnt signaling is divided up into three distinct pathways that has been proposed to work as a network of interacting rather than individual pathways (Kestler & Kuhl, 2008). The primary cilium and basal body have been proposed to act as regulator in both the non‐canonical and canonical Wnt pathways due to the ciliary/basal body localization of essential proteins like PCP (planar cell polarity) protein inversin (Morgan et al., 2002), Vangl‐2 (Ross et al., 2005) in addition to members of the degradation complex Glycogen synthase kinase 3 beta, GSK‐3β, (Wilson & Lefebvre, 2004) and Adenomatous Polyposis Coli, APC (Corbit et al., 2008).

Wnt signaling is affected in ORPK mice which show abnormal cyst formation in the pancreas where the cysts and dilated ducts have an increased cytosolic localization of β‐catenin in addition to an increased expression of Tcf/Lef, which activates transcription of Wnt target genes (Willert & Nusse, 1998; Roose & Clevers, 1999; Cano et al., 2004; Zhang et al., 2005). In neural tube development inhibition of Shh via the Gli3 repressor inhibit the canonical Wnt‐mediated transcriptional activation by physical interaction with the carboxy‐terminal domain of β‐catenin (Ulloa et al., 2007). However, the primary cilium does not seem to be essential for the canonical Wnt signaling pathway demonstrated in recent findings in the mouse embryo and mouse embryonic fibroblasts (Ocbina et al., 2009). The Wnt signaling appears intact in the absence of primary cilia in Ift88 and Ift172 knockout mice or in the anterograde motor Kif3a and retrograde motor Dync2h1 knockouts (Ocbina et al., 2009). These data contradict previous findings, suggesting that IFT proteins and especially Kif3a have specific roles in


Figure 14: Brief overview of signaling pathways in cardiogenesis. A: Ciliary Hh and Wnt signaling pathways. With Hh ligand present the Gli transcription factor activators can control timely activation of early heart transcription factors. The non‐Canonical Wnt signaling pathway favors cardiogenesis through degradation of beta‐catenin. Wnt 11 has been observed to induce cardiomyogenesis (Flaherty & Dawn, 2008). In terms of Hh signaling, binding of ligands to Ptc in the cilium activate the Hh pathway by removal of Ptc from the cilium in a process that is associated with ciliary enrichment of Smo. The red arrow indicate a possible crosstalk between the Gli2 activator and the Wnt signaling pathways B: Bone morphogenetic protein (BMP) and transforming growth factor (TGF)‐beta signaling pathways function in formation of the endocardial cushion tissue development. The cushion tissue is formed in the outflow tract and in the atrioventricular regions during cardiogenesis (Yamagishi et al., 2009). Whether the two pathways work via the primary cilium is not yet understood. The red arrow indicates strong crosstalk between the two pathways. C: Platelet‐derived growth factor receptor (PDGF‐R) and fibroblast growth factor receptor (FGF‐R) signaling pathways are also involved in cardiogenesis. PDGF‐R signaling plays a role in the migration of epicardial cells that form the coronary artery and myocardium (Mellgren et al., 2008). FGF‐R signaling regulates the early heart transcription factors Gata5 and TBX6 and TBX16 (Neugebauer et al., 2009). Little is known about FGF signaling in the primary cilium in contrast to PDGF‐R signaling that has been documented to work through the cilium (Schneider et al., 2005; 2009a). The red arrow indicates strong crosstalk between the two pathways. D: Examples of potential signaling crosstalk between Wnt, BMP and Hh signaling pathways in embryonic development. Lack of Hh signaling was observed to delay BMP4 signaling in P19 cardiac progenitor cells (Gianakopoulos & Skerjanc, 2009). Furthermore, Gli3‐repressor activity was shown to negatively regulate Wnt/beta‐catenin signaling (Ulloa et al., 2007). The red arrow indicates possible crosstalk between the individual signaling pathways, although we still know very little as to the potential association with the primary cilium.


regulation of the canonical Wnt pathway (Gerdes et al., 2007; Gerdes & Katsanis, 2008; Corbit et al., 2008). In addition, zebrafish studies show that oval mutants (ovl;ift88) lack primary cilia but still have normal canonical and non‐canonical Wnt signaling but show defects in Hh signaling (Huang & Schier, 2009). A possible hypothesis is that the Wnt signaling pathway is intact because the basal body adapts the role of the primary cilium and compensate for the absence of cilia. Consequently, it is plausible that the ciliary axoneme and the basal body are two distinct signaling organelles with separable functions, where the ciliary axoneme is required for Hh signal transduction and the basal body might be essential for the Wnt signaling response. How the Wnt and Hh signaling pathway crosstalk in the primary cilium or not is not fully understood. Of the other cardiogenic pathways in figure 14, both BMP and (TGF)‐beta signaling pathways function in the formation of the endocardial epithelial‐mesenchymal transformation (EMT), which is a critical process in endocardial cushion tissue development. The cushion tissue is formed in the outflow tract and in the atrioventricular regions during cardiogenesis (Yamagishi et al., 2009). Whether the two pathways work via the primary cilium is not yet known, however the findings that Hh and BMP signaling crosstalk in cardiomyocyte formation in P19 cells (Gianakopoulos & Skerjanc, 2009), might indicate a possible ciliary mechanism for the BMP signaling pathway. The PDGF‐R and FGF‐R signaling pathways also play a role in cardiogenesis. PDGF‐R signaling plays a role in the migration of epicardial cells that form the coronary artery and myocardium (Mallgren et al., 2008), where FGF‐R signaling regulate the early heart transcription factors Gata5, TBX6 and TBX16 which are proposed to play a role for regulating ciliary length (Neugebauer et al., 2009). Little is still known about FGF signaling in the primary cilium where PDGF‐R signaling previously has been shown to regulate ciliary PDGF‐Rαα signaling that control tissue homeostasis, migration and mitogenic signaling pathways in fibroblasts (Schneider et al., 2005; 2009a; 2009b).

The complexity in early heart development makes it difficult to assess how individual signaling pathways function because of the possible crosstalk. Future in vitro experiments on Ift88‐/‐ mESCs, P19 and P19.CL6 EC cells with focus on the Wnt, Hh, FGF, PDGF and BMP signaling pathways should help determine how the individual pathways interact with one and other. In some cases, siRNA may have off target effects and may not induce a complete loss of function. In order to eliminate these concerns, more experiments with Ift88‐/‐ mESCs and wt as well as rescued mutant ES cells would clarify these possible off target effects. Further and in contrast to P19.CL6 cells, mESCs are true pluripotent stem cells that will give more clear information on the function of the primary cilium in the earliest steps of cell fate determination and prior to formation of cardiac progenitor cells. Copying the same characterization done on P19.CL6 EC cells onto the Ift88‐/‐ and wt mESCs will reveal interesting knowledge on how the early heart development progress in mESCs and possibly how Wnt and Hh signaling function in these cells, with and without inhibitors like KAAD‐cyclopamine. Preliminary data in the P19.CL6 differentiation studies reveal fluctuations of Ptc1 and Smo in and out of the cilium around the forming clusters of cardiomyocytes which is something that would be interesting to study in detail [4]. This might give more insight on how the Hh signaling pathway is regulated locally in cardiogenesis. Additionally, more in vivo studies are needed with chimera Ift88‐/‐ESC lines and Ift88‐/‐ mice followed by histological sectioning to pinpoint where the primary cilia are needed in embryonic development, in stem cell positioning and differentiation during the various stages of heart development from E8.0 old embryos and in adult mice. Moreover, immunofluorescence microscopy‐3D reconstruction and in situ hybridization analysis would identify ciliary signaling components during in vivo heart development, and reveal how defects in ciliary assembly affect the level of their activation in Ift88‐/‐ null embryos.


CHAPTER 4 – SENSORY CILIA IN THE PANCREAS AND REPRODUCTIVE ORGANS


This chapter will discuss and summarize the data from the secondary objective articles that are found in chapter 7 in the following sections: Collaborative work with Stefan Teilmann on progesterone receptor localization and expression in the female reproductive organs (Chapter 7: [1]) and on pancreatic development in humans in addition to cancer cell lines (Chapter 7: [2]).

4.2 HEDGEHOG SIGNALING IN PANCREATIC DEVELOPMENT AND CANCER

Precise and timely Hh signaling is required for proper regulation and development of the pancreas, but even in the mature adult tissue, Hh signaling take part in the general maintenance of the pancreatic tissue (Hebrok et al., 2000). Therefore, mal‐regulation of the Hh signaling pathway results in a series of diseases, including annular pancreas, diabetes mellitus, chronic pancreatitis and pancreatic cancer (Lau et al., 2006). The role of the primary cilium in pancreatic development and cancer via the Hh signaling pathway is not fully known and needs to be investigated further before treatments are effective enough to cure cases with pancreatic cancer.

4.2.1 PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREATIC DEVELOPMENT

In the early mouse development, the pancreas initiates its development around E8.0 and is situated in the anterior midgut region of the endoderm epithelium. The pancreas is formed by two primary tissues, the exocrine compartment, which contains acinar and ductal cells, and the endocrine compartment with cells that localize to the islet of Langerhans structure. The exocrine acinar cells compose the majority of the mature organ and produce enzymes that drain into the intestinal tract through the ductal tissue. The islets of Langerhans are imbedded within the exocrine tissue, where they produce important hormones that regulate blood glucose levels. These islets contain four different cell types, the glucagon producing α–cells, somatostatin producing δ‐cells, insulin producing β‐cells and the pancreatic polypeptide producing PP‐cells. Hence the main function of the pancreas is to produce enzymes and secrete hormones that aid digestion and controls blood glucose homeostasis (Slack, 1995).

As discussed in [2] a number of previous investigations have indicated a link between primary cilia and development of the pancreas in mice. In addition to kidney defects, the loss of primary cilia in the Tg737ORPK mouse causes a series of abnormalities in the pancreas, such as extensive cyst formation in ducts (Cano et al., 2004; Zhang et al., 2005). This may indicate a possible functional similarity between cilia in kidney and pancreatic duct systems. Cells of both exocrine and endocrine systems in the pancreas possess primary cilia, including islet cells and the ducts, but not in the acini (Kodama, 1983; Ashizawa et al., 1997; Cano et al., 2004; 2006; Zhang et al., 2005). In the Tg737ORPK mouse pancreas abnormalities begin with dilations of the ducts in late gestation, which after birth are accompanied by extensive formation of large, interconnected cysts as well as apoptosis and vacuolization of acini. In the dilated ducts and cysts PC‐2 is mislocalized to the intracellular compartments. These changes are reminiscent of chronic pancreatitis, supporting the speculation that primary cilia of ducts play an essential role in the development of the pancreas (Cano et al., 2004; Zhang et al., 2005). As


will be discussed below, primary cilia may also play a major role in coordinating Hh signaling during human pancreatic development and aberrant Hh signaling in pancreatic cancer may be associated with defects in ciliary assembly in pancreatic adenocarcinoma cell lines.

Hh signaling appears to play multiple roles during mouse embryonic pancreatic development. During the early stages of gut formation in mice, expression of both Shh and Ihh genes is found throughout the endoderm epithelium (Bitgood & McMahon, 1995; Aubin et al., 2002), even though both genes are absent from the early endodermal area specified to become pancreas (Hebrok et al., 1999; Kim & Melton, 1998). In situ hybridization experiments have shown that Ptc1 expression is found in the mesenchyme adjacent to, but missing from, the pancreas anlage in E9.5 old embryos (Apelqvist et al., 1997). This difference in Hh signaling may ensure the correct establishment of organ boundaries. Subsequently Hh signaling is activated to promote proliferation and maturation of the tissue, as observed at embryonic day E13.5 where the developing pancreas expresses several Hh genes such as Ihh, Dhh, Hhip and Ptc1 (Kawahira et al., 2005; Lau et al., 2006; Cano et al., 2007; van den Brink, 2007). In the adult pancreas both Ptc1 and Smo has been observed in the islet and ductal cells, which indicate that Hh signaling is present and active during later stages of pancreas development and in the mature organ (Hebrok et al., 2000; Kawahira et al., 2003). To sum up on the collaborative work with Sonja Brorsen [2], we found primary cilia projecting into the pancreas duct lumen. These primary cilia are up to 20µm long and show increased Smo and Gli2 localization when the embryos enter the fetal stages of development at weeks 14 and 18, compared to the 7.5 week old embryos in the embryonic stage. Contrarily, the nuclear and cytosolic expression levels of the repressor form of Gli3 decreases as the embryos enter the fetal stages, suggesting an increased Hh activity. These changes in localization correlate with known activity of the Hh pathway during pancreas development. The primary cilium may be the critical organelle that coordinates pancreatic development to promote maturation of the tissue and function in tissue homeostasis in adult individuals. Loss of primary cilia function in the Tg737ORPK mouse further strengthens the hypothesis that primary cilia are key regulators of pancreatic development, since these mice show severe pancreatic abnormalities including extensive cyst formation in the ducts (Cano et al., 2004; Zhang et al., 2005).

4.2.2 PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREATIC CANCER Pancreatic cancer is a severe disease that is diagnosed in 33.000 patients annually in the USA alone, world wide it is estimated to cause more than 200.000 deaths each year (Parker et al., 2003). Notch, Hh, and Wnt signaling pathways play an important role in multiple tissues during development and are for most part turned off in adult somatic cells, including the exocrine pancreas. Abnormal transcriptional activation of these pathways has been reported in both human and mouse models of pancreatic neoplasia. Aberrant activation of the Hh signaling pathway has been reported in pancreatic intraepithelial neoplasia (PanIN) (Miyamoto et al., 2003; Berman et al., 2003; Thayer et al., 2003; Zeng et al., 2006). In addition, activation of the Hh pathway in a human pancreatic ductal epithelial cell line resulted in up‐regulation of extra‐pancreatic foregut markers observed in the early PanIN lesions (Prasad et al., 2005), which normally are not present in normal ductal epithelium (Koorstra et al., 2008).

In the collaborative work with Sonja Brorsen [2] we further investigated the role of primary cilia in pancreatic adenocarcinoma cell lines, CFPAC‐1 and PANC‐1, which are isolated from metastatic and primary tumors, respectively (Schoumacher et al., 1990; Lieber et al., 1975). Initially, we show that CFPAC‐1 and PANC‐1 cells have long primary cilia, up to 20µm long, which have ciliary localization of Ptc, Gli2 and Smo. This localization is consistent with the idea that the primary cilium continues to coordinate Hh signaling in cells derived from the mature pancreas. Aberrant Hh signaling in these two cancer cell lines may be associated with the autonomous activation of the signaling pathway in the cilium, judged by the relative high levels of Smo and low levels of Ptc in the cilium. Furthermore, the high expression of full‐length form of Gli2 in the cilium and low levels of Gli3 repressor in the nucleus suggest even further that there is a high activation level of Hh signaling pathway. The fact that Hh signaling is highly active in the CFPAC‐1 and PANC‐1 cell lines suggests that ciliary Hh signaling


plays a potential role in tumorigenesis. However these results proposes that a certain level of Hh signaling is required for proper organ formation, as observed in gain of function studies that have demonstrated that de‐regulation of Hh signaling activity results in critical changes of pancreas morphogenesis and function (Lau et al., 2006).

4.3 SENSORY MOTILE CILIA IN THE OVIDUCT

The mouse ovary is enclosed in a thin epithelial bursa. The inner surface epithelium of the bursa is continuous with the ovarian surface epithelium in the areas around the hilus or ovarian stalk. It encloses the ovary and the upper part of the oviduct, the infundibulum. Ovulated oocytes are prevented of escaping into the peritoneum by the bursa and are led to the opening of the oviduct (salpinx), which collects the oocytes. The mouse oviduct has a coiled appearance and can be divided into four parts (Nielsson & Reinius, 1969), see figure 15. In the ovary the follicles begin to grow soon after birth and continue until the pool of follicles is depleted (Peters et al., 1975). Initiation of follicle growth begins with mitotic activity of the granulosa cells surrounding the oocyte, which also increases in volume (Pedersen & Peters, 1968). In the late follicular development, multiple layers of granulosa cells develop around the oocyte where fluid begins to accumulate in the space around the oocyte (called antrum). A wide range of signaling pathways and hormones control the follicular development including progesterone. Ovulation is controlled by the luteinizing hormone, LH, which induces luteinization of granulosa cells that particularly increases the expression of progesterone and its receptor (PR), (Shimada & Terada, 2002; Park & Mayo, 1991). Progesterone is an important local regulator of ovulation, lutenization, oviductal gamete transport and implantation. Additionally progesterone mediates various effects in the female reproductive organs through its cognate nuclear receptors, PR‐A and PR‐B which often are co‐expressed within the same cells, e.g. in granulosa cells of pre‐ovulatory follicles. The two receptor types although co‐expressed have distinct roles and show different phenotypes in knockout mice. In PRA‐/‐ mice ovulation is critically impaired and the implantation is no longer possible, demonstrating that only PR‐A is obligatory for female fertility, in contrast to PRB‐/‐ mice that has defective mammary gland development (Mulac‐Jericevic et al., 2000).

Motile and primary cilia are found in abundance in the oviduct. In the ovaries, primary cilia are particularly found on the granulosa cells in the antral follicles, which have been shown to have the TRP ion channel, polycystin‐1 and ‐2 localizing to the primary cilia (Teilmann et al., 2005). Furthermore, the angiopoietin receptors such as the Tie‐1 and Tie‐2 receptor tyrosine

Figure 15. Schematic view of the mouse reproductive organs. The infundibulum: the outer part closest to the ovary (also known as fimbriae or ostium). This part of the oviduct is thin walled with an epithelium almost completely covered by ciliated cells that facilitate pickup and transport of ovulated eggs. The ampulla: a thicker walled part of the oviduct with slightly less ciliated cells, fertilization occurs here. The isthmus: has a thicker wall characterized by increased musculature. Ciliated epithelial cells are becoming rarer. The juncture: which is connecting the oviduct to the uterus is not shown, as well as the ovarian bursa for simplicity (Modified from; Stefan Teilmann PhD thesis, 2005).


kinases localize to motile cilia of the oviduct. Tie‐2 specifically localize to primary cilia of the surface epithelium of the ovary, bursa and extra‐ovarian rete ducts as well as to plasma membranes of the ovarian theca and endothelial cells (Teilmann & Christensen, 2005). In the oviduct ciliated epithelial cells of both adult human and mice, revealed progesterone localization to the lower half of the motile cilia, whereas the nuclei were not stained or otherwise only faintly. It is possible that ciliary progesterone receptors in the oviduct play a role in progesterone signaling after ovulation, possibly via non‐genomic events [1]. The presence of progesterone/Tie receptors in addition to polycystins in the cilia population of the female reproductive organs, support the hypothesis that cilia both motile and primary, play an important sensory role in coordinating and regulating hormonal and reproductive events.

To sum up on the collaborative work with Stefan we found PR localization in the lower half of the motile cilia in the oviduct. The ciliated cells with PR localization did not show PR localizing to the nuclei or if the case, only very faintly. In the pubertal mice the localization of PR was increased in the cilia, in contrast to the primary granulosa cell cilia, which lacked PR staining at all stages. Since progesterone is a regulator of ciliary beat frequency, we suggest that ciliary PR directly modulates the ciliated oviduct epithelium by operating as a fast means to sense and relay changes in the levels of progesterone in the oviduct, such as those induced through release of follicular fluid at ovulation or released by the oocyte cumulus complex. In this scenario, ciliary beat frequency may be regulated directly by progesterone via ciliary receptors to control uptake and/or transport of the oocyte cumulus complex. Additionally, the findings of polycystins 1 and 2 as well as Tie receptors to motile cilia in the oviduct further support the hypothesis that cilia of the female reproductive organs play a significant sensory role in relaying physiochemical changes from the extracellular environment to epithelial cells of the oviduct and ovary (Teilmann et al., 2005; Teilmann & Christensen, 2005).

4.4 SECONDARY OBJECTIVE CONCLUSIONS AND PERSPECTIVES

Motile cilia in the mouse and human oviduct revealed progesterone receptor localization in the lower half of the cilia. This localization was increased in pubertal mice, which suggest a possible role for the cilia in progesterone signaling after ovulation. Hedgehog signaling also plays a significant role in development of the pancreas in mammals. Ciliary localization of Gli2 and Smo in both cultures of human pancreatic duct adenocarcinoma cell lines and in duct epithelial tissue indicates that Hh signaling is a strong regulator in pancreatic development, which may also be responsible for tumorigenesis. Tight regulation of the Hh pathway in the embryo, fetus and adult is critical judged by the observed changes from the 7.5 weeks old embryos to the 14 and 18 weeks old fetuses. These findings support the hypothesis that primary as well as motile cilia play significant roles in cell signaling to maintain tissue homeostasis and control differentiation, in addition to coordinate developmental events.


CHAPTER 5 – THESIS CONCLUSIONS

5.1 THESIS CONCLUSIONS

This thesis presents data to support the conclusion that primary cilia as well as motile cilia in the oviduct are critical organelles in coordinating signaling transduction pathways during development and in tissue homeostasis. Several of the studies presented have focused on the Hh signaling pathway, which is an important pathway in heart development and differentiation. Aberrant Hh signaling can lead to cancer if not properly coordinated, however very little is still known about the Hh signaling pathway and how it is coordinated in vivo during heart development and in the adult. The connection between cilia and various human diseases has clearly demonstrated the importance of cilia. Many functions in cilia assembly and maintenance are still not known and will need to be investigated to determine how diseases arise from ciliary defects. A future challenge will be to further improve our understanding of the ciliary signaling pathways and how receptors in addition to signaling molecules work via the primary cilium, which impinges on cellular responses and gene expression. Especially in heart development and stem cell research, the primary cilium may play a significant role in regulating cellular processes that can be used as effective therapeutic tools against cancer and in regeneration of damaged tissue in the near future.


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CHAPTER 7 – ARTICLES [1‐5]

525

Expression and localization of the
progesterone receptor in mouseand human reproductive organs
Stefan Cuoni Teilmann, Christian Alexandro Clement2, Jørgen Thorup1, Anne Grete Byskov

and Søren Tvorup Christensen2

Laboratory of Reproductive Biology, Juliane Marie Center, Rigshospitalet, Copenhagen, Denmark1Department of Paediatric Surgery, Section 4072, Rigshospitalet, Copenhagen, Denmark2Department of Biochemistry, Institute of Molecular Biology and Physiology, University of Copenhagen, The August Krogh Building, Universitetsparken 13,

DK-2100 Copenhagen, Denmark

(Requests for offprints should be addressed to S T Christensen; Email: [email protected])

Abstract

The effects of gonadotropins on progesterone receptor (PR)

expression and localization in the mouse oviduct, uterus, and

ovary was examined. In the oviduct ciliated epithelial cells of

adult mice and human revealed a unique PR localization to

the lower half of the motile cilia whereas the nuclei were

unstained or faintly stained. Pubertal female mice were

further studied by confocal laser scanning microscopy and

western blotting before and after injection with FSH and LH

followed by human chorionic gonadotropin (hCG) injection

after a 48-h period. PR immunolocalization to the oviduct

cilia was greatly increased in pubertal mice upon hCG

stimulation. In neighboring goblet cells, the PR staining was

confined to the nuclei. Nuclear PR localization was evident

in epithelial cells of the uterus as well as in a fraction of stromal

and muscle cells. Staining intensity and number of stained

cells was not affected by hormone stimulation. In the ovary,

weak PR immunolocalization was observed in unprimed

Journal of Endocrinology (2006) 191, 525–5350022–0795/06/0191–525 q 2006 Society for Endocrinology Printed in Great

animals but increased significantly after hCG stimulation. In

granulosa cells of preovulatory follicles PR was exclusively

observed in mural cells, whereas cumulus cells remained

negative. At all stages examined, primary granulosa cell cilia

lacked PR staining. SDS-PAGE and western blotting analysis

of tissues from oviduct, uterus, and ovary confirmed antibody

specificity, and identified two bands corresponding to the PR

isoforms PR-A and PR-B. Upon hCG stimulation, a new

band cross-reacting with anti-PR emerged above the PR-A

form in oviduct fractions, suggesting LH-induced phos-

phorylation of PR-A. We suggest that ciliary PR in the

oviduct plays a role in progesterone signaling after ovulation,

possibly via non-genomic events. These novel findings

warrant further studies of oviduct and postovulatory signaling

events and suggest a sensory role for oviduct cilia in the

process of oocyte transport/fertilization.

Journal of Endocrinology (2006) 191, 525–535

Introduction

Many reproductive functions in mammals are regulated by the

ovarian steroids estradiol and progesterone (P). P is an important

local regulator of critical reproductive events such as ovulation,

luteinization, gamete transport within the oviduct, and

implantation. P mediates various effects in the female

reproductive organs through its cognate nuclear receptors,

PR-A andPR-B,which often are co-expressedwithin the same

cells, e.g., in granulosa cells of preovulatory follicles (Hild-Petito

et al. 1988, Sheridan et al. 1989, Park &Mayo 1991, Gava et al.

2004). Both the PR forms are ligand-activated transcription

factors that upon ligand binding undergo phosphorylational and

conformational changes (Weigel 1996, Clemm et al. 2000). The

importance of the two PR isoforms has been demonstrated in

PR null mice (PRKO), which are anovulatory and unable to

properly respond to exogenous gonadotropins (Lydon et al.

1995). Mice deficient in either PR-A (PRAKO) or PR-B

(PRBKO) have different phenotypes, demonstrating a complex

and tissue-specific interplay between the two receptor forms.

In PRAKO mice, ovulation is severely hindered and

implantation impossible showing that only PR-A is obligatory

for mouse female fertility (Mulac-Jericevic et al. 2000).

Cyclic changes of P and estradiol control homeostasis of the

oviduct epithelium as well as formation and beat frequency of

the cilia (Brenner 1969, Wessel et al. 2004), although little is

known about the underlying regulatory mechanisms.

Present research has positioned cilia as key sensory organelles

in human health and reproduction (Satir & Christensen

2006), and increasing evidence suggests a role for sensing cilia

in the female reproductive organs. We have previously

described the unique localization of signal components to

both motile cilia of the oviduct and to primary cilia of ovarian

and extraovarian tissues of the mouse, and signaling through

these cilia were suggested to play a role in follicular

maturation, ovulation, and gamete transport. In the ovary,

DOI: 10.1677/joe.1.06565Britain Online version via http://www.endocrinology-journals.org

S C TEILMANN and others . PR expression and localization526

the Ca2C-selective transient receptor potential (TRP) ion

channels localize to primary cilia of granulosa cells of antral

follicles and to motile cilia of the oviduct (Teilmann et al.

2005). In addition, receptor tyrosine kinases, angiopoetin

receptors, localize to cell type-specific primary cilia of the

reproductive organs and to the tip of motile cilia in the

oviduct (Teilmann & Christensen 2005).

In the present study, we analyzed murine PR in oviduct,

uterus, and ovary by immunolocalization and western

blotting techniques upon follicle stimulating hormone

(FSH) and human chorionic gonadotropin (hCG) stimu-

lation. Using specific antibodies against PR, we were able to

show that hCG radically upregulates PR in nuclei and cytosol

of mural granulosa cells of the preovulatory follicles and in

motile cilia of the oviduct. These results indicate the

discovery of a novel function of PR in reproductive biology,

in which ciliary beat frequency may be regulated directly by

progesterone via ciliary receptors.

Materials and Methods

Animals and tissues

Pubertal female C57Bl/6J mice (21–23 days old,w14 g) were

injected intraperitoneallywith 30 IUgonadotropins (Menopur:

15 IU FSH and 15 IU luteinizing hormone (LH) (Ferring

Pharmaceuticals, Copenhagen, Denmark) followed by 5 IU

menotropin (hCG) (Pregnyl, Organon AS, Skovlunde, Den-

mark) injection 48 h later. Mice were killed by cervical

dislocation 48 h after FSH or 6 h after hCG. Non-stimulated

mice of coeval age served as controls. Oviducts, uteri, and

ovaries were immediately removed and processed for immu-

nohistochemistry (IHC) or western blot and SDS-PAGE

analysis. Furthermore, lung, heart, and reproductive organs of

3-month-old femalemice in estrouswere used as control tissues.

Ovaries from three 30-days-old Wistar rats were collected in

PBS rinsed from fat and extraovarian tissue, fixed in 4% para

formaldehyde, and processed for IHC. Animal experiments

were conducted in accordance with EU guidelines and

approved by the Danish Ministry of Justice, Animal Ethics

Committee no. 2003/561-713 (A G B).

Human material

Humanmaterial was used after ethical approval was obtained by

the ethics committee forCopenhagen andFrederiksbergno.KF

01-170/99, and after informed consent was obtained from each

patient following the guidelines in the Declaration of Helsinki.

Immunohistochemistry

After fixation, oviduct, uterus, and ovary were embedded in

paraffin and cut in 8 mm thick sections that were collected on

microscope slides (SuperFrost/Plus, Menzel Glaser, Germany).

Importantly, we found what appeared to be time- and

temperature-dependent degradation of specific PR epitopes in


the tissue sections examined, leading to decreased or absent

staining intensity. Multiple sections from ovaries from three

different animals in each group were therefore always stored at

4 8C and used within a week after cutting. Sections were

deparaffinized, rehydrated, and rinsed in PBS. For antigen

retrieval, slides were boiled in citric acid buffer (0.01 M, pH 6),

and incubated for 15 min in PBS (pH 6.5) containing 5% (w/v)

BSA and 1% (v/v) preimmune goat serum (Dako, Glostrup,

Denmark). Sections were incubated with one of the following

primary antibodies diluted inPBS containing5% (w/v)BSAand

0.02% (w/v) NaN3 overnight at 4 8C: diagnostic grade rabbit

monoclonal PR antibody (1:300, Clone SP2, LabVision,

Westinghouse Drive, Fremont, CA, USA) directed against an

epitope corresponding to amino acid (aa) sequence 410–516 in

mouse PR and to aa sequence 412–526 in human PR was a

gift from AH Diagnostics, Aarhus, Denmark. Mouse mono-

clonal PR antibody (1:300, Clone Ab-4, NeoMarkers,

LabVision) directed against the N-terminal region of PR, i.e.

aa sequence 1–557 in mouse PR and aa sequence 1–566 in

human PR. Mouse monoclonal anti-acetylated a-tubulin(1:3000, Cat no. T6793, Sigma-Aldrich) for localization of

cilia and cytosolic network of acetylated microtubules (Alieva

et al. 1999). Non-specific binding of the PR antibody was

evaluated by substitution with preimmune rabbit IgG (Dako)

with the same concentration as primary antibody. Primary

antibodieswere detected by 1-h incubation at room temperature

with species-specific Alexa Fluor anti-IgG F(ab0)2 secondary

antibody (5 mg/ml, Molecular Probes, Eugene, OR, USA)

and counterstained with propidium iodide (1 mg/ml) or

TO-PRO-3 iodide (2 mg/ml Molecular Probes) in PBS for

8 min.Afterwashing, slidesweremounted in 1:1 (v/v) glycerol/

PBSwith 2% (w/v)NaN3 and sealedwith nail polish. A series of

sections as well as isolated and fixed single cell preparations

(see below) were double labeled with anti-acetylated a-tubulinand anti-PR.

Tissue and immunofluorescence analysis

The three parts of the oviduct studied were divided into the

following separate groups: fimbriae (the cranial part), ampulla

(the middle part), and isthmus (the caudal part). For single cell

analysis, the cranical and middle part of the oviduct were

collected in PBSwithCa2C/Mg2C and cut into small pieces to

expose the ciliated epithelium. Then the tissues were placed in

ice-cold incubation buffer containing 0.25 M sucrose, 0.02 MHEPES, 2 mM EDTA, and 25 mM KCl supplemented with

freshly made phenylmethylsulfonyl fluoride (1 mM) and

N-ethylmaleimide (10 mM). For the next 50 min, the sample

was kept on ice and vortexed intermittently. Pieces of oviduct

were removed and an equal volume of incubation buffer was

added. After 60 s of incubation, the material was vortexed and

spun down (500 g for 10 min). The pellet (containing ciliated

cortices, single cells and nuclei) was fixed and loaded onto glass

cover slips for immunohistochemical analysis.Ovarian follicles

were categorized according to Pedersen & Peters (1968) and

antral follicleswere classified as atreticwhenO5 granulosa cells

www.endocrinology-journals.org

PR expression and localization . S C TEILMANN and others 527

were pyknotic, corresponding to the stage one atresia

described (Byskov 1974) and was excluded from analysis.

Stained sections and isolated cells were observed on an IX70

confocal laser scanning microscope (Olympus, Tokyo) with a

Krypton/Argon laser using a 60! oil immersion objective

(NA:1.25) and a 40! air objective (NA:0.85), both equippedwith appropriate Normarski optics. Care was taken to avoid

bleedthrough between channels, and at the beginning of each

evaluation, image settings was optimized so that it contained

the maximum number of gray levels, and during subsequent

image acquisition all settings (laser power, photomultiplier

tube gain and offset) were kept constant so that the images

could be compared.

SDS-PAGE and western blot analysis

In the isolation of oviduct infundibulum samples, carewas taken

only to include the outer cranial part in order to minimize non-

ciliated epithelium from the sample. Ovaries and oviducts were

cleaned from fat and connective tissues before protein isolation.

Protein was extracted using a common protocol (VanSlyke &

Musil 2001). The protein concentrationswere estimated using a

BCA protein kit (Pierce Biotechnology, Rockford, IL, USA),

and proteins were resolved by gel electrophoresis under

denaturing and reducing conditions and electrophoretically

transferred to nitrocellulose membranes as previously described

(Christensen et al. 2001). The membranes were incubated with

anti-PR (1:300) and anti-b-tubulin (1:300; Cat no. T4026,

Sigma-Aldrich) and antibody cross-reactivities were identified

with species-specific alkaline phosphatase-coupled secondary

antibodies (1:1200, Jackson Laboratory, Bar Harbor, ME,USA)

followed by developing with 5-bromo-4-chloro-3-indolyl

phosphate/nitro blue tetrazolium (BCIP/NBT) (KPL, Cessna

Court, Gaithersburg, Maryland, USA).

Statistical analysis

Band intensities of PR proteins in western blot analysis were

measured using UN-SCAN-IT Version 5.1 (Silk Scientific,

Inc., Orem, Utah, USA). Data are presented as mean valuesGS.E.M. from a minimum of three individual experiments, in

which tissue homogenates were obtained from a minimum of

six animals. Significant differences in the level of PR expression

between non-stimulated and hormone-stimulated mice were

estimated using a two-tailed paired t-test. For all statistical

evaluations, P values !0.05, !0.01, and !0.001 were

considered statistically significant, very significant, and extre-

mely significant respectively.

Results

Localization and expression of PR in the ovary upon hormonestimulation of pubertal mice

In the untreated pubertal ovary anti-PR (SP2) weakly

localized to theca and granulosa cells (Fig. 1A and D), and a


slight nuclear anti-PR (SP2) staining was observed in the

interstitial tissue. Forty-eight hours after FSH a subpopulation

of theca cells around large antral follicles (stage 6) began to

show nuclear PR immunoreactivity (SP2), whereas the PR

signal seemed reduced in a subpopulation of granulosa cells of

any follicle stage (Fig. 1B and E). Six hours after hCG, nuclei

of many theca cells in the large preovulatory follicles (stage 7)

were anti-PR (SP2) positive (Fig. 1C). At this stage, follicular

granulosa cells are divided into two distinct populations:

mural granulosa cells facing the theca cell layer and cumulus

granulosa cells facing the oocyte. Clearly, hCG stimulation

dramatically increased the level of anti-PR (SP2) immuno-

fluorescence of mural granulosa cells, whereas cumulus cells

remained PR negative (Fig. 1F). To further characterize PR

localization in the granulosa cells of hCG-stimulated ovaries,

we used anti-acetylated a-tubulin to detect primary cilia in

mural and cumulus granulosa cells in stage 7 follicles. Most

non-dividing mural and cumulus granulosa cells had a

primary cilium that was often presented into the antrum

(Fig. 1G). In double labelings of anti-acetylated a-tubulin andanti-PR in granulosa cells, nuclear and cytosolic PR

expression did not correlate with the presence of a cilium

and no detectable co-localization between acetylated

a-tubulin and PR was observed (Fig. 1G). As a further

control on PR localization in the ovary, tissue sections of mice

were subjected to co-localization analysis with anti-PR (clone

Ab-4) that recognizes the entire N-terminal region of PR

(Fig. 1H). It is seen that SP2 and Ab-4 co-localize to mural

granulosa cells in hCG-stimulated ovaries, confirming

specific immunoreactivity to PR in this cell population.

A strong immunofluorescent signal in the cytoplasm of

mouse oocytes from all follicle stages including atresia was

observed with Anti-PR (SP2; Fig. 1A–F). However, no

immunofluorescent signal was detected with this antibody in

oocytes from either rat or human (Fig. 1I). Importantly, the

Ab4 antibody did not label oocytes from either rat or mouse

(Fig. 1I). Since, unspecific immunohistochemical staining of

oocytes is a well known problem, these observations suggest

that excessive oocyte staining can be considered a phenom-

enon restricted to that particular antibody when used on

mouse tissue. Although we do not know whether this signal

represents a true receptor population within the oocytes, it is

most likely an artifact and has not been investigated further.

Using western blot analysis anti-PR (SP2) specifically

recognized two PR forms of approximately 115 and 83 kDa,

corresponding to the B- and A-form respectively. In the ovary

protein fraction of the non-stimulated mice both PR forms

could be detected at a low level (Fig. 1J). After 48 h with FSH,

the level of PR-A was significantly reduced to about 40%

comparedwith that of the non-stimulatedmice (Fig. 1J andK).

In contrast, 6 h after hCG, PR-A and PR-B were present at a

level about 19- and 8-fold higher than in the non-stimulated

mice respectively (Fig. 1J and L). In some experiments, we also

observed a protein migrating in SDS-PAGE as a 60 kDa

protein, which was recognized by anti-PR (SP2) and

upregulated upon hCG stimulation (data not shown).


Figure 1 Immunolocalization of progesterone receptor in ovary tissue sections before and after gonadotropic stimulation of pubertal femaleC57Bl/6J mice. (A) Control ovary (no stimulation; TZ0). (B) Forty-eight hours after FSH stimulation (TZ48). (C) Six hours after additionalstimulation with hCG (TZ48C6). Scale bar (A–C): 200 mm. Corresponding close ups of pubertal (TZ0) stage 5b follicle and granulosa cells(D and D 0), stage 6 follicle and granulosa cells after 48 h FSH/LH (TZ48) (E and E 0), and stage 7 follicle and cumulus granulosa cells (arrows)6 h after hCG (TZ48C6) (Fand F 0). Cumulus granulosa cells are marked with asterisks. Nuclei are detected with propidium iodide (red). Scalebar (D–F): 50 mm. (G) Single mural granulosa cell presenting a primary cilium detected with anti-acetylated a-tubulin (red, arrow head). Nucleiare detected with TO-PRO (blue). nu, nucleus; cyt, cytoplasm; PM, plasma membrane. Progesterone receptor is localized with monoclonalrabbit anti-PR (SP2; green). (H) Co-immunolocalization of progesterone receptor with monoclonal rabbit anti-PR (SP2; green) and monoclonalmouse anti-PR (Ab-4; red) in mural granulosa cells (arrows) of a stage 7 follicle and 6 h after hCG (TZ48C6). Nuclei are detected with TO-PRO(blue). Scale bar: 50 mm (I) Immunolocalization of anti-PR (SP2; green) and anti-PR (Ab-4; red) in tissue sections of adult rat, human, and mouseovaries. Scale bar: 50 mm. (J). SDS-PAGE and western blot analysis with anti-PR (SP2) showing the expression of progesterone receptor forms,PR-A (ca. 83 kDa) and PR-B (ca. 115 kDa), at different times after the gonadotropic stimulation. Anti-b-tubulin was used as a loading control.Molecular markers (kDa) are shown to the left. (K and L) Relative levels of PR-A (open bars) and PR-B (solid bars) in ovaries after 48 h FSH (TZ48)and 6 h hCG (TZ48C6) compared with pubertal controls (TZ0). Error bars indicate standard errors from five separate experiments. Significantchanges in PR levels are marked with one, two and three asterisks, for P!0.05, P!0.01, and P!0.001 respectively.


Localization of PR in the oviduct and uterus of adult mice

Localization and expression of PR isoforms in the oviduct

and uterus were initially examined in tissues of adult mice

(Fig. 2). In all parts of the oviduct, nuclear PR

immunoreactivity was detected in a fraction of the stromal

cells and the luminal non-ciliated epithelial cells. In ciliated


epithelial cells, anti-PR (SP2) uniquely localized to the cilia,

whereas the antibody exclusively localized to the nucleus in

non-ciliated glandular goblet epithelial cells (Fig. 2A). In the

uterus, anti-PR (SP2) localized to nuclei of epithelial,

stromal, and muscle cells (Fig. 2B). In the stroma and muscle

layers, PR staining had a mosaic-like pattern with


A: oviduct B:uterus

PR(SP2)

PR(SP2)

PR(SP2)

C

ovid

uct

uter

uslu

ng

PR-B

PR-A

116.397.4

hear

t

D: Lung

A´

F: human oviduct I

PR (SP2)PR (Ab-4)PR (SP2)

PR (SP2)

DIC

tissue

tb PR (SP2) merged

DIC

tb PR (SP2) merged

isolated cell

E

PR (Ab4)

G

PR (Ab4)

H

*

*

**

Figure 2 Immunolocalization of progesterone receptor in tissue sections of oviduct (A) and uterus (B) of adult C57Bl/6J female mice.Cilia are marked with arrow heads and nuclei are stained with propidium iodide (red). Progesterone receptor is localized withmonoclonal rabbit anti-PR (SP2; green). Scale bars: 50 mm. (A0) Close up of the mouse oviduct epithelium showing ciliated and gobletcells. Cilia are marked with arrow heads. Nuclei are stained with propidium iodide (red) and progesterone receptor is localized withmonoclonal rabbit anti-PR (SP2; green). Scale bar: 10 mm. (C) SDS-PAGE and western blot analysis with anti-PR (SP2) showing theexpression of progesterone receptor forms, PR-A (ca. 83 kDa) and PR-B (ca. 115 kDa) in extracts of oviduct, uterus, lung, and heart ofadult mice. Molecular markers (kDa) are shown to the left. (D) Immunolocalization of anti-PR (SP2) in a tissue section of ciliated lungepithelium from mouse (arrow heads indicate cilia). Nuclei are stained with propidium iodide (red). Scale bar: 50 mm. (E) Ciliaryimmunolocalization of anti-PR (SP2; green) in the mouse oviduct in a tissue section (upper row; scale bar: 5 mm) and in an isolated ciliatedepithelial cell (lower row; scale bar: 1 mm). Cilia are detected with anti-acetylated a-tubulin (red) and marked with arrow heads. Nucleiare detected with TO-PRO (blue). Nuclear localization of anti-PR is indicated with asterisks. (F) Immunolocalization of anti-PR (SP2;green) in a tissue section of ciliated epithelial cells from adult human oviduct (arrow heads indicate cilia). Nuclei are stained withpropidium iodide (red). Scale bar: 5 mm. (G and H) Immunolocalization of anti-PR (Ab-4; green) in tissue sections of ciliated epithelialcells from adult mouse (arrow heads indicate cilia). Nuclei are stained with propidium iodide (red). Scale bars: 25 mm (G) and 5 mm (H). (I)Co-immunolocalization of anti-PR (Ab-4; red) and anti-Pr (SP2; green) in a tissue section of ciliated epithelial cells from adult mouse(arrow heads indicate cilia). Nuclei are stained with propidium iodide (red) and nuclear localization of anti-PR is indicated with asterisks.


www.endocrinology-journals.org Journal of Endocrinology (2006) 191, 525–535


neighboring cells having PR-positive or negative nuclei.

SDS-PAGE and western blot analysis showed that anti-PR

(SP2) recognized PR-A and PR-B isoforms in isolated

tissues of both oviduct and uterus (Fig. 2C). PR expression

was not detected in lung extracts (Fig. 2C) and no PR

immunoreactivity was detected in lung cilia and epithelial

cells (Fig. 2D). This confirms specificity of PR to cilia of the

oviduct. Further, PR was not detected in heart extracts

(Fig. 2C).

The subcellular localization and expression level of PR in

the oviduct was investigated by high resolution confocal laser

scanning microscopy. Double labeling of anti-acetylated

a-tubulin and anti-PR (SP2) in tissue sections and isolated

cells from the oviduct epithelium of hCG-stimulated mice

showed that PR was confined to the lower region of the cilia

and confirmed that nuclear PR is mostly limited to goblet

cells in the epithelium (Fig. 2E). A similar pattern of

immunolocalization was observed in human oviduct epi-

thelium (Fig. 2F). As a further control, PR localization in the

mouse oviduct was investigated using anti-PR (clone Ab-4)

raised against the entire N-terminal region of PR. This

antibody localized in a similar fashion as SP2, i.e. it localized

to the nuclei of stromal cells, to the nuclei of goblet cells, and

to the lower part and at the base of the cilia in ciliated

epithelial cells (Fig. 1G–I). Importantly, we observed that

Figure 3 Subciliary immunolocalization of progesterone receptormice. (A) Lateral fluorescence intensity profile of PR immunofluore(white bar). DIC profile was used to define the base and the tip ofimmunofluorescence (SP2) along the cell surface (white bar). Anti-heads) along the epithelial surface.


tissue sections kept at room temperature for several days show

no or very little ciliary PR localization, whereas localization

to nuclei remains intact. This may mean that ciliary PR is

subjected to rapid degradation if sections are not stored

properly.

In order to analyze ciliary localization of PR in more detail,

we performed a lateral inspection of anti-PR (SP2)

fluorescence along the length of the cilia in tissue sections

of oviduct epithelial cells in hCG-stimulated mice. We used

the differential interference contrast (DIC) signal as well as a

vertical intensity plot of PR fluorescence and acetylated

a-tubulin fluorescence that mark individual cilia along the

surface of the cells (Fig. 3). The lateral inspection showed that

PR localization was restricted to the lower half of the ciliary

area (Fig. 3A), and that this localization was specifically

assigned to individual cilia (Fig. 3B). These results show that

PR in ciliated epithelial cells localizes to the lower half of the

cilia and not to microvilli, which are positioned at the base

between individual cilia.

Localization and expression of PR in the oviduct and uterus uponhormone stimulation of pubertal mice

In the untreated pubertal oviduct, PR (SP2) predominantly

localized to stromal cell nuclei, whereas ciliary localization

(green) in tissue sections of oviduct of adult C57Bl/6J femalescence (SP2) along the length of the cilia of the infundibulumthe cilium. (B) Vertical fluorescence intensity profile of PRacetylated a-tubulin (red) was used to define the cilia (arrow



was weak (Fig. 4A). Upon 48 h FSH stimulation nuclear PR

immunoreactivity was increased in the stromal cell nuclei of

the ampulla and isthmus, whereas this was less pronounced in

stromal cells of the infundibulum (not shown). Six hours after

hCG, the stromal as well as the non-ciliated epithelial cell nuclei

PR immunoexpression was increased in all regions of the

oviduct with the weakest relative staining intensity observed in

the infundibulum (Fig. 4B). At this time, ciliary PR localization

was heavily increased along the entire oviduct; this was also

associated with a minor nuclear PR staining in the ciliated

epithelial cells of the ampulla (Fig. 4B). The increase in ciliary

PR along the oviduct was predominantly confined to the lower

half of the cilia (Fig. 4C).Uponelicitationofovulation16 h after

hCG treatment, PR was still absent in mural granulosa cells but

present at a high level in the cilia of the oviduct (Fig. 4D). The

figure shows a close up of the cumulus cells in close proximity

with thecilia of theoviduct.These results support theconclusion

that ciliary PR is important during transport of the cumulus-

oocyte complex, since cumulus cells secrete progesterone to the

oviductal fluid.

In protein samples from whole infundibulum tissue, PR-A

and PR-B immunoreactivity could be demonstrated with anti-

PR (SP2) in western blot analysis in both non-stimulated and

hormone-stimulatedmice (Fig. 4E). However, the level of these

proteins increased significantly uponhormonal stimulation, such

that the levels of PR-A and PR-B increased about twofold after

48 h of FSH stimulation, and three- and fourfold respectively

after an additional 6 h treatment of hCG(Fig. 4E andF). Further,

after 6 h of hCG, we observed the appearance of a distinct and

major PR-immunoreactive protein band migrating just above

PR-A (Fig. 4E). The level of this band was increased about

25-fold upon hCG treatment (Fig. 4F). It was also observed that

the increase in the level of PR-Buponhormonal stimulationwas

associated with a slight migration shift such that the protein

migrated at a higher molecular mass (Fig. 4E).

In the uterus, the luminal epithelium PR localization was

less intense after 48 h of FSH, although after 6 h of hCG, the

nuclear epithelial cell staining slightly intensified (data not

shown). Uterus stroma and muscle cell nuclei PR staining was

comparable in all groups examined. Western blot analysis of

protein samples from uterus indicated that the levels of PR-A

and PR-B are not significantly altered upon hormonal

stimulation (Fig. 4H and G), although the PR-A protein

band appeared slightly more diffuse after hCG.

Discussion

In the mouse oviduct, we show here for the first time a

unique PR localization to motile cilia of the epithelial cells.

The biological significance of this finding is strengthened by

the fact that the PR staining intensity increased upon

gonadotropic stimulation of pubertal mice. The gonado-

tropin-primed mouse has close similarities with the pre-

ovulatory cyclic mouse and the staining pattern of PR in the

fallopian tube, ovary, and uterus in the present study is largely


comparable (Gava et al. 2004). However, previous studies

failed to identify PR in ciliated cells of the oviduct and it was

speculated that effects on ciliary activity were mediated by

goblet or stroma cells positive for PR (Okada et al. 2003). Our

findings on ciliary PR suggest a novel function of PR that

could aid in the regulation of ciliary activity and function of

oviduct epithelium.

The observed PR localization is confined to the lower half

and at the base of the cilium, suggesting that ciliary activity

regulated by P is mediated through effector molecules that

specifically localize to this part of the cilium. Fliegauf et al.

(2005) showed that outer arm dynein (OAD) heavy chains

regulating ciliary beat frequency in human respiratory cilia

and sperm flagella may be regionally and differentially

distributed along the axoneme, indicating that regulated

beat frequency is controlled by regional localization of OAD

heavy chains. Controlled ciliary beating is essential for proper

pickup and transport of the ovulated cumulus–oocyte

complex (COC), although the identity of extracellular signals

that regulate beat frequency and form are not clear. It has been

shown that estrogen accelerates and P decelerates oviduct egg

transport (Mahmood et al. 1998, Orihuela & Croxatto 2001,

Orihuela et al. 2001) and antiprogestins added to natural

cyclic rats accelerates ovum transport and results in premature

arrival to the uterus (Fuentealba et al. 1987). More recently,

Wessel et al. (2004) used explants of bovine oviduct to show

that P regulates ciliary beat frequency by a fast, non-genomic

hormonal interaction. Ciliary beat frequency in the different

parts of the oviduct is regulated by Ca2C (Verdugo 1980), and

ciliary beat frequency changes during the natural cycle and

during pregnancy (Lyons et al. 2002). We suggest that ciliary

PR directly modulates the ciliated oviduct epithelium by

operating as a fast means to sense and relay changes in the

levels of P in the oviduct, such as those induced through

release of follicular fluid at ovulation or released by COCs.

Cumulus cells from ovulated COCs are known to produce

and secrete large amounts of P (Vanderhyden & Macdonald

1998), and thus we speculate that these signaling pathways

involve specific Ca2C-regulated functions in these cells.

Recent studies of the mouse oviduct have shown that the

Ca2C-selective TRP ion channel, polycystin-2, and the

Ca2C-binding receptor protein, polycystin-1, are highly

upregulated in the cilia along the entire oviduct upon hCG

stimulation (Teilmann et al. 2005). In addition, the Ca2C

permeable cation channel gated by thermal and osmotic

stimuli, TRP vanilloid 4 (TRPV4), localizes to a

subpopulation of motile cilia on epithelial cells of the

ampulla and isthmus (Teilmann et al. 2005), and the TRPV4

channel was suggested to be involved in the coupling of

fluid viscosity changes to oviduct epithelial ciliary activity

(Andrade et al. 2005). Together, these findings favor a model

where ciliated oviduct epithelial cells perceive signals from

the extracellular milieu in a previously unappreciated

manner. Thus, ciliary signaling components such as

membrane-associated P receptors and TRP ion channels



Journal of Endocrinology (2006) 191, 525–535 www.endocrinology-journals.org


may act in concert to co-ordinate uptake, transport, and

fertilization of the gamete.

Western blotting analysis of isolated infundibulum

suggests that PR-A is the principal PR form undergoing

transcriptional and/or posttranslational changes in this part

of the oviduct. Cyclic changes in PR localization in the

uterus and oviduct can easily be appreciated by immuno-

histochemistry during the estrous cycle, but when analyzed

by immunoblotting only small changes in total PR

expression are observed (Ohta et al. 1993, Gava et al.

2004). Changes in the relative amount and distribution of

the two PR isoforms can alter the signaling capacity

dramatically and both PR isoforms undergo phosphoryl-

ation upon ligand binding that results in increased sensibility

to changes in the levels of P (Lange 2004). Phosphorylation

of PR may also facilitate subcellular relocalization (Qiu &

Lange 2003, Lange 2004). We suggest that the PR band

observed migrating just above PR-A in SDS-PAGE analysis

of oviduct protein fractions, and emerging specifically after

hCG stimulation, may represent a phosphorylated and

sensitized/activated form of PR-A as described previously

(Sheridan et al. 1989, Denner et al. 1990, Poletti et al. 1993,

Takimoto & Horwitz 1993).

We also investigated the expression and localization of PR

in the ovary and confirmed previous findings that both

nuclear and cytoplasmic PR localization in theca and

granulosa cells of large preovulatory follicles greatly increases

upon administration of LH (Iwai et al. 1991, Park & Mayo

1991, Robker et al. 2000, Jo et al. 2002). PR mRNA

expression was suggested to be confined to mural granulosa

cell compartment (Conneely et al. 2003) and no PR

expression was reported in isolated mouse cumulus cells

immediately after isolation (Conneely et al. 2001). In

agreement with these studies, we find that PR localization

in preovulatory follicles is restricted to mural granulosa cells.

In our study, luteinizing granulosa cells displayed less staining

than non-luteinized granulosa cells 6 h after hCG. Induction

of PR isoforms in granulosa cells upon the preovulatory

gonadotropin surge have been reported in bovine (Jo et al.

2002), mouse (Shao et al. 2003, Gava et al. 2004), rat (Park

& Mayo 1991), porcine (Machelon et al. 1996, Slomczynska

et al. 2000), and in the primate (Hild-Petito et al. 1988,

Figure 4 Immunolocalization of progesterone receptor with anti-PR (Sstimulation of pubertal female C57Bl/6J mice. (A) Control ovary (no stimafter additional stimulation with hCG (TZ48C6). Cilia are marked withScale bars: 25 mm. (C) Close up of the ciliated oviduct epithelium of stia-tubulin (red, arrow heads) and nuclei were detected with TO-PRO (blu16 h after additional stimulation with hCG (TZ48C16). Cilia are markedmarked with asterisk and nuclei are detected with propidium iodide (reanti-PR showing the expression of progesterone receptor forms, PR-A (cgonadotropic stimulation. Molecular markers (kDa) are shown to the leupper protein band (hatched bars), and PR-B (solid bars) in oviduct aftepubertal controls (TZ0). (G) SDS-PAGE and western blot analysis with a(ca. 83 kDa) and PR-B (ca. 115 kDa), in uterus before and after gonadotrRelative levels of PR-A (open bars) and PR-B (solid bars) protein bands iwith pubertal controls (TZ0). (F and H) Error bars indicate standard erroare marked with one, two and three asterisks for P!0.05, P!0.01, an


Sheridan et al. 1989, Iwai et al. 1990, Suzuki et al. 1994).

Although species differences clearly exist, some PR

expression in granulosa cells appears to sustain in the corpus

luteum. Our western blot analysis of ovary protein fractions

supported the immunohistochemical findings in that PR-A

and PR-B forms are highly upregulated in ovaries in mice

after hCG. Further, we observed a shift in molecular mass

for PR-A in western blot analysis of ovary extracts upon

hCG treatment, but this shift was less prominent compared

with that observed in the oviduct. Most interestingly, the

protein level of ovarian PR-A is reduced in mice treated

with FSH and prior to stimulation with hCG compared

with the non-stimulated mice. To our knowledge, these

observations are the first to show a downregulation of

ovarian PR specifically upon FSH stimulation, suggesting

additional roles of PR in preovulatory follicle development.

The mechanism by which ovarian PR is suppressed by FSH

remains to be determined.

It is now clear that P in many cell types may act via non-

genomic pathways (Luconi et al. 2002, Peluso et al. 2003,

Wessel et al. 2004) and in non-luteinized granulosa cells P

stimulation rapidly increases intracellular levels of Ca2C via

membrane receptors (Machelon et al. 1996). The increased

non-nuclear PR localization in granulosa cells of preovula-

tory follicles could therefore represent membrane-bound PR

forms that act through Ca2C signaling and possibly in

coordination with Ca2C channels (Teilmann et al. 2005) and

Ca2C currents (Asem et al. 2002). In rat ovaries, P was

suggested to inhibit granulosa cell apoptosis through the

binding and activation of a 60 kDa membrane protein that

functions as a low-affinity, high-capacity receptor for P

(Peluso et al. 2001, Peluso 2004). It remains an open question

whether the ca. 60 kDa anti-PR immunoreactive protein

observed in some western blot analysis of granulosa cells in

preovulatory follicles (data not shown) represents a mem-

brane-bound PR form.

In conclusion, PR principally localizes to the lower half

and at the base of the cilium in ciliated oviduct epithelium,

whereas non-ciliated epithelial cells primarily show nuclear

receptor localization. In contrast, ciliated mural granulosa

cells uniquely express cytosolic and nuclear PR. Further, cell-

specific localization and expression of PR are highly regulated

P2; green) in oviduct tissue sections before and after gonadotropiculation; TZ0); (B) Forty-eight hours after FSH stimulation and 6 harrow heads and nuclei are detected with propidium iodide (red).

mulated mice (TZ48C6). Cilia are detected with anti-acetylatede). Scale bar: 5 mm. (D) Forty-eight hours after FSH stimulation andwith arrow heads, cumulus granulosa cells from ovulated ovary are

d). Scale bars: 25 mm. (E) SDS-PAGE and western blot analysis witha. 83 kDa) and PR-B (ca. 115 kDa), in oviduct before and afterft. (F) Relative levels of PR-A lower protein band (open bars), PR-Ar 48 h FSH (TZ48) and 6 h of hCG (TZ48C6) compared withnti-PR showing the expression of progesterone receptor forms, PR-Aopic stimulation. Molecular markers (kDa) are shown to the left. (H)n uterus after 48 h FSH (TZ48) and 6 h hCG (TZ48C6) comparedrs from three separate experiments. Significant changes in PR levelsd P!0.001 respectively.



by gonadotropins, in which P released around the time of

ovulation may act directly through the ciliated oviduct

epithelium to immediate receptivity of the ovulated eggs.

Acknowledgements

This work was supported by EU project QLRT-2000-00305

(A G B), the Danish Medical Research Science Council 22-

02-0233 (A G B), the NOVO Foundation (S T C), The

Danish Research Science Council (S T C). The authors

declare that there is no conflict of interest that would

prejudice the impartiality of this scientific work.

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Received in final form 22 August 2006Accepted 23 August 2006Made available online as an Accepted Preprint25 September 2006


SPECIAL ISSUE RESEARCH ARTICLE

Characterization of Primary Cilia andHedgehog Signaling During Development ofthe Human Pancreas and in Human PancreaticDuct Cancer Cell LinesSonja K. Nielsen,1 Kjeld Møllgård,2 Christian A. Clement,1 Iben R. Veland,1 Aashir Awan,1

Bradley K. Yoder,3 Ivana Novak,1 and Søren Tvorup Christensen1*

Hedgehog (Hh) signaling controls pancreatic development and homeostasis; aberrant Hh signaling isassociated with several pancreatic diseases. Here we investigated the link between Hh signaling andprimary cilia in the human developing pancreatic ducts and in cultures of human pancreatic ductadenocarcinoma cell lines, PANC-1 and CFPAC-1. We show that the onset of Hh signaling from humanembryogenesis to fetal development is associated with accumulation of Hh signaling components Smo andGli2 in duct primary cilia and a reduction of Gli3 in the duct epithelium. Smo, Ptc, and Gli2 localized toprimary cilia of PANC-1 and CFPAC-1 cells, which may maintain high levels of nonstimulated Hh pathwayactivity. These findings indicate that primary cilia are involved in pancreatic development and postnataltissue homeostasis. Developmental Dynamics 237:2039–2052, 2008. © 2008 Wiley-Liss, Inc.

Key words: primary cilia; pancreas; exocrine duct; development; cancer; Hedgehog signaling; Smoothened; Patched;Gli2; Gli3

Accepted 9 May 2008

INTRODUCTION

Primary cilia are microtubule-basedorganelles that project into the extra-cellular environment from the mothercentriole of most quiescent cells in thehuman body. Recent research hasdemonstrated that cilia function assensory units that coordinate a vari-ety of signal transduction pathwaysessential for embryonic and postnataldevelopment as well as tissue ho-meostasis in the adult body (Chris-tensen et al., 2007; Eggenschwiler andAnderson, 2007; Kiprilov et al., 2008).

These pathways include Hedgehog(Hh), Wnt, and platelet-derivedgrowth factor receptor (PDGFR) sig-naling as well as Ca2�-signaling bymeans of TRP ion channels. Further-more, primary cilia may control be-havioral responses, such as occurs inthe central nervous system whereneuronal cilia function in a pathwaythat controls satiety responses (Dav-enport et al., 2007). Consequently, de-fects in ciliary assembly lead to aplethora of diseases and disorders, in-cluding kidney, liver and pancreatic

cysts, blindness, obesity, diabetes, de-velopmental disorders, and other hu-man diseases now collectively referredto as ciliopathies (Badano et al., 2006;Satir and Christensen, 2007; Yoder,2007; Fliegauf et al., 2007).

Primary cilia on the epithelium of re-nal tubules control tissue homeostasisprobably by acting as mechanosensorsthat monitor the composition and flowrate of urine in the nephron (Praetoriusand Spring, 2003; Yoder, 2007). Ob-struction of this flow-sensing responsein, for example, the IFT88Tg737Rpw

1Department of Biology, University of Copenhagen, Copenhagen, Denmark2Department of Cellular and Molecular Medicine, The Panum Institute, University of Copenhagen, Copenhagen, Denmark3Department of Cell Biology, University of Alabama at Birmingham, Birmingham, AlabamaGrant sponsor: National Institutes of Health; Grant number: RO1 R01-HD056030.*Correspondence to: Søren Tvorup Christensen, Department of Biology, Section of Cell and Developmental Biology, TheAugust Krogh Building, University of Copenhagen, Universitetsparken 13, DK-2100 Copenhagen OE, Denmark.E-mail: [email protected]

DOI 10.1002/dvdy.21610Published online 10 July 2008 in Wiley InterScience (www.interscience.wiley.com).

DEVELOPMENTAL DYNAMICS 237:2039–2052, 2008

© 2008 Wiley-Liss, Inc.

mouse (hereafter referred to asTg737orpk), causes disruption of tissueorganization and cyst formation. TheTg737orpk mouse has a mutation inthe Tg737/IFT88 gene that encodes asubunit of the IFT particle complex Brequired for ciliary assembly (Pazouret al., 2000; Yoder et al., 2002a;Rosenbaum and Witman, 2002; Ped-ersen et al., 2008). Among the ciliarysignaling modules involved in renaltissue homeostasis are the TRP ionchannel polycystin 2 (PC-2) and poly-cystin-1 (PC-1), which are disruptedin human ADPKD (Yoder et al.,2002b; Pazour et al., 2002; Nauli etal., 2003). Bending of the ciliary axon-eme due to fluid movement has beenshown to induce a Ca2�-response(Praetorius and Spring, 2001), whichis dependent on PC-1 and PC-2. Thiscalcium response is thought to be im-portant in detecting fluid movementfor maintaining tissue function, be-cause mutations leading to its loss re-sult in cyst development (Delmas etal., 2004; Liu et al., 2005b; Kottgen,2007; Weimbs, 2007; Yoder, 2007). Inaddition, in the absence of flow PC-1may be proteolytically processed(Chauvet et al., 2004), leading to thenuclear translocation of the transcrip-tion factor STAT6 and co-activatorP100 (Low et al., 2006). Furthermore,defects in ciliary assembly may resultin up-regulation of the canonical Wntpathway and impair the ability of non-canonical Wnt signals to suppress thecanonical pathway (Gerdes et al.,2007; Corbit et al., 2008). Altered reg-ulation of the Wnt signaling pathwayin the kidney has been associated withuncontrolled cell proliferation and dif-ferentiation and was found to result incystic kidney disease (Cano et al.,2004; Merkel et al., 2007).

In addition to kidney defects, theloss of primary cilia in the Tg737orpk

mouse causes a series of abnormali-ties in the pancreas, including exten-sive cyst formation in ducts (Cano etal., 2004; Zhang et al., 2005). Thismay indicate a functional similaritybetween cilia in kidney and pancreaticduct systems. Cells of both exocrineand endocrine systems in the pan-creas possess primary cilia, includingislet cells and the ducts, but appar-ently not the acini (Kodama, 1983;Ashizawa et al., 1997; Cano et al.,2004, 2006; Zhang et al., 2005). Pan-

creas abnormalities in the Tg737orpk

mouse begin with dilations of the pan-creas ducts in late gestation, whichafter birth are accompanied by exten-sive formation of large, interconnectedcysts as well as apoptosis and vacuol-ization of acini. These changes arereminiscent of chronic pancreatitis,supporting the speculation that pri-mary cilia of ducts play an essentialrole in the development of the pan-creas (Cano et al., 2004; Zhang et al.,2005). In the dilated ducts and cysts,PC-2 is mislocalized to intracellularcompartments, the cytosolic localiza-tion of �-catenin is increased andthere is an increased expression ofTcf/Lef transcription factors. Thisfinding suggests that, as in the kid-ney, there is an alteration in the Wntsignaling pathway (Cano et al., 2004).

Another signaling pathway essen-tial for pancreatic development andtissue homeostasis is the Hedgehog(Hh) pathway. Hh regulates cell pro-liferation and differentiation in nu-merous embryonic tissues and Shh isexpressed in many regions of the em-bryo where it functions as a key orga-nizer of tissue morphogenesis (Odentet al., 1999). One embryonic tissuewhere the initial absence of Shh sig-naling is required for development isin the pancreatic anlage, because ec-topic expression of Shh leads to trans-formation of pancreatic mesoderm intointestinal mesenchyme (Apelqvist etal., 1997). In addition to being an impor-tant inductive signal in embryonic de-velopment, Hh signaling is required inhomeostasis of mature tissues and isalso implicated in many human can-cers, including endodermal derived car-cinomas of the esophagus, stomach, andpancreas (Beachy et al., 2004) as well asneurodegenerative disorders (Bak etal., 2003). Although the mechanism isnot completely understood, it has beenshown that Hh signaling is coordinatedby the primary cilium (e.g., Huangfu etal., 2003; Corbit et al., 2005; Huangfuand Anderson, 2005; May et al., 2005;Haycraft et al., 2005; Liu et al., 2005a;Koyama et al., 2007; Vierkotten et al.,2007; Ruiz-Perez et al., 2007; Casparyet al., 2007; Rohatgi et al., 2007;Kiprilov et al., 2008). Regulation of theHh pathway is complex with Hh re-sponsive Gli transcription factors hav-ing either activator or repressor func-tions that depend on IFT proteins.

Binding of Hh to its receptor Patched(Ptc) occurs in the cilium and results intranslocation of Ptc out of the ciliumand into the cell body (Rohatgi et al.,2007; Kiprilov et al., 2008). In contrast,Smoothened (Smo), the transmem-brane effector of the pathway whose ac-tivity is suppressed by Ptc is targeted tothe cilium in the presence of Hh. Thisrelieves the inhibitory effects of Ptc onSmo and controls the activation or de-activation of Gli transcription factors.In the absence of IFT or cilia, the re-pressor and activator functions of theGli transcriptional regulators becomederegulated with the resulting pheno-type in a tissue being determined bywhether it is the activator or repressorfunction of the Gli transcription factorsthat is most critical. For example, theprocessing of full-length Gli3 (Gli3FL)to repressor form (Gli3R) is affected inIFT mutants, showing the importanceof functional IFT for regulating Glitranscription factors.

In the developing and mature pan-creas, deregulated Hh signaling re-sults in a series of diseases, includingannular pancreas, diabetes mellitus,chronic pancreatitis and pancreaticcancer (Lau et al., 2006). During theearly mouse embryonic stages, Hh sig-naling is kept at a low level in thepancreatic anlage to ensure the cor-rect establishment of organ bound-aries, as well as tissue architecture.At later developmental stages, Hh sig-naling is activated to promote prolif-eration and maturation of the tissue(Kawahira et al., 2005; Lau et al.,2006; Cano et al., 2007; van denBrink, 2007). In the mature pancreas,Hh signaling maintains endocrinefunctions, and aberrant Hh signalingcauses pancreatic ductal adenocarci-noma and chronic pancreatitis(Thayer et al., 2003; Berman et al.,2003; Kayed et al., 2003, 2006; Mortonet al., 2006; Cano et al., 2007).

The aim of our study was to exam-ine the expression of primary cilia inhuman pancreatic ducts and to inves-tigate the sensory competence of pri-mary cilia with regards to the Hh sig-naling pathway during pancreaticdevelopment and in pancreatic adeno-carcinoma cell lines, CFPAC-1 andPANC-1, which are isolated from met-astatic and primary tumors, respec-tively (Lieber et al., 1975; Schouma-cher et al., 1990). Initially, we show

2040 NIELSEN ET AL.

that localization of Hh signaling com-ponents is regulated during humanpancreatic development with Gli2 andSmo accumulating in the cilium atlater stages while the level of nuclearand cytosolic Gli3 expression is re-duced. These changes in localizationcorrelate with known activity of theHh pathway during pancreas develop-ment. We demonstrate that compo-nents of the Hh pathway also localizeto the cilium in CFPAC-1 and PANC-1cells and that these cells may maintainhigh levels of nonstimulated Hh path-way activity. These findings indicatethat Hh signaling coordinated by theprimary cilium in the pancreas is essen-tial for normal pancreatic developmentas well as postnatal tissue function.

RESULTS

Early Development of theHuman Pancreas

To investigate the presence of primarycilia in human pancreatic progenitorcells of the initial duct epithelium weanalyzed 3-�m-thick tissue sectionsfrom 7.5-week-old embryos and from14- and 18-week-old fetuses. Hema-toxylin and eosin (HE) staining of thepancreatic tissue and identification ofthe developing ducts at these stagesare presented in Figure 1A–C. In 5- to6-week-old embryos a dorsal and aventral out pocketing of the foregutduodenal endoderm result in the for-mation of separate dorsal and ventralpancreatic primordia. During the sev-enth embryonic week a fusion of thetwo epithelial pancreatic ducts takesplace followed by a rapid expansionand branching of the ductal epithe-lium (Fig. 1A). Pancreatic progenitorcells in this epithelium engage in twoseparate differentiation programs atdistinct developmental stages. At thefirst stage from week 8 to 10, the en-docrine progenitor cells appear as cellclusters budding from the central duc-tal epithelium. These buds form theislets of Langerhans in the centralpancreas at week 12, and at midges-tation, the endocrine compartment isestablished. At a much later stage, theexocrine progenitors are responsiblefor the differentiation and expansionof the exocrine acinar epithelium,which is mainly a late fetal or earlyperinatal event. Figure 1B shows a

section of the ductal system from a14-week-old fetus (Jackerott et al.,2006) with initial endocrine progeni-tors budded from the ductal epithelium,and Figure 1C shows a section from an18-week-old fetus with endocrine pro-genitors budded from the ductal epithe-lium as well as acinar progenitors sur-rounding the ductal epithelium.

Characterization of PrimaryCilia in Ducts of theDeveloping Human Pancreas

To identify primary cilia, we used twomarkers for cilia (anti-acetylated�-tubulin [acet. tb] and anti-detyrosi-nated �-tubulin [glu-tub]) in immuno-fluorescence microscopy (IF) analysisand found that epithelial cells in ductsat all stages form primary cilia thateither project into the duct lumen,form spiral-like structures or alignparallel to the lumen surface (Fig.1D–H). The cilia varied in lengthsfrom approximately 5 to 20 �m, and insome ducts long cilia seemed to formcontact or bridge with cilia fromneighboring epithelial cells or fromcells on the opposite side of the duct(Fig. 1E,F). At the late stage of embry-onic development (week 7.5), longacetylated-tubulin structures oftenprojected from epithelial cells in a“star”-like configuration by pointinginto the site of the developing duct(Fig. 1F and inset). Both short andlong primary cilia were observed dur-ing all three developmental stages.Long cilia were predominantly ob-served in week 7.5, possibly due to theprevalence of small intercalated andintralobular type ducts that do havelonger cilia (Kodama, 1983). The for-mation and orientation of primarycilia in the human developing ducts issimilar to that reported for the pan-creas in the rat and mouse (e.g., Ko-dama, 1983; Hidaka et al., 1995; Ash-izawa et al., 1997; Cano et al., 2004;Zhang et al., 2005).

Development of the HumanPancreas Correlates WithCiliary Localization of HhSignaling Components inDuct Epithelial Cells

It was previously shown that thegraded response to Hh-signaling con-

trols pancreatic organogenesis in themouse, where Hh signaling is at lowlevels during early embryonic stagesto ensure the correct establishment oforgan boundaries and tissue architec-ture. Hh signaling is then activated atlater developmental stages to promoteproliferation and maturation of thetissue (Kawahira et al., 2005; Lau etal., 2006; Cano et al., 2007; van denBrink, 2007). In Hh signaling, Gli2 invivo acts primarily as a transcrip-tional activator, whereas Gli3 mainlyworks as a transcriptional repressorto control development. To addressthe potential role of primary cilia inhuman pancreatic development, weinvestigated the expression and cellu-lar distribution of Smo, Gli2, and Gli3in developing pancreatic ducts by IFanalysis. In other cell types, such asfibroblasts (Rohatgi et al., 2007) andhuman embryonic stem cells, hESC(Kiprilov et al., 2008), Smo becomesconcentrated while Ptc levels decreasein primary cilia upon stimulation witha Hh ligand, which may result in ac-tivation or deactivation of Gli tran-scription factors, possibly in the ci-lium (Huangfu and Anderson, 2006;Christensen and Ott, 2007). We foundthat Smo (Fig. 2A) and Gli2 (Fig. 2B)are absent from primary cilia in pan-creatic ducts of embryos (week 7.5),but are concentrated in the cilia ofducts of early fetuses (week 14), andeven more so in 18-week-old fetuses.We then investigated the localizationof Gli3 in IF analysis using two differ-ent Gli3 antibodies that were raisedagainst amino acids 1-280 of Gli3(Gli3-H-280) and against a peptidemapping at the N-terminus of Gli3(Gli3-N-19). Both antibodies showed adecrease in staining intensity and cel-lular distribution of Gli3 during devel-opment. At week 7.5, Gli3 strongly lo-calized to duct epithelial cells and waspresent in the nuclei and in an intensepunctate pattern in the cytosol of ductepithelial cells. In contrast, in ducts of14- and 18-week-old fetuses, Gli3staining was reduced (Fig. 2C–E). Asa control, a blocking peptide againstanti-Gli3-N-19 decreased localizationof the antibody to the duct epithelium(Fig. 2D). These results could reflectactivation of the pathway because Hhis known to repress Gli3 expression(Marigo et al., 1996; Buscher andRuther, 1998; Schweitzer et al., 2000).

PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREAS 2041

Characterization of PrimaryCilia in Cultures of HumanPancreatic Duct Cell Lines

CFPAC-1 and PANC-1 are humanpancreatic ductal adenocarcinoma celllines, which form adherent epithelialmonolayers in culture (Fig. 3A,B) andare often used as model cell lines forstudying ion transport and signaling

in human pancreas (Novak et al.,2008). PANC-1, derived from epithe-liod carcinoma, forms a monolayer ofheterogeneous duct cells also withsome cells growing on top of eachother. Actin is mainly arranged in thecortex and lamellipodia (Fig. 3B). CF-PAC-1 cells, derived from a patientwith cystic fibrosis containing deletionin Phe-508 in CFTR, appear homoge-

nous and on glass surface they form amonolayer interrupted by some voidspaces. In most CFPAC-1 cells actin isconcentrated in well-organized stressfibers and no subcortical actin net-work is apparent (Fig. 3A). Also thecytosolic network of microtubulesseems dispersed throughout the cells,as also observed by Hollande et al.(2005). The epithelial-like cytoskeletal

Fig. 1. Characterization of primary cilia in human pancreatic progenitor cells of the initial ductal epithelium. A–C: Hematoxylin and eosin (HE) stainingof sections of the human developing pancreas from 7.5-week-old embryos (A), 14-week (B), and 18-week-old (C) fetuses. Lower panels show highermagnification images of the developing ducts. d, ducts; mPD, main pancreatic duct; PI, pancreatic islets budding from the duct (d); D, duodenum. D–H:Immunofluorescence microscopy (IF) analysis of primary cilia in the developing ducts at the stages presented in (A–C) by using two markers for primarycilia (arrows; anti-acetylated �-tubulin, acet. tb, red; and anti-detyrosinated �-tubulin, glu-tub, green). Nuclei were stained with DAPI (blue,4�,6-diamidine-2-phenylidole-dihydrochloride). Shifted overlays (E) shows the combined staining with acet. tb and glu-tb, where the images for eachantibody are slightly shifted from one another. Scale bars � 50 �m in A–D, 10 �m in E,G,H, 20 �m in F.

2042 NIELSEN ET AL.

arrangement of PANC-1 cells may aidthe targeting and function of adeno-sine receptors, which regulate iontransport (Novak et al., 2008). Thedistribution of these receptors, car-bonic anhydrase, and cytoskeleton ar-rangement is defective in CFPAC-1cells (Novak et al., 2008).

To study the time course of primarycilia formation the cells were analyzedby IF microscopy both in cultures with10% serum (at 50% and 100% conflu-ency) and following serum starvationof confluent cultures in medium with0.5% serum for periods of 48 and 72 hr(Fig. 3G). At 50% confluency and inthe presence of 10% serum PANC-1

cells are most often in interphasegrowth and mitosis, such that veryfew cells are ciliated. At 100% conflu-ency, approximately 4% of the cellshad formed single primary cilia, pre-sumably as a consequence of cellularcontact inhibition of growth. After 48and 72 hr of serum starvation, thefrequency of ciliated cells increased toapproximately 12 and 46%, respec-tively, and cilia had lengths of approx-imately 10–20 �m (Fig. 3D, left panel,and 3G). Similar results on appear-ance of cilia were obtained with cul-tures of CFPAC-1 cells; these cells,however, started to loose contact withthe polylysine-coated glass coverslips

at 72 hr of serum starvation. The rel-atively low percentage of ciliated cellsin pancreatic cell lines is in sharp con-trast to that of other cell types such ascultured fibroblast, in which morethan 95% of the cells are ciliated after24 hr of serum starvation (Schneideret al., 2005). The low frequency of cil-iated cells in the adenocarcinoma celllines could be related to their cancer-ous origin because cells that have anincreased rate of growth more rarelyenter growth arrest, which is requiredfor assembly and maintenance of theprimary cilium (Satir and Chris-tensen, 2007). Indeed, IF analysis re-vealed many mitotic and dividing cells

Fig. 2. Expression of Hedgehog signal components in human pancreatic progenitor cells of the initial ductal epithelial cells and their primary cilia from7.5-week-old embryos and 14-week and 18-week-old fetuses. A–E: Localization of Smo (green) in 7.5-week-old embryos and 18-week-old fetuses (A),and localization of Gli2 (H-300; green, B), Gli3 (H-280, green, C), and Gli3 (N-19, green, D,E) in 7.5-week-old embryos and 14-week and 18-week-oldfetuses. Primary cilia (arrows) were localized with acet. tb (red) and nuclei were stained with DAPI (blue, 4�,6-diamidine-2-phenylidole-dihydrochloride).d, ducts. Scale bars � 10 �m.


in the CFPAC-1 and PANC-1 culturesas evidenced by the presence of mi-totic spindles and midbodies (Fig.3C,D, right panels). As in other celltypes, the primary cilia emanate fromthe centrosomes (labeled by anti-Pctn,Fig. 3E,F left panels) and, in particu-lar, from one of the two centrioles (la-beled by anti-centrin, Fig. 3E,F rightpanels), presumably the mother cent-riole, which also functions as the basalbody (Satir and Christensen, 2007).PANC-1 cells grew longer cilia (10–20�m) compared with CFPAC-1 cells

(5–15 �m), where in some cases theshort cilia projected vertically into themedium, visualized as “dots” in the IFanalysis (Fig. 3C, middle panel).

Localization of Ptc and Smoto Primary Cilia of HumanPancreatic Duct Cell Lines

To evaluate whether Hh-signalingand primary cilia might be linked todevelopment of cancer in adult pan-creas, we next investigated whethercomponents of the Hh signaling sys-

tem are present in CFPAC-1 andPANC-1 primary cilia. Initially, wedemonstrate by IF analysis that as inother cell types, Ptc and Smo localizedto primary cilia of both cell lines (Fig.4A,F). As a control, we exogenouslyexpressed Smo from a YFP-Smo con-struct and show that it localized to thecilia of CFPAC-1 cells (Fig. 4G). ThePtc antibody recognized Ptc as a sin-gle protein band in Western blot anal-ysis (Fig. 4B, upper panel), and thisband was removed in the presence ofanti-Ptc blocking peptide (Fig. 4C).

Fig. 3. Characterization of primary cilia in cultures of human pancreatic adenocarcinoma cells. A,B: Adherent epithelial monolayer cultures ofCFPAC-1 (A) and PANC-1 (B). Cells were either double-labeled with DAPI (blue, 4�,6-diamidine-2-phenylidole-dihydrochloride) and phalloidin (red) fornuclear and F-actin staining, respectively, or triple-labeled with DAPI (blue), phalloidin (red) and tubulin (green) (insets). C,D: Formation of primary cilia(bold arrows, acet. tb, red) in quiescent CFPAC-1 (C) and PANC-1 (D), i.e., serum-starved for 48 and 72 hr, respectively. Acet. tub (red) also stainedmitotic spindles (*: metaphase cells; ¤: anaphase cells) and midbodies of cells that have recently undergone cytokinesis (open arrows). Nuclei werestained with DAPI (blue). E,F: Primary cilia (acet. tub, red, arrows) in CFPAC-1 (E) and PANC-1 (F) serum-starved for 48 and 72 hr, respectively, emergefrom the centrosome marked with anti-pericentrin at the base of the cilium (Pctn, green, #, left panels) and in particular from one of the centriolesmarked with anti-centrin (centrin, green, *, right panels). Nuclei were stained with DAPI (blue). G: Frequency of ciliated PANC-1 cells in the presenceof 10% serum at 50 and 100% confluency and in 100% confluent cultures after 48 and 72 hr of serum starvation (0.5% serum). Scale bars � 100 �min A,B, 10 �m. in E,F.

2044 NIELSEN ET AL.

Expression of Ptc at the mRNA levelwas previously shown to be up-regu-lated as a consequence of the autono-mous operation of an active Hh signal-ing process in pancreatic cancer cellsin which Ptc mRNA in PANC-1 cells isup-regulated at a higher level thanthat of CFPAC-1 cells (Berman et al.,2003; Thayer et al., 2003). Consistentwith these earlier findings, we showthat the protein level of Ptc in PANC-1was higher than that in CFPAC-1cells (Fig. 4B, upper and lower pan-els). As further controls for antibodyspecificity, we show that Ptc localizedto primary cilia of wild-type mouseembryonic fibroblasts (wt MEFs) butnot to mutant Ptc�/� MEFs (Fig. 4D)as previously demonstrated by Ro-hatgi et al. (2007). Then we stimu-lated wt MEFs with the Smo agonist,SAG, that activates Hh-signaling(Chen et al., 2002) to analyze changesin protein levels of Ptc, which is up-regulated in response to activation ofthe Hh pathway. As shown in Figure4E, the Ptc is present at a low level inwt MEFs in the absence of SAG,whereas it increased after Hh path-way activation. This increase was de-pendent on the presence of the pri-mary cilium, because the level of Ptcwas kept at a low level in the presenceof SAG in MEFs derived from theTg737orpk mouse (Fig. 4E). These re-sults support the idea that the ciliumis required for activation of the Hhpathway, and that aberrant Hh sig-naling in pancreatic cancer may beassociated with increased Hh signal-ing by means of the primary cilium.

Another observation was that Ptclocalized only weakly to the primarycilium (Fig. 4A), while Smo localizedvery strongly and in a punctate pat-tern in the cilium of the pancreaticduct cancer cells (Fig. 4F). This is insharp contrast to that observed in cul-tures of normal cells such as MEFsand hESCs, where Ptc is highly con-centrated in the cilium and Smo ismostly absent from the cilium in non-stimulated cells (Rohatgi et al., 2007;Kiprilov et al., 2008). Stimulation ofthe pancreatic duct cells with SAG didnot further increase ciliary localiza-tion of Smo in CFPAC-1 cells (Fig.4H), supporting the conclusion thatthe Hh pathway is maintained at ahigh and autonomous level in thesecells.

Localization of GliTranscription Factors toPrimary Cilia of HumanPancreatic Duct Cell Lines

Gli2 and Gli3 are the primary tran-scription factors that are being regu-lated in Hh signaling. The full-lengthtranscription factors mainly functionas transcriptional activators, but inthe absence of the Hh signaling theymay undergo proteolytic processingand function as transcriptional re-pressors (Wang et al., 2000; Liting-tung et al., 2002; Pan et al., 2006). Toinvestigate the expression and local-ization of Gli2 in growth-arrestedPANC-1 and CF-PAC-1 cells we usedthree different antibodies raisedagainst amino acids 841-1140 map-ping near the C-terminus of Gli2(Gli2-H-300), a peptide mapping nearthe N-terminus of Gli2 (Gli2-N-20),and a peptide mapping within an in-ternal region of Gli2 (Gli2-G-20). Allthree antibodies localized uniquely tothe primary cilium and most often asan enrichment at the tip of the cilium(Fig. 5A,B, left panels), similar towhat has been observed in primarycilia of the murine limb bud and hESC(Haycraft et al., 2005; Kiprilov et al.,2008). Ciliary localization was abol-ished in the presence of blocking pep-tides to the two peptide antibodies(Fig. 5B, right panels). Also, localiza-tion of Gli2 to the ciliary tip was con-firmed by transfection and overex-pression of a green fluorescent protein(GFP) -Gli2 construct in NIH3T3 fi-broblasts (Fig. 5C). To confirm anti-body specificity, we performed West-ern blotting analysis with the peptideantibodies in the absence and in thepresence of blocking peptides. Asshown in Fig. 5D, Gli2-N-20 specifi-cally recognized both full-length (ca.170 kDa) and processed (ca. 70 kDa)forms of Gli2 that function as activa-tor (Gli2(FL)) and repressor (Gli2(R))forms in Hh signaling, respectively(Pan et al., 2006). In contrast, Gli2-G-20 recognized only Gli2(FL) (Fig.5E). This finding suggests that the ac-tivator forms of Gli2 are present in theprimary cilium and that ciliary Gli2may be part of the Hh signaling ma-chinery that is up-regulated in pan-creatic cancer cells.

We then analyzed lysates from CF-PAC-1 cells by Western blotting anal-

ysis using the Gli3 antibodies, Gli3-N-19 and Gli3-H-280. Gli3-N-19recognized both full-length (Gli3(FL)at ca. 180 kD) and processed forms(Gli3(R) at ca. 80 kDa) of Gli3 (Fig.5F,G). In contrast, Gli3-H-280 pre-dominantly recognized the processedversion of Gli3 (Fig. 5F), supportingthe conclusion that this antibodymainly identifies the repressor form ofGli3 under the experimental condi-tions carried out in this work. Thesedata further support the assumptionthat the observed decrease in stainingintensity and cellular distribution ofGli3 in the epithelium of the pancre-atic ducts during fetal development(Fig. 2) is due to the increased activa-tion of the Hh signaling that favorsthe processing of transcriptional acti-vators of the Hh pathway. Based onthese observations, it was thereforeinteresting to investigate the expres-sion and localization of Gli3(R) in pan-creatic cancer cells upon stimulationof the Hh pathway using Gli3-H-280in IF analysis. As seen in Figure 5H,Gli3-H-280 stained CFPAC-1 cellsvery weakly in both the absence andin the presence of SAG. In contrast,Gli3-H-280 strongly stained the nu-cleus as well as small cytoplasmicpuncta in NIH3T3 cells and this stain-ing was largely absent in the presenceof SAG (Fig. 5I). Furthermore, weused transfected NIH3T3 cells withGFP constructs of either the GFP-Gli3(R) and Gli3(FL) to evaluate thecellular localization of activator andrepressor forms of Gli3. Similar tothat seen in nonstimulated NIH3T3cells with Gli3-H-280 (Fig. 5I) andpreviously in mouse limb bud mesen-chyme cells, we found that GFP-Gli3(R) localized predominantly to thenucleus and to small cytosolic puncta,but not to the primary cilium (Fig. 5J,left panels). In contrast, GFP-Gli3(FL)seemed to localize to the primary ci-lium with minimal localization in thenucleus and without cytosolic puncta(Fig. 5J, right panels). These resultsare in line with our conclusion thatGli3-H-280 recognizes the repressorform of Gli3, which is concentrated inthe nucleus in the absence of Hh sig-naling, and that Gli3-repressor formsare expressed at a low level in CF-PAC-1 cells.


Fig. 4.

Fig. 5.

Fig. 4. Localization of Patched (Ptc) andSmoothened (Smo) in cultures of quiescent hu-man pancreatic adenocarcinoma cells. Primarycilia are marked with arrows and were localizedwith acet. tb (red). A: Localization of Ptc (green)in PANC-1 cells (upper panels) and CFPAC-1cells (lower panels). B: The upper panel showssodium dodecyl sulfate-polyacrylamide gelelectrophoresis, SDS-PAGE, and Western blot-ting analysis of anti-Ptc cross-reactivity to pro-teins in CFPAC-1 and PANC-1 cells. The lowerpanel shows protein relative levels of Ptc inCFPAC-1 and PANC-1 cells. Values are givenrelative to the level of Ptc in CFPAC-1 cells (n �3). C: Western blotting analysis of anti-Ptccross-reactivity in the absence and in the pres-ence of blocking peptide in PANC-1 cells. D:Localization of Ptc (green) to the primary ciliumin wild-type mouse embryonic fibroblasts (wtMEFs, upper panels) and in mutant Ptc�/�MEFs (lower panels). E: Protein levels of Ptcbefore and after stimulation with SAG for 24 hrin quiescent wt and Tg737orpk MEFs. F: Local-ization of Smo (green) in PANC-1 cells (upperpanels) and CFPAC-1 cells (lower panels). G:Localization of YFP-Smo (green) in CFPAC-1cells. Localization of Smo following 24 hr SAGstimulation in CFPAC-1 cells. Scale bars � 10�m.

2046 NIELSEN ET AL.

DISCUSSION

Recent research has shown that pri-mary cilia coordinate signal path-ways, which are essential in mamma-lian development, tissue homeostasisand behavioral responses. Accord-ingly, defective primary cilia assemblyand maintenance lead to a plethora ofdiseases and disorders, now collec-tively referred to as ciliopathies(Badano et al., 2006; Fliegauf et al.,2007; Satir and Christensen, 2007;Yoder, 2007). Previous reports haveshown that loss of primary cilia in theTg737orpk mouse causes a series of de-velopmental defects in exocrine andendocrine tissues of the pancreas, in-cluding extensive cyst formation inducts (Cano et al., 2004; Zhang et al.,2005). However, the function of pri-mary cilia in pancreatic organogenesisand adult tissue homeostasis islargely unknown. In this report, wedemonstrate that essential compo-nents of the Hedgehog (Hh) signalingpathway localize to primary cilia ofboth human pancreatic progenitorcells of the initial ductal epitheliumand in cultures of human pancreaticadenocarcinoma duct cell lines, andthat expression of ciliary Hh compo-nents may be linked to pancreatic or-ganogenesis and pancreatic cancer.

Hh Signaling and PrimaryCilia in the DevelopingHuman Pancreas

Previous studies have shown that thegraded response to activation of theHh signaling pathway is critical inpancreatic organogenesis in themouse, where Hh signaling is acti-vated only at late developmental

stages (Kawahira et al., 2005; Lau etal., 2006; Cano et al., 2007; van denBrink, 2007). It is likely that organo-genesis of the human pancreas alsodepends on the graded response to Hhsignaling, because the transcriptionalnetwork that controls pancreatic de-velopment is highly conserved inmammals (Wilson et al., 2003), al-though the time point of onset of de-velopment and maturation of the en-docrine system differs in humans androdents (Ostrer et al., 2006). Recently,the primary cilium was shown to actas a sophisticated switch by whichcells turn Hh signaling on and off bythe regulated movement of Smo intothe cilium and Ptc out of the ciliumupon stimulation of the Hh pathwayin fibroblasts (Rohatgi et al., 2007)and hESC (Kirprilov et al., 2008),where Smo probably functions to acti-vate Gli transcription factors (Chris-tensen and Ott, 2007). To analyze thesignificance of the primary cilium inHh signaling during organogenesis ofthe human pancreas, we studied sec-tions of the developing human pan-creas at different developmentalstages. These stages included embry-onic stage 7.5 weeks, which is beforedevelopment of the endocrine system,and stages 14 and 18 weeks, whereendocrine progenitors bud from theductal epithelium to form the endo-crine compartment (Fig. 1). Based onthe recent data from Rohatgi et al.(2007), our findings that Smo and Gli2are absent from pancreatic primarycilia at embryonic stage 7.5, buthighly concentrated in cilia in 14- and18-week-old fetuses (Fig. 2) suggestthat the Hh pathway has become ac-

tivated during developmental stages.This further supports the conclusionthat development of the human pan-creas is regulated by a graded Hh sig-naling response, and that this could becoordinated by the primary cilium. In-deed, the loss of Gli3 staining in theductal epithelium after week 7.5, thatis, before formation of the endocrinesystem, is consistent with the inter-pretation that Gli3 staining at week7.5 may represent the up-regulatedlevel of the repressor form of Gli3 (Fig.5F), consistent with low level of Hhsignaling at this developmental stage.Therefore, disruption of pancreatic de-velopment in mice with defects in pri-mary ciliary assembly (Cano et al.,2004, 2006; Zhang et al., 2005), maypartly be due to loss of coordinated Hhsignaling during genesis of the pan-creas.

Another interesting observation isthe presence of long cilia-like struc-tures that seem to bridge with corre-sponding structures emerging fromadjacent or opposite cells of the lumensurface (Fig. 1). This is particularlyprominent in the developing ducts atweek 7.5. Bridging of primary ciliawas also observed in collecting ductsof the mouse developing kidney (Liu etal., 2005b), but the function of physi-cal contact between individual pri-mary cilia is unknown. Althoughhighly speculative at this point, theintertwining of cilia could be part ofthe sensory signaling machinery thatcontrols the initial development ofducts and perhaps endocrine progeni-tors. In the developing kidney, the for-mation of tubular systems is con-trolled by inversin, which acts as a

Fig. 5. Expression and localization of Gli transcription factors in quiescent human pancreatic adenocarcinoma cells and NIH3T3 fibroblasts. Primarycilia are marked with arrows and were localized with acet. tb (red). A: Localization of Gli2 (H-300; green) to primary cilia of CFPAC-1 cells. The insetin the lower panel shows a shifted overlay of the cilium (red) and Gli3 (H-300, green). B: Localization of Gli2 (N-20) and Gli2 (G-20, both in green) inthe absence and in the presence of their corresponding blocking peptides in PANC-1 and CFPAC-1 cells. C: Localization of green fluorescent protein(GFP) -Gli2 (green) in NIH3T3 cells. D,E: SDS-PAGE and Western blotting analysis of anti-Gli2 (N-20) (D) and anti-Gli2 (G-20) (E) specificity to proteinsin CFPAC-1 and PANC-1 cells in the absence and in the presence of their corresponding blocking peptides. Anti-Gli2 (G-20) only recognized a bandat ca. 170 kDa. Abbreviations: Gli2(FL): Full length (activator) form of Gli2; Gli2(R): Processed (repressor) form of Gli2. F: SDS-PAGE and Westernblotting analysis of anti-Gli3 (N-19) and anti-Gli3 (H-280) specificity to proteins in CFPAC-1 cells. Gli3(FL): Full-length (mild activator) form of Gli3;Gli3(R): Processed (repressor) form of Gli3. Anti-Gli (N-19) recognized both Gli3 forms, while Gli3 (H-280) predominantly recognized the repressor formof Gli3. G: Western blotting analysis of anti-Gli3 (N-19) specificity in the absence and in the presence of its blocking peptide in CFPANC-1 cells.H,I: Changes in localization of Gli3 (H-280) in nonstimulated (left panels) vs. SAG-stimulated (right panels) in CFPAC-1 (H) and NIH3T3 (I) cells. Innonstimulated NIH3T3 cells, Gli3 localized similar to that of GFP-Gli3(R), but after SAG stimulation the nuclear and cytosolic levels of Gli3 weremarkedly decreased. In CF-PAC-1 cells the nuclear and cytosolic levels of Gli3 were low in both stimulated and nonstimulated cells. J: Expression oftwo different GFP-Gli3 (green) constructs in quiescent NIH3T3 cells, where the repressor form of Gli3 (GFP-Gli3(R)) strongly localizes to the nucleusand to cytosolic puncta (left panels) and the full-length form of Gli3 (GFP-Gli3(FL)) localizes to the primary cilium (right panels). Insets: Shifted overlayswith high resolution of GFP in the cilium. Nuclei were stained with DAPI (blue, 4�,6-diamidine-2-phenylidole-dihydrochloride). Scale bars � 5 �m in A,10 �m in J.


molecular switch between the canoni-cal and nonconical Wnt pathways (Si-mons et al., 2005), and inversin poten-tially signals from the primary ciliumto suppress �-catenin expression inthe canonical pathway and to up-reg-ulate the noncanonical pathway to or-ganize planar cell polarity and correcttubular formation (Christensen et al.,2007). Because loss of the primary ci-lium in the developing mouse pan-creas is associated with increased�-catenin expression (Cano et al.,2004; Zhang et al., 2005), it is possiblethat cilia in the developing pancreasregulate both Wnt and Hh signaling,although it is presently unknownwhether Hh or Wnt signaling is af-fected by physical contact betweenprimary cilia in the ductal epithelium.Further experiments are required toexplore this.

Hh Signaling and PrimaryCilia in Human PancreaticDuct Cell Lines

Hh is implicated as an important me-diator of human pancreatic carci-noma, as evidenced by, for example,the increased expression of positiveHh regulators and decreased expres-sion of negative Hh regulators in CF-PAC-1 or PANC-1 cells (Thayer et al.,2003; Berman et al., 2003). Here, weshow that essential components of theHh pathway, including Smo, Ptc, andGli2, are present in primary cilia ofhuman pancreatic ductal adenocarci-noma cell lines, CFPAC-1 andPANC-1, consistent with the idea thatthe primary cilium continues to coor-dinate Hh signaling in cells derivedfrom the mature pancreas. We alsosuggest that aberrant Hh signaling inthese cancer cell lines may be associ-ated with the autonomous activationof the signaling pathway in the ciliumas judged by high levels of Smo andlow levels of Ptc in the cilium, which isaccompanied by the formation ofGli2(FL) forms in the cilium and lowlevels of Gli3(R) in the nucleus.

In support of the hypothesis thatthe primary cilium may comprise thesite for aberrant Hh signaling in hu-man pancreatic cancer tumorigenesis,we show that in contrast to normalcells, which require Hh pathway stim-ulation in the cultures for Smo to en-

ter the cilium and activate Hh targetgenes (Rohatgi et al., 2007; Kiprilov etal., 2008), Smo is strongly localized tothe primary cilium in both CFPAC-1and PANC-1 cells, even in the absenceof external stimulation of the Hhpathway by SAG. Further and in con-trast to primary cultures of mouse em-bryonic fibroblasts, Ptc is expressed ata higher level than that of MEFs,which require the primary cilium andstimulation of the Hh pathway to in-crease their expression of Ptc. The ac-tivator form of Gli2, Gli2(FL), local-izes to the tip of primary cilium, whichmay comprise the site for activation ofGli transcription factors in Hh signal-ing. We then examined the expressionand localization of Gli3 before and af-ter SAG stimulation by comparing thelocalization of endogenous Gli3 by IFassessed using Gli3-H-280, which rec-ognizes the processed form of Gli3with localization of GFP-Gli3(FL) andGFP-Gli3(R) expressed in transfectgrowth-arrested NIH3T3 fibroblasts.As shown in Figure 5J, only GFP-Gli3(R) localized to the nucleus and tocytoplasmic puncta. These data areconsistent with the conclusion thatthe repressor form is absent from theprimary cilium and is repressing Hhtarget genes in the absence of Hh sig-naling. We then performed IF analysisin NIH3T3 cells in the absence and inthe presence of SAG and compared thestaining with Gli3-H-280 to localiza-tion of the protein expressed from theGFP constructs. As shown in Figure5I (left panels) anti-Gli3 stains non-stimulated cells similar to that of theGFP-Gli3(R) construct. Upon activa-tion of the Hh pathway nuclear Gli3staining is strongly reduced, support-ing the conclusion that the antibody inIF analysis is detecting the repressorform of Gli3. Finally, we examined thelocalization of Gli3 in CFPAC-1 cells,which showed very weak cytosolic andnuclear staining of Gli3 in both stim-ulated and nonstimulated cells. Thisresult would be expected if Hh signal-ing activity is elevated in these cellsbecause Hh signaling represses Gli3expression (Marigo et al., 1996; Bus-cher and Ruther, 1998; Schweitzer etal., 2000), and is further evidenced bythe accumulation of Smo in the ciliumof these cells.

CONCLUSIONS

Our data indicate a functional role ofthe primary cilium in coordinating Hhsignaling during development of thehuman pancreas and potentially in tu-morigenesis of the adult human pan-creas. During development the pri-mary cilium may sense and relay thegraded response to activation of theHh signaling pathway that controlshuman pancreatic organogenesis atthe onset of fetal development. In theadult, the autonomous activation ofthe Hh pathway in pancreatic tumor-igenesis may be linked to aberrant Hhsignaling in the primary cilium of thepancreatic cells. Further analysis willbe required to understand the poten-tial significance of Gli processing inthe cilium and how the switch be-tween activator and repressor forms ofGli2 and Gli3 is regulated by the ci-lium during development and tumori-genesis of the pancreas and poten-tially of other human tissues andorgans. Furthermore, it will be impor-tant to identify the molecular mecha-nisms that coordinate translocation ofSmo and Ptc in and out of the cilium,which ultimately may impinge on theactivation and deactivation of the Hhpathway, which is of relevance in hu-man health and development.

EXPERIMENTALPROCEDURES

Cell Cultures

The human pancreatic exocrine ductcell lines PANC-1 (ATCC, #CRL-1469)and CFPAC-1 (ATCC, #CRL-1918;passages 8–36) were from the Ameri-can Type Culture Collection and werecultured in Dulbecco�s modified Ea-gles medium with glutamax (DMEM;Invitrogen) and Iscove’s modified Dul-becco’s medium (IMDM; Invitrogen),respectively. Both growth media weresupplemented with 10% heat inacti-vated fetal calf serum (FCS; Invitro-gen) and 1% penicillin-streptomycin(Penicilin G sodium: 10,000 U/ml andstreptomycin G sodium: 10,000 �g/ml)(P/S; Invitrogen) and kept in a humid-ified air chamber at 37°C and 5% CO2.Passing of cells was performed bytrypsination. NIH3T3 Swiss mouseembryonic fibroblasts and wt, Ptc�/�and Tg737orpk mouse embryonic fibro-blasts (MEFs) were cultured as de-

2048 NIELSEN ET AL.

scribed previously for MEFs (Schnei-der et al., 2005).

Tissues

Human embryos and fetuses were ob-tained from extra uterine pregnan-cies, Caesarean sections, or prosta-glandin induced legal abortionsdonated to the Developmental BiologyUnit, ICMM, at the Panum Institute,University of Copenhagen, Denmark.The embryos and fetuses ranged from8 to 180 mm crown-rump length(CRL), corresponding to 6th–19th ovu-lation weeks. Informed consent wasobtained according to the guidelines ofthe Helsinki Declaration II. Addi-tional samples from legal first trimes-ter abortions from the Laboratory ofReproductive Biology, Rigshospitalet,and Frederiksberg Hospital (bothUniversity of Copenhagen) were alsoincluded in this study. Informed con-sent was obtained according to theHelsinki declaration II and approvedby the ethical committee of Copenha-gen and Frederiksberg Communities(KF 258206). Calculation of embry-onic and fetal age was based on infor-mation about the last menstrual pe-riod, and measurements of CRL andfoot lengths.

Antibodies, BlockingPeptides, and StainingReagents

Primary antibodies (dilutions in pa-renthesis for IF analysis): mousemonoclonal antibodies from Sig-maAldrich: anti-acetylated �-tubulin(acet. tb, cat. no. T6793; 1:5,000), anti-�-actin (cat. no. A5441; 1:5,000), and�-tubulin (cat. no. T5168; 1:2,000).Rabbit polyclonal anti-detyrosinated�-tubulin (glu-tub; ab48389; 1:500)was from Abcam. Polyclonal antibod-ies from Santa Cruz Biotechnology:goat anti-centrin (cat. no. sc-8719;1:500), goat anti-pericentrin (Pctn;cat.no. sc-28145; 1:500), rabbit anti-Gli2 transcription factor [Gli2(H-300),cat. no. sc-28674; Gli2(N-20), cat. no.sc-20290; Gli2(G-20), cat no. sc-20291;all diluted 1:200], rabbit anti-Gli3transcription factor [Gli3(H-280), cat.no. sc-20688; Gli3(N-19), cat. no. sc-6155; all diluted at 1:100], goat anti-Patched (Ptc; cat. no. sc-6149; 1:200).Two different rabbit anti-Smoothened

were used: (1) Smo (cat. no. LS-A2668;1:100) from MBL, and (2) Smo (cat. no.38686) from Abcam. Secondary anti-bodies for immunofluorescence mi-croscopy analysis (all from MolecularProbes and diluted at 1:600): AlexaFluor 568-conjugated goat anti-mouseIgG (cat. no. A11019), Alexa Fluor568-conjugated rabbit anti-mouse IgG(cat. no. A11061), Alexa Fluor 488-conjugated goat anti-rabbit IgG (cat.no. A11008), and Alexa Fluor 488-con-jugated donkey anti-goat IgG (cat. no.A11055). Secondary antibodies forWestern blot analysis (all from Jack-son ImmunoResearch and diluted1:5,000): alkaline phosphatase conju-gated goat anti-rabbit, rabbit anti-goat,and goat anti-mouse. Nuclei werestained with 4�,6-diamidine-2-pheny-lidole-dihydrochloride (DAPI, 1:600)from Molecular Probes, and F-actin wasstained with rhodamine phalloidin (1:100; Molecular Probes, Invitrogen).Blocking peptides sc-6155P, sc20290P,sc-20291P, and sc- sc-6149P were fromSanta Cruz Biotechnology.

Immunohistochemistry ofthe Developing HumanPancreas

Tissue specimens were dissected intoappropriate tissue blocks and fixed for12–24 hr at 4°C in one of the followingfixatives: 10% buffered formalin, 4%Formol-Calcium, Lillie’s AAF, orBouin’s fixatives. The specimens weredehydrated with graded alcohols,cleared in xylene, and embedded inparaffin wax (Merck, melting point52°C). Serial sections, 3–5 �m thick,were cut in transverse, sagittal, orhorizontal planes and placed on si-lanized slides. Representative sec-tions of each series were stained withhematoxylin and eosin, or with tolu-idine blue. For immunohistochemistry(IHC) sections were dewaxed, rehy-drated and washed in phosphate buff-ered saline (PBS; 137 mM NaCl, 2.6mM KCl, 6.5 mM Na2HPO4, and 1.5mM KH2PO4) as previously described(Teilmann and Christensen, 2005),followed by rinsing with blockingbuffer (5% bovine serum albumin inPBS) for 15 min before incubationwith primary antibodies for at least1.5 hr at room temperature or over-night at 4°C. The sections were thenwashed three times in blocking buffer,

incubated for 45 min in dark with flu-orochrome-conjugated secondary anti-bodies and DAPI in blocking buffer.After washing, sections were mountedin PBS with 70% glycerol and 2% N-propylgallate and sealed with nail pol-ish. Samples were observed on anEclipse E600 microscopes (Nikon, To-kyo, Japan) with EPI-FL3 filters andMagnaFire cooled CCD camera(Optronics, Goleta, CA), and digitalimages were processed using AdobePhotoshop 6.0.

Immunocytochemistry

Cells were grown on 25-mm-diametersterile polylysine-coated glass cover-slips for 3–5 days to �80% confluency,followed by serum starvation inDMEM/IMDM with 0.5% (PANC-1and CFPAC-1) and 0% (MEF) FCSand 1% P/S for 24, 48, and 72 hr, toinduce growth arrest and formation ofprimary cilia. The cells were brieflywashed in PBS, fixed for 15 min in 4%paraformaldehyde followed by washin blocking buffer and then permeabil-ized with 1% Triton X-100 in PBS for10 min. The coverslips were rinsedwith blocking buffer for 30 min beforeincubation with antibodies as de-scribed above for IHC, and mountedon microscope slides in anti-fademounting solution and sealed withnail polish. Samples were observed onan Eclipse E600 microscopes (Nikon,Tokyo, Japan) with EPI-FL3 filtersand MagnaFire cooled CCD camera(Optronics, Goleta, CA), and digitalimages were processed using AdobePhotoshop 6.0.

Sodium Dodecyl Sulfate-Polyacrylamide GelElectrophoresis and WesternBlot Analysis

Cell lysates were prepared by using1% sodium dodecyl sulfate (SDS) lysisbuffer. Lysates were sonicated andcentrifuged to separate off cell debris.Protein concentrations were mea-sured using a DC Protein Assay fromBio-Rad. Proteins were resolved bySDS-polyacrylamide gel electrophore-sis (SDS-PAGE) as described byChristensen et al. (2001). Briefly, pro-teins were separated under reducingconditions with NuPAGE 10% Bis-Tris precast gels (Invitrogen), fol-


lowed by electrophoretic transfer tonitrocellulose membranes (Invitro-gen). Before incubation with antibod-ies for 2 hr/over night the membraneswere blocked with 5% milk/TBST (10mM Tris/HCl [pH 7.5], 120 mM NaCl,0.1% Tween 20) and 0.5% Na-azide.Antibodies were diluted in 5% milk/TBST as indicated: anti-Ptc (1:200),anti-Gli2 (N-20) and (G-20) (1:200),anti-Gli3 (H-280) and (N-19) (1:100),anti-�-actin (1:5,000), and anti-�-tu-bulin (1:2,000). Membranes werewashed several times in TBST fol-lowed by incubation with alkalinephosphatase-conjugated secondaryantibodies. Blots were developed withBCIP/TNBT from KPL. The developedblots were scanned and band intensitywas estimated from arbitrary densito-metric values obtained using UN-SCAN-IT software. The blots werefurther processed for publication inAdobe Photoshop version 6.0.

SAG Stimulation

Cultures of PANC-1, CF-PAC1 cellsand NIH3T3 fibroblasts (80% conflu-ency) were serum starved for 48–72hr and incubated in the presence andin the absence of 1 �M SAG (AlexisBiochemicals, San Diego, CA) for 0and 24 hr, followed by IF analysiswith anti-Smo and anti-Gli3 and byWB analysis with anti-Ptc and anti-Gli2. Primary cilia were visualizedwith anti-acet. tb and nuclei withDAPI. All images were taken withequivalent time exposures.

Transfection of Cells WithFluorescent-Tagged Plasmids

CFPAC-1 cells were cultured on cov-erslips placed in six-well test plates(NUNC A/S, Denmark) and grown to70% confluency. The medium waschanged to DMEM/IMDM for 1 hr, fol-lowed by incubation with 1 �g/ml plas-mid (YFP-Smo, provided by P. Beachy)and 5 �l of lipofectamine 2000 (Invitro-gen) per well. After 4 hr of transfectionthe medium was changed to DMEM/IMDM with 10% FCS, and after thecells had reached 90% confluency theywere serum-starved for 48 hr to inducegrowth arrest. Cells were then fixed andpermeabilized and subjected to immu-nocytochemistry analysis as describedabove. NIH3T3 cells were cultured

and transfected under the same con-ditions as above, except that serum-free DMEM was used during the 4-hrtransfection period. The following plas-mids were used: full length Gli2::GFP,GFP-Gli2(FL), full length Gli3::GFP,GFP-Gli3(FL), Gli3repressor::GFP, GFP-Gli(R) (Buttitta et al., 2003). All the vec-tors were constructed in the pShuttlebackbone (Clontech).

ACKNOWLEDGMENTSWe thank Drs. Melissa Lutterodt andAnne Grete Byskov (Laboratory of Re-productive Biology, Rigshospitalet,Denmark) for expert help with collect-ing of human embryonic tissues. Wealso thank Dr. Kathryn Anderson(Memorial Sloan-Kettering CancerCenter, NY, USA) for the gift of thePtc�/� MEFs, and Simon Krabbeand Mette R. Hansen (Department ofBiology, University of Copenhagen,Denmark) for help with cultures of hu-man pancreatic ductal adenocarci-noma cell lines. This work was sup-ported by the Lundbeck Foundationand the Danish Science ResearchCouncil (S.T.C.), the Novo Nordisk/Novozymes Foundation (S.K.N.), theDanish Science Research Council(I.N.), funds from the Department ofBiology, University of Copenhagen(C.A.C.), and by a National Instituteof Health Grant (B.K.Y.).

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© The Rockefeller University Press $30.00The Journal of Cell Biology, Vol. 180, No. 5, March 10, 2008 897–904http://www.jcb.org/cgi/doi/

JCB 89710.1083/jcb.200706028

JCB: REPORT

E.N. Kiprilov and A. Awan contributed equally to this paper.

Correspondence to Rhoda Elison Hirsch: [email protected]

Abbreviations used in this paper: AcTb, acetylated tubulin; hESC, human embry-onic stem cells; hFF, human foreskin fi broblast; Hh, hedgehog; IF, immunofl uores-cence; Ptc1, patched 1; SAG, Smo agonist; SEM, scanning electron microscopy; SHh, sonic hedgehog; Smo, smoothened; TEM, transmission electron micros-copy; Tra-1-85, tumor rejection antigen 1-85.

The online version of this paper contains supplemental material.

Introduction Driving human embryonic stem cells (hESCs) along specifi c

differentiation pathways remains a signifi cant challenge for

translational medicine and the development of hESC therapies.

During early embryology, signaling pathways, such as hedgehog

(Hh) and Wnt, are critical for human development ( Corbit et al.,

2005 ; Haycraft et al., 2005 ; Huangfu and Anderson, 2005 ; Liu

et al., 2005 ; May et al., 2005 ) and, recently, have been shown to

be mediated by the primary cilium (for reviews see Michaud

and Yoder, 2006 ; Singla and Reiter, 2006 ; Christensen et al.,

2007 ; Satir and Christensen, 2007 ). Therefore, in the search for

mechanisms regulating hESC differentiation, it is vital to fi rst

establish the existence of primary cilia and the localization of

signaling components in undifferentiated hESCs.

Primary cilia are single, generally nonmotile, cilia with a

9 + 0 axoneme, differing from the 9 + 2 arrangement of motile

cilia. Primary cilia are implicated as key cellular sensory structures

involved in signal transduction and coordination of intra- and

intercellular signaling pathways (for reviews see Michaud and

Yoder, 2006 ; Singla and Reiter, 2006 ; Christensen et al., 2007 ;

Satir and Christensen, 2007 ). Signaling in primary cilia is thought

to be initiated by receptors positioned within the cilium and re-

layed through transcription factors, which may become activated

directly in the cilium or in the cell body via basal body scaffold

proteins. Specifi c growth factor receptors in the primary cilium,

such as PDGF receptor- � , enable the cell to respond differentially

to ligands and to initiate cell division ( Schneider et al., 2005 ).

Mutations giving rise to defective primary cilia or improper

placement of signaling molecules within the cilium result in a

plethora of clinical manifestations ( Pazour, 2004 ; Badano et al.,

2006 ). These include obesity, rod – cone dystrophy, renal ab-

normalities, polycystic kidney disease, polydactyly, genital ab-

normalities, learning disabilities, congenital heart disease, hearing

loss, situs inversus, and Bardet-Biedl syndrome ( Blacque and

Leroux, 2006 ). In particular, mutations in genes encoding intra-

fl agellar transport proteins impair Hh signaling and result in

limb bud and neural tube defects, which are similar to those seen in

Hh signaling mutations ( Corbit et al., 2005 ; Haycraft et al., 2005 ;

Huangfu and Anderson, 2005 ; Liu et al., 2005 ; May et al., 2005 ).

Hh signaling is essential during embryonic development for

Human embryonic stem cells (hESCs) are potential

therapeutic tools and models of human develop-

ment. With a growing interest in primary cilia in

signal transduction pathways that are crucial for embryo-

logical development and tissue differentiation and interest

in mechanisms regulating human hESC differentiation,

demonstrating the existence of primary cilia and the local-

ization of signaling components in undifferentiated hESCs

establishes a mechanistic basis for the regulation of hESC

differentiation. Using electron microscopy (EM), immuno-

fl uorescence, and confocal microscopies, we show that

primary cilia are present in three undifferentiated hESC

lines. EM reveals the characteristic 9 + 0 axoneme. The num-

ber and length of cilia increase after serum starvation.

Important components of the hedgehog (Hh) pathway,

including smoothened, patched 1 (Ptc1), and Gli1 and 2,

are present in the cilia. Stimulation of the pathway results

in the concerted movement of Ptc1 out of, and smoothened

into, the primary cilium as well as up-regulation of GLI1

and PTC1 . These fi ndings show that hESCs contain pri-

mary cilia associated with working Hh machinery.

Human embryonic stem cells in culture possess primary cilia with hedgehog signaling machinery

Enko N. Kiprilov , 1 Aashir Awan , 1,4,5 Romain Desprat , 3 Michelle Velho , 2 Christian A. Clement , 4 Anne Grete Byskov , 5

Claus Y. Andersen , 5 Peter Satir , 1 Eric E. Bouhassira , 2,3 S ø ren T. Christensen , 4 and Rhoda Elison Hirsch 1,2

1 Department of Anatomy and Structural Biology, 2 Department of Medicine, and 3 Department of Cell Biology, Albert Einstein College of Medicine, Bronx, NY 10461 4 Institute of Molecular Biology, University of Copenhagen, DK-2100 Copenhagen OE, Denmark 5 Laboratory of Reproductive Biology, Rigshospitalet, DK-2100 Copenhagen OE, Denmark

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JCB • VOLUME 180 • NUMBER 5 • 2008 898

hESCs. The presence of this organelle is not limited to specifi c

culture conditions. HESCs from H1 (male) and H9 (female) lines

(approved by the National Institutes of Health; Olivier et al.,

2006 ) were grown on matrigel without feeder cells (described in

Yao et al. [2006] ) with serum replacement for 6 d. Primary cilia

were fi rst identifi ed by immunofl uorescence (IF) markers of

acetylated tubulin (AcTb) in both H1 and 9 hESCs after 6 d of

culture in DME:F12 with serum replacement ( Figs. 1 A and 2 A ).

Primary cilia became more prominent after starvation of hESCs

by placement in DME:F12 without serum replacement for 24 h

( Fig. 1 B ). Another hESC line, LRB003 (female; studied in the

Denmark laboratory and supported by funds independent of the

National Institutes of Health; Laursen et al., 2007 ), was cultured

in monolayers on 0.1% gelatin with conditioned medium from

cultured human foreskin fi broblasts (hFF), and primary cilia

were observed after 4 d as the cells entered growth arrest in con-

fl uent colonies in the culture dish ( Fig. 1, D and E ).

Confi rmation that the hESCs remained undifferentiated

was made by IF using the transcription factor OCT-4 ( Fig. 1,

A, D, and E ) and stage-specifi c embryonic antigen 4 (not depicted).

Both markers were used to assure undifferentiated hESCs. Anti-

AcTb identifi ed potential primary cilia ( Fig. 1, B and C ) and

antibodies against tumor rejection antigen 1-85 (Tra-1-85) –

marked human cells (see Fig. 3 D ). After 5 d in culture, short

( � 2 – 3 μ m) AcTb extensions characteristic of primary cilia were

seen on � 33% of H1 hESCs (25 cilia/75 cells counted from fi ve

left – right asymmetry axis, limb and heart development, and

neurogenesis ( Corbit et al., 2005 ; Haycraft et al., 2005 ; Huangfu

and Anderson, 2005 ; Liu et al., 2005 ; May et al., 2005 ). In the

adult, Hh signaling is involved in stem cell maintenance and

tissue homeostasis.

We hypothesized that primary cilia might be found in

hESCs, wherein they could play a critical role in hESC differ-

entiation parallel to that in normal early embryogenesis. In this

study, we demonstrate that primary cilia are a general feature of

hESC lines and that essential signaling components of the Hh

pathway are present and functional in primary cilia of undiffer-

entiated hESCs. Transmission electron microscopy (TEM) and

scanning electron microscopy (SEM) images provide defi nitive

evidence and reveal novel features of hESCs and their primary

cilia. To date, this is the fi rst study conclusively showing the pres-

ence of these unique organelles in hESCs by defi nitive confocal

and electron micrographs of hESC primary cilia and by dynamic

colocalization of key signaling molecules essential for early

development and known to be functional in the Hh signaling

pathway, as was recently demonstrated in primary cilia of cul-

tured mouse fi broblasts ( Rohatgi et al., 2007 ).

Results and discussion In this study, we demonstrate that the primary cilium is a dynamic

ultrastructural feature in three different lines of undifferentiated

Figure 1. Immunolabeling of primary cilia in undifferentiated hESCs. (A) Characterization of undifferentiated colonies of H1 hESCs grown on matrigel for 5 d in DME:F12 with serum replacement. Undifferentiated cells are identifi ed by nuclear colocalization of anti – OCT-4 (OCT-4, green) and DAPI (dark blue) in the merged image (light blue). More than 97% of cells on average expressed OCT-4 in a nuclear pattern indicating their undifferentiated state. Primary cilia stained with anti-AcTb (tb, red) are indicated by arrows. (B) H1 hESCs grown on matrigel for 5 d in DME:F12 with serum replacement (i.e., unstarved), labeled with anti-AcTb (tb, green) to show primary cilia (arrows). (C) H1 hESCs grown on matrigel for the same period of time in DME:F12 without serum replacement (i.e., starved) for 24 h and labeled with anti-AcTb (tb, green). Primary cilia are indicated by arrows. (D) Characterization of undifferentiated colonies of LRB003 hESC grown on 0.1% gelatin with conditioned medium. Undifferentiated cells are identifi ed by nuclear colocalization of anti – OCT-4 (OCT-4, green) and DAPI (dark blue) in the merged image (light blue). Anti-AcTb (tb, red) marks the primary cilia (arrows) as well as the microtubular network in the cytoplasm. Less than or equal to 95% of the cells were ciliated and positive for OCT-4. Primary cilia are not easily visualized at the low resolution images (as shown in the insets). (E) A primary cilium (tb, red, arrow) in undifferentiated LRB003 hESCs emerges from one of the centrioles (asterisks) marked with anti-centrin (centrin, green). Nuclear localization of anti – OCT-4 (OCT-4, blue) denotes that the cell has not differentiated. The inset shows anti-pericentrin (Pctn, green) marking the centrosome ( ¤ ) at the base of the primary cilium (arrow). on N

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899HUMAN EMBRYONIC STEM CELLS AND PRIMARY CILIA • KIPRILOV ET AL.

surfaces of the cells, which is in contrast to the many microvilli

that are shorter and have a smaller diameter ( Fig. 2 B ). SEM also

demonstrated paddle tips at the ends of some primary cilia ( Fig. 2,

C and D ). Confocal images ( Fig. 2 A ) show the outward orienta-

tion of primary cilium from growth-arrested cells in a monolayer,

whereas mitotic cells lack a primary cilium ( Pan and Snell, 2007 ).

To show defi nitively that the structures are primary cilia,

we fi xed hESC cultures in situ and processed them for TEM.

Some colonies were cut parallel to and just above their free sur-

faces to give cross-sectional views of projecting structures, and

other sections were oriented through the cell bodies perpendicular

to this direction to show longitudinal views of the cilia and

their basal bodies. Cross sections near the apical surfaces of

the cells showed axonemes, which are enclosed by a unit mem-

brane ( Fig. 2 E ). The 9 + 0 pattern can be clearly observed in

cross sections close to the basal body, as are a disarray of nine

doublets, including 8 + 1, 6 + 1, and other patterns ( Fig. 2 E ),

either from the same cell but at different sections along the length

of the cilia approaching the tip or in different cells at varying

stages of ciliary growth. One centriole pair can also be observed

close to the cell surface with a primary cilium growing from one

of the centrioles ( Fig. 2 F ), which has become the ciliary basal

body. Primary cilia often emerge from a concavity in the cell,

different fi elds of colonies on a plate; Fig. 1 B ). In the remaining

H1 cells, anti-AcTb was often seen as a single concentrated spot,

which likely represents a very short cilium of length < 1 � m.

When cultures were starved in DME:F12 alone (without serum

replacement) for 24 h, the AcTb extensions increased in length

to � 4 – 6 μ m, and the number of cells with these extensions in-

creased to � 50% (41/80 cells, counted as the H1 hESCs; Fig. 1 C ).

H9 behaved essentially similarly (unpublished data).

In the LRB003 hESCs, after � 7 d in culture and indepen-

dent of starvation, primary cilia with lengths of 5 – 10 μ m emerged

as solitary organelles from > 90% of the confl uent cells ( Fig. 1 D ).

To show unequivocally that cilia were present on undifferenti-

ated LRB003 hESCs, we used triple colocalization with anti-

AcTb, anti – OCT-4, and either anti-centrin or anti-pericentrin

( Fig. 1 E ). Single primary cilia (labeled by anti-AcTb) were shown

to emanate from the centrosomes (labeled by anti-pericentrin;

Fig. 1 E , inset) of OCT-4 – positive cells and, in particular, from one

of the two centrioles (labeled by anti-centrin; Fig. 1 E ), prob-

ably the mother centriole, which also functions as the ciliary

basal body.

SEM of H1 and 9 hESCs (95% OCT-4 positive) revealed

single AcTb-positive projections � 4 – 6 μ m in length and � 0.25 μ m

in diameter (characteristic of primary cilia) seen at the free

Figure 2. Electron microscopy of hESC primary cilia. (A) Confocal microscopy of hESCs grown on matrigel in a monolayer viewed from the side (a) and the top (b), showing primary cilia (arrows) on undifferentiated hESCs but not on dividing cells (arrowhead). (B) SEM of H1 cells grown for 7 d on matrigel in N2/B27-supplemented medium and starved in DME:F12 without serum replacement for the last 24 h of growth. See Online supplemental material for details. Primary cilia (arrows) are seen on two adjacent cells among numerous smaller microvilli. (C and D) Enlarged images of primary cilia. Note paddle tips. (E) TEM cross sections of primary cilia of H1 and 9 hESCs grown for 7 d on matrigel in N2/B27-supplemented medium and starved in DME:F12 without serum replacement for the last 24 h before fi xation. The fi rst panel shows the 9 + 0 arrangement, which becomes disorganized along the ciliary length as shown on the rest of the panels. (F) TEM longitudinal section of a primary cilium (arrowhead) emerging from one of a pair of centrioles/basal bodies. The perpendicular orientation of the basal bodies suggests that the cilium arises from the mother centriole (arrow). A lamellar vesicle (asterisk) is seen budding from the cell surface. (G) TEM of lamellar-type bodies secreted by hESCs.

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Gli transcription factors that enter the nucleus to control differ-

ential processes during early and late embryogenesis. Smo was

previously reported to be a constituent of nodal cilia, Madin Darby

canine kidney cell cilia, and other primary cilia ( Corbit et al.,

2005 ; May et al., 2005 ), and Gli2 was found at the tip of mesen-

chymal primary cilia during limb formation ( Haycraft et al., 2005 ).

Time-dependent studies in mammalian differentiated cells sup-

port a model in which SHh triggers the removal of Ptc from the

primary cilium, permitting Smo to enter the cilium and initiat-

ing signaling ( Rohatgi et al., 2007 ). We therefore tested whether

Smo, Ptc, and Gli2 are present in hESC primary cilia, and we

followed the movement of Smo and Ptc in and out of the cilium

upon stimulation by Smo agonist (SAG). The use of SAG to in-

duce activation of SHh signaling has been established by Chen

et al. (2002) . In transfected LRB003 hESCs, YFP:Smo strongly

and almost exclusively localizes to the primary cilium ( Fig. 3 A ).

The ciliary staining of YFP:Smo was remarkably higher than

that of anti-Smo ( Fig. 4 A ) because of overexpression of Smo

from the construct. Furthermore, with a Gli2-specifi c antibody,

which may be interpreted as a small depression in the cell ’ s apical

surface as shown in the SEM ( Fig. 2 D ) and TEM (Fig. S1 A, avail-

able at http://www.jcb.org/cgi/content/full/jcb.200706028/DC1)

images. Rarely (in < 1% of observed cells), two primary cilia

originate within one cell (unpublished data). A rich array of poly-

somes and cytoplasmic microtubules, running parallel to the api-

cal surface, are seen near the basal body (Fig. S1 A). Immediately

below this level, a ciliary rootlet emerging from the basal body

and microfi lament bands of the adherens junctions of the confl uent

hESCs can be found (Fig. S2). In addition, lamellar-type vesicles

are observed both intracellularly and extracellularly, adherent to

the hESC surface ( Fig. 2, F and G ; and Fig. S1 B).

Next, we examined whether components of the Hh signal-

ing system were present and functional in the hESC primary cilia.

It has been reported previously that the sonic Hh (SHh) recep-

tors patched (Ptc) and smoothened (Smo) and their downstream

effectors Gli1, 2, and 3 are expressed in hESCs ( Rho et al., 2006 ).

In various cells, upon binding of SHh to its receptor Ptc, Smo is

activated, which is followed by the processing and activation of

Figure 3. Hh signaling proteins localize to hESC primary cilia. (A) Localization of YFP:Smo (green) to the primary cilium (tb, red, arrow) in LRB003 hESCs. The merged image shows colocalization. Nuclei are stained with DAPI (blue). (B) Immunolocalization of Gli2 (green) to primary cilia (tb, red, arrows) in LRB003 hESCs. The merged image shows colocalization. Nuclei are stained with DAPI (blue). Gli data from H1 and LRB003 cells were obtained with different antibodies. (C) H1 cells grown for 7 d on matrigel in N2/B27-supplemented medium and labeled for primary cilia (tb, green) and anti-SHh (SHh, red). In addition, a z series of a fi eld showing separate SHh labeling (red) located distinctly to the side of the primary cilium (green) is depicted. (D) Similar cells labeled with anti – Tra-1-85 (red) and anti-Gli2 (Gli2, green). Arrows indicate punctuate localization of the Gli2 protein (green dots) and the inset specifi cally localizes the Gli2 protein (arrowhead) at one end of a primary cilium (tb, red). (E) Anti-SHh (SHh, red) localizes near the base of most primary cilia. 33/71 cells possess primary cilia (46.5%), which is consistent with Fig. 1 C . 22/33 cells with primary cilia (66%) exhibit SHh near their base (arrows). SHh also localizes to points not associated with the cilia (arrowheads). The asterisk points to the midbody of cells that have recently under-gone cytokinesis. These structures are not associated with Hh signaling molecules. (F) Anti-Smo (Smo, red) localizes predominantly to the base of primary cilia. This pattern of Smo expression is similar to that observed at 0 and 1 h of SAG stimulation (see Fig. 4 ). Arrows point to primary cilia (tb, green) and arrowheads indicate Smo localization.

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proteins and their Hh cargoes in hESCs that would establish

mechanisms of traffi cking. Knockdown experiments, for exam-

ple, using siRNA of KIF3A, would be informative and are pres-

ently underway.

The addition of 5 μ M SHh or 10 μ g/ml SAG to H1 hESCs

for 18 h up-regulated GLI1 (approximately twofold) and PTC1

(approximately fi vefold) mRNA levels compared with baseline

levels of these components without exogenous ligand stimulation,

as determined by real-time PCR with GAPDH as an internal

control ( Fig. 5 ). As expected, GLI2 mRNA was essentially non-

responsive. Cyclopamine, a Smo inhibitor ( Lipinski et al., 2006 ),

modestly inhibited the up-regulation in the presence of inducers

under the conditions used ( Fig. 5 ). GLI2 mRNA was not affected.

These data are consistent with the dynamics of the Hh signaling

machinery, as described by Rohatgi et al. (2007) , in differentiated

cells and, together with the localization studies of Hh signaling

proteins, support the conclusion that Hh signaling proceeds

through hESC primary cilia. Whether or not the SHh ligand is

produced by the hESC and whether the function of the signal is

to maintain the cells undifferentiated or act as a precursor to

differentiation remains to be determined.

The presence of the extracellular lamellar bodies in un-

differentiated hESCs may also be related to Hh or other signaling

pathways. Similar vesicles have been reported to be involved in

we show that Gli2 strongly localizes in a punctuate pattern along

the entire length of primary cilia but is absent in the nucleus of

these cells ( Fig. 3 B ). Also, in H1 and 9 hESCs, anti-Gli2 local-

izes to the primary cilia ( Fig. 3, B and D ), whereas Smo local-

izes to the base of � 3/4 of the cells with primary cilia ( Fig. 3 F ).

In addition, by fl uorescence immunolocalization, small amounts

of SHh can be localized near the base of the cilia, which is

clearly located to the side of the primary cilium ( Fig. 3 C , z series)

in � 2/3 of the ciliated H1 cells ( Fig. 3, C and E ). In LRB003

cells, upon stimulation with SAG, the ciliary level of Smo starts

to increase beginning at 1 h ( Fig. 4 B ) as compared with 0 h

( Fig. 4 A ). This is followed by a major accumulation of Smo

along the length of the cilium at 4 h of SAG treatment ( Fig. 4 C ).

This infers that translocation of Smo along the cilium is initi-

ated by the docking of Smo at the base of the cilium. The oppo-

site pattern of translocation can be seen for Ptc, which leaves

the cilium upon SAG stimulation ( Fig. 4, D – F ). This movement

of Hh components into and out of the cilium ( Fig. 4 ), along with

the z series showing SHh located to the side of the primary cilium

( Fig. 3 C ), eliminates the possibility of nonspecifi c antibody

binding to the centrosome in light of the fact that centrosomes

never migrate up the cilium. Experiments similar to that described

by Orozco et al. (1999) are planned for the direct viewing of intra-

cellular and ciliary transport of intrafl agellar transport motor

Figure 4. Translocation of Smo and Ptc in and out of the primary cilium after SAG stimulation. (A) Localization of anti-Smo (green) to the pri-mary cilium of LRB003 cells (tb, red, arrows) at 0 h of SAG treatment. (B and C) Localiza-tion of anti-Smo to the primary cilium (arrows) at 1 and 4 h, respectively. The insets in A – C show high resolution (shifted overlays) of Smo (green) in the primary cilium (red). The asterisk indicates the base of the cilium. (D) Localization of anti-Ptc (green) to the primary cilium (tb, red, arrows) at 0 h of SAG treatment. (E and F) Local-ization of anti-Ptc at 1 and 4 h, respectively, to primary cilium (arrows). The insets in D – F show high resolution images (shifted overlays) of Ptc (green) in the primary cilium (red). Nuclei were stained with DAPI.

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L- glutamine and 15 mM Hepes, supplemented with the serum replacements N2 (chemically defi ned supplement containing 1000 mg/liter human trans-ferrin, 50 mg/liter insulin recombinant full chain, 0.6 mg/liter progesterone, 161 mg/liter putrescine, and 173 mg/liter selenite; Invitrogen) as 100 × con-centrate of Bottenstein ’ s N2 formulation ( Bottenstein, 1985 ) and B27 (50 × serum supplement designed for the long-term viability of hippocampal and other neurons of the central nervous system; Invitrogen), in addition to 20 ng/ml of basic FGF (R & D Systems), BSA fraction V, 1% nonessential amino acids, 50 U/ml penicillin, 50 ng/ml streptomycin, 1 mM L- glutamine, and 1-thioglycerol added for 6 d (as described in detail in Yao et al. [2006] ) and observed by phase microscopy using an inverted light microscope (CK40; Olympus). To passage the cells, differentiated cells were scraped in PBS under a binocular magnifi er with a Pasteur pipette scraper (elongated and twisted using heat), treated with prewarmed collagenase type IV for 5 min to detach the hESC colonies, aspirated, concentrated using a macrocentrifuge (Eppendorf), and either plated on 6-well tissue culture plates (Thermo Fisher Scientifi c) coated with 1:4 matrigel for propagation, on gamma-irradiated 35-mm glass-bottom microwell dishes (MatTek Cultureware) covered with 1:4 matrigel for IF, or on carbon-coated glass coverslips on the bottom of each well of a 6-well plate covered with 1:4 matrigel for TEM or SEM. The cultures were monitored microscopically and at day 6 were either maintained for an additional day in the same supplemented medium or starved in plain DME:F12 for 24 h. The cells were then prepared for IF microscopy using the proto-col described in IF Microscopy (Albert Einstein College of Medicine).

The LRB003 cell line (not approved by the National Institutes of Health) was studied, as described in the next section, in the Copenhagen laboratory and solely supported by Danish funding agencies (see Acknowledgments).

Cell cultures (Copenhagen) The hESC line LRB003 ( Laursen et al., 2007 ) was initially cultured on 35-mm dishes (Thermo Fisher Scientifi c) coated with 0.1% gelatin (Sigma-Aldrich) on a confl uent layer of mitotically inactivated hFF (Line #CCD-1112Sk; American Type Culture Collection). The hESC culture medium consisted of the following: knockout DME, 15% knockout serum replacement, 2 mM GlutaMAX, nonessential amino acids, 50 U/ml penicillin, 50 ng/ml strepto-mycin, and 0.1 mM � -mercaptoethanol (Invitrogen); and 4 ng/ml basic FGF (R & D Systems). Cells were maintained in a humidifi ed incubator at 37 ° C with an atmosphere consisting of 6% CO 2 , 7% O 2 , and 87% N 2 . After 5 – 7 d of incubation, hESCs were passaged using trypsin (Invitrogen) for ex-perimental culturing conditions in a feeder-free environment. The cells were plated on 16-well glass slides (Thermo Fisher Scientifi c) coated with 0.1% gelatin (BD Biosciences) in the absence of hFF. The conditioned media used consisted of hFF supernatant and hESC culture media (1:1).

IF microscopy (Albert Einstein College of Medicine) hESCs from H1 and 9 cell lines were washed in Dulbecco ’ s PBS without cal-cium and magnesium (Mediatech, Inc.) at RT, and then fi xed in 3.7% para-formaldehyde in PBS for 15 min. They were then rinsed three times with PBS, incubated in 0.1% Triton X-100 (Sigma-Aldrich) in PBS for 10 min, and blocked with 2% BSA in PBS for 1 h at RT or overnight at 4 ° C, and primary antibodies (monoclonal anti-AcTb mouse anti – human IgG2b [Sigma-Aldrich];

signaling in association with cells with nodal or primary cilia in

several embryonic tissues. It would be interesting if the lamellar

vesicles seen here are indeed akin to nodal vesicular parcels con-

taining SHh signals, as described in early embryonic nodal cells

( Tanaka et al., 2005 ; Hirokawa et al., 2006 ), or to prominin-1 –

containing particles of dividing neuroepithelial cells of the

developing mammalian central nervous system (Dubreuil et al.,

2007). The content of the H1 lamellar vesicles remains to be in-

vestigated further.

In summary, defi nitive confocal and transmission electron

micrographs, coupled with SEM and IF microscopy, conclu-

sively demonstrate the presence of primary cilia with many

known features in hESCs. For a detailed review of primary cilia

ultrastructure in differentiated cells, see Satir and Christensen

(2007 ). Because Hh signaling pathways of embryological

development and patterning operate via primary cilia, it is per-

haps not surprising to fi nd Ptc, Smo, and Gli2 localized and po-

tentially functional within the hESC primary cilia. Whether SHh

is released in cultures before or after differentiation of hESCs is

unclear, and whether important receptors of other signaling path-

ways, such as Wnt ( Gerdes et al., 2007 ; Pan and Thomson, 2007 ),

are localized in hESC primary cilia remains to be determined.

Collectively, our results suggest that primary cilia may be

involved in the regulation and coordination of the fi rst steps of

hESC differentiation and/or the maintenance of the undifferenti-

ated state/self-renewal. Because hESCs hold promise for the treat-

ment of many diseases and provide an excellent system for

studying mechanisms involved in early human development, these

fi ndings provide the groundwork to determine specifi c aspects of

early differentiation controlled by the machinery of primary cilia.

This knowledge may ultimately reveal pathways for manipulation

of hESC differentiation into specifi c cell and tissue lineages.

Materials and methods Cell cultures (Albert Einstein College of Medicine) hESCs from H1 and 9 lines (National Institutes of Health approved) were maintained in a humidifi ed incubator at 37 ° C with an atmosphere consisting of 6% CO 2 , 7% O 2 , and 87% N 2 and were grown on matrigel (BD Bio-sciences) without feeder cells in DME nutrient mixture F12 (Ham; Invitrogen) with

Figure 5. Up-regulation of mRNAs for Hh com-ponents in H1 HESCs by SHhN or SAG induction as determined by real-time PCR (Materials and methods). A comparison of the relative increase in mRNA compared with baseline levels (without stimulation) for the respective components is shown. This experiment was reproduced twice with essentially the same qualitative results, and the mean values are shown as numbers above each bar with the corresponding stan-dard error bars, which represent the standard error of the mean. Each experiment used three pooled samples to obtain suffi cient RNA for the technique.

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RNA purifi cation from H1 hESC lines and real-time quantitative PCR (Albert Einstein College of Medicine) After 18 h, RNA was extracted from three pooled wells of H1 hESCs after stimulation, pretreated with DNase, and further purifi ed by RNeasy columns (QIAGEN). cDNA synthesis was performed using the SuperScript III double-stranded cDNA synthesis kit (Invitrogen) on a Mastercycler Gradient (Eppendorf). Single-stranded cDNA was cleaned on a QIAquick PCR purifi ca-tion kit (QIAGEN) and 50 ng was used for the PCR quantifi cation. Gene ex-pression was assayed by quantitative real-time RT-PCR using TaqMan gene expression master mix (Applied Biosystems) and TaqMan gene expression assay primer and probe sets (Applied Biosystems) of PTCH1 (Assay ID, Hs00181117_m1), GLI1 (Hs00171790_m1), and GLI2 (Hs00257977_m1) on the iCycler (Applied Biosystems) and normalized using the internal control gene human GAPDH (FAM/MGB Probe, non – primer limited; Ap-plied Biosystems), which was used as the endogenous reference in the H1 hESC line assays. Each sample was run twice in triplicate. PCR reactions were run for 40 cycles. The log-linear phase of amplifi cation was monitored to obtain threshold cycle values. The comparative threshold cycle method was used to determine levels of expression. Absence of primer dimers was verifi ed by running the PCR product on a 1.5% agarose gel.

Transfection of cells (Copenhagen) 800 ng of pSmo:YFP (provided by P. Beachy, Stanford University School of Medicine, Stanford, CA) was mixed with FuGene6 (Roche) at 6:1 at RT for 45 min. Afterward, 16 μ l of the mix was aliquoted to each well containing hESC colonies in 100 μ l knockout DME. After 3 h at 37 ° C, the medium was replaced with conditioned medium and incubated at 37 ° C for an ad-ditional 48 h. The cells were fi xed and permeabilized, and anti-AcTb was added at 1:10,000 and visualized with Alexa Fluor 568 – conjugated rabbit anti – mouse IgG along with DAPI staining.

TEM (Albert Einstein College of Medicine) The carbon-coated matrigel-covered samples with hESC colonies of H1 and 9 cells, respectively, were fi xed with 2.5% glutaraldehyde and 0.5% tannic acid in 0.1 M sodium cacodylate buffer, postfi xed with 1% osmium tetroxide followed by 2% uranyl acetate, dehydrated through a graded se-ries of ethanol, and embedded in LX112 resin (Ladd Research Industries). Ultrathin sections were cut on a Ultracut UCT (Reichert), stained with uranyl acetate followed by lead citrate, and viewed on a transmission electron microscope (1200EX; JEOL) at 80 kV.

SEM (Albert Einstein College of Medicine) The carbon-coated matrigel-covered samples with hESC colonies of H1 and 9 cells, respectively, were fi xed in 2.5% glutaraldehyde, 0.1 M sodium cacodylate, 0.2 M sucrose, and 5 mM MgCl 2 , pH 7.4, dehydrated through a graded series of ethanol, critical point dried using liquid CO 2 in a critical point drier (Samdri 795; Tousimis), sputter coated with gold-palladium in a sputter coater (Vacuum Desk-2; Denton), and examined in a scanning electron microscope (JSM6400; JEOL) using an accelerating voltage of 10 kV.

Online supplemental material Fig. S1 shows a transmission electron micrograph of an hESC showing the details of lamellar vesicles with a ciliary necklace around a forming pri-mary cilia. Fig. S2 shows a transmission electron micrograph of a section taken just below the cell cortex with a centriole/basal body showing a cili-ary rootlet and microfi lament bands at the adherens junctions of confl uent H9 hESCs. Online supplementary material is available at http://www.jcb.org/cgi/content/full/jcb.200706028/DC1.

We thank Aaron Bell for his expert assistance with preliminary IF and EM data collection and Frank Macaluso, Michael Cammer, Juan Jimenez, and Leslie Gunther of the Albert Einstein College of Medicine Analytical Imaging Facility for their expert technical assistance. The Copenhagen team thanks Roberto Oliveri for his expert assistance with culturing LRB003 hESCs.

This work was supported in part by grants from the National Institutes of Health (NIDDK DK 41296, DK41918, and NIAAA AA008769 to P. Satir and DK064123 to R.E. Hirsch) and National Institute of General Medical Sciences (GM075037 to E.E. Bouhassira). It must be emphasized that National Institutes of Health funds were only used for experiments with the H1 and 9 hESC cell lines that are listed on the approved National Institutes of Health registry eligible for federal funding support. Enko Kiprilov is a doctoral candidate in the Albert Einstein College of Medicine medical scientist training program and is supported in part by The National Institutes of Health (grant T32-GM007288). Romain Desprat is a doctoral candidate in the Sue Golding Division of the Albert Einstein College of Medicine. The studies by the team at the University of Copenhagen were supported in part by Rigshospital Science Foundation

purifi ed polyclonal anti – � -tubulin rabbit anti – human IgG [BioLegend]; puri-fi ed polyclonal anti-zinc fi nger protein Gli2 rabbit anti – human IgG [Aviva Systems Biology]; monoclonal anti – Tra-1-85 mouse anti – human IgG1 [Millipore]; and PE-conjugated monoclonal anti – stage-specifi c embryonic antigen 4 mouse anti – human IgG3 [R & D Systems]) were added in 1:300 dilution in blocking buffer for 1 h at RT or overnight at 4 ° C. Rabbit anti – human Oct-3/4 polyclonal IgG (Santa Cruz Biotechnology, Inc.), rabbit anti – human Smo polyclonal IgG (Santa Cruz Biotechnology, Inc.), and rab-bit anti – human SHh antibody polyclonal IgG (Cell Signaling Technology) were used at dilutions of 1:100 in blocking buffer and incubated overnight at 4 ° C. The cells were then washed three times in PBS with 5-min incuba-tions between washes. The secondary antibodies Cy3-conjugated Affi ni-Pure goat anti – mouse IgG (H + L) and Cy5-conjugated Affi niPure goat anti – rabbit IgG (H + L) (Jackson ImmunoResearch Laboratories) were added at 1:400 dilution in blocking buffer and incubated for 1 h at RT in the dark. All appropriate controls were done for the IF experiments described. Negative controls consisted of cells incubated with secondary antibody only. The cells were then washed again three times in PBS with 5-min incubations between washes and taken for IF imaging or stored at 4 ° C. The cells were incubated in DAPI (1:1,000 dilution) for 15 min in PBS before imaging. IF imaging was performed on an inverted (IX70; Olym-pus) and a confocal microscope (described in detail in Confocal micros-copy; TCS SP2 AOBS; Leica) and viewed at a fi nal magnifi cation of 600 using CY3 (red) and 5 (far red) fl uorescence fi lters. A cooled charge- coupled device camera (Sensicam QE; Sony) and IP Laboratory software (BD Biosciences) were used to capture the images, whereas ImageJ (National Institutes of Health) and Photoshop CS2 version 9.0.2 (Adobe) were used to view and analyze the data.

IF microscopy (Copenhagen) After 1 wk of incubation, hESCs on 16-well glass slides were washed once with PBS (136.89 mM NaCl, 2.68 mM KCl, 8.1 mM Na 2 HPO 4 , and 1.7 mM KH 2 HPO 4 ) and then fi xed with 4% paraformaldehyde for 20 min. After three 5-min washes with PBS, the wells were permeabilized with 0.1% Triton X-100 for 20 min. After three 5-min washes with PBS, the wells were blocked with 4% FBS for 45 min. Wells were incubated overnight at 4 ° C in the following primary antibodies: monoclonal mouse anti-AcTb at 1:10,000; polyclonal goat anti-pericentrin, polyclonal goat anti-centrin, polyclonal rabbit anti-Gli2, polyclonal rabbit anti – OCT-4, polyclonal rabbit anti-Ptc (Santa Cruz Biotechnology, Inc.) at 1:200; and polyclonal rabbit anti-Smo (MBL International) at 1:200. The next day, cells were washed fi ve times with PBS and allowed to stand 5 min, followed by three more quick washes with PBS. The cells were incubated 1 h with the follow-ing secondary antibodies: Alexa Fluor 488 – conjugated goat anti – rabbit IgG, Alexa Fluor 488 – conjugated donkey anti – goat IgG, and Alexa Fluor 568 – conjugated goat or rabbit anti – mouse IgG (1:600; Invitrogen); and coumarin/aminomethylcoumarin acetate – conjugated donkey anti – rabbit IgG (Jackson ImmunoResearch Laboratories). Secondary antibody incubation was occa-sionally followed by DAPI incubation. Cells were visualized on a micro-scope (Eclipse E600; Nikon) with EPI-FL3 fi lters and a cooled charge-coupled device camera (MagnaFire; Optronics), and digital images were processed using Photoshop.

Confocal microscopy (Albert Einstein College of Medicine) Images were collected with a confocal microscope (TCS SP2 AOBS) with 60 × oil immersion optics. Laser lines at 488, 543, and 633 nm for excita-tion of DAPI, Cy3, and Cy5, respectively, were provided by an Ar laser and a HeNe laser. Detection ranges were set to eliminate crosstalk be-tween fl uorophores.

SAG stimulation (Copenhagen) Confl uent cultures of LRB003 cells were incubated in the presence of 1 μ M SAG (Qbiogene) for 0, 1, and 4 h, followed by IF microscopy analysis with rabbit anti-Smo and anti-Ptc. Primary cilia were visualized with anti-AcTb and nuclei with DAPI. All images were taken with equivalent time exposures.

Exposure of H1 hESCs to SHhN, SAG, and cyclopamine (Albert Einstein College of Medicine) Recombinant human SHh (C24II), amino terminal peptide (SHhN; R & D Systems), and SAG were dissolved in PBS containing 0.1% BSA. Cyclopa-mine (Toronto Research Chemicals) was dissolved in 95% ethanol. SHhN, SAG, and cyclopamine, in medium containing 0.5% serum, were applied to H1 hESCs in culture (in triplicate) at concentrations of 5 μ M, 10 μ g/ml, and 1 μ M, respectively, for 18 h. The exposure time and concentrations used were derived from Lipinski et al. (2006) .

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Satir , P. , and S.T. Christensen . 2007 . Overview of structure and function of mam-malian cilia. Annu. Rev. Physiol. 69 : 377 – 400 .

Schneider , L. , C.A. Clement , S.C. Teilmann , G.J. Pazour , E.K. Hoffmann , P. Satir , and S.T. Christensen . 2005 . PDGFR � signaling is regulated through the primary cilium in fi broblasts. Curr. Biol. 15 : 1861 – 1866 .

Singla , V. , and J.F. Reiter . 2006 . The primary cilium as the cell ’ s antenna: signaling at a sensory organelle. Science . 313 : 629 – 633 .

Tanaka , Y. , Y. Okada , and N. Hirokawa . 2005 . FGF-induced vesicular release of Sonic hedgehog and retinoic acid in leftward nodal fl ow is critical for left-right determination. Nature . 435 : 172 – 177 .

Yao , S. , S. Chen , J. Clark , E. Hao , G.M. Beattie , A. Hayek , and S. Ding . 2006 . Long-term self-renewal and directed differentiation of human embryonic stem cells in chemically defi ned conditions. Proc. Natl. Acad. Sci. USA . 103 : 6907 – 6912 .

(A.G. Byskov and C.Y. Anderson), The Lundbeck Foundation (grant numbers 150/05 and R9-A969 to S.T. Christensen, A.G. Byskov, and A. Awan), and funds from the University of Copenhagen (S.T. Christensen and C.A. Clement).

Submitted: 6 June 2007 Accepted: 4 February 2008

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3070 Research Article

IntroductionHeart development in vertebrates is initiated in embryos shortly

after gastrulation by aggregation of cardiomyocyte progenitor cells

that become allocated from the mesodermal population (Sucov,

1998). The mouse embryonal carcinoma (EC) P19 cell line is a

common cell model system to study early heart differentiation in

vitro because the P19 EC cells can differentiate into beating

cardiomyocytes when stimulated with dimethyl sulfoxide (DMSO)

(Skerjanc, 1999; Paquin et al., 2002). The heart transcription factors

Gata4 and Nkx2-5 are markers for early cardiomyocyte

differentiation (Grépin et al., 1997; Lints et al., 1993). Gata4 is a

tissue-restricted transcription factor that is found in the heart but

not in skeletal muscle (Grépin et al., 1994) and is necessary for

proper heart tube development at the ventral midline (Kuo et al.,

1997). The Gata4 and Nkx2-5 genes are, together with MEF2C,

desmin and cardiac actin, expressed in the cardiomyocyte population

before fusion of the linear heart tube (Lyons, 1994). Mice lacking

Gata4 and Nkx2-5 die because of severe defects in heart formation

(Sucov, 1998).

A series of signal transduction systems have been implicated as

essential coordinators of early cardiogenesis, including hedgehog

(Hh), Wnt, bone morphogenetic protein (BMP) and platelet-derived

growth factor receptor (PDGFR) signaling (Washington Smoak et

al., 2005; Kwon et al., 2008; Hirata et al., 2007; van Wijk et al.,

2007). In mammals, Hh signaling is induced by three different

ligands, including Sonic hedgehog (Shh), which controls left-right

asymmetry, digit patterning in the limbs and development of the

lung and heart (Tsukui et al., 1999; Johnson et al., 1994; Bellusci

et al., 1997; Washington Smoak et al., 2005). In the adult mouse

heart, Hh signaling is required for proangiogenic gene expression

and maintenance of the adult coronary vasculature, and it

specifically controls the survival of small coronary arteries and

capillaries (Lavine et al., 2008). In vertebrates, the secreted Hh

proteins bind to the transmembrane patched protein-1 (Ptc1) hereby

abolishing the inhibitory effect of Ptc1 on the seven-transmembrane

receptor Smoothened (Smo). This allows Smo to transduce a signal

via Gli transcription factors to the nucleus for expression of Hh

target genes. There are three Gli transcription factors, Gli1-Gli3.

Gli1 functions as a constitutive activator (Hynes et al., 1997; Ruiz

I Altaba, 1999), whereas Gli2 and Gli3 have an N-terminal

transcriptional repressor domain and a C-terminal transcription

activator domain. Smo might be the controlling molecule in the Hh

signaling pathway that mediates the proteolytic events between the

activating and repressing form of Gli2 and Gli3 in an Hh-dependent

manner (Huangfu and Anderson, 2006). P19 EC cells normally

require aggregation to form embryoid bodies in suspension induced

by DMSO before differentiation analysis (Skerjanc, 1999).

However, overexpression of Shh has been observed to induce the

expression of cardiac muscle factors Gata4 and Nkx2-5 via Gli1and

Gli2, which results in differentiation of the cells into cardiomyocytes

in the absence of DMSO (Gianakopoulos and Skerjanc, 2005). This

supports the conclusion that Hh signaling is critical during early

Defects in the assembly or function of primary cilia, which are

sensory organelles, are tightly coupled to developmental defects

and diseases in mammals. Here, we investigated the function

of the primary cilium in regulating hedgehog signaling and early

cardiogenesis. We report that the pluripotent P19.CL6 mouse

stem cell line, which can differentiate into beating

cardiomyocytes, forms primary cilia that contain essential

components of the hedgehog pathway, including Smoothened,

Patched-1 and Gli2. Knockdown of the primary cilium by Ift88

and Ift20 siRNA or treatment with cyclopamine, an inhibitor

of Smoothened, blocks hedgehog signaling in P19.CL6 cells, as

well as differentiation of the cells into beating cardiomyocytes.

E11.5 embryos of the Ift88tm1Rpw (Ift88-null) mice, which form

no cilia, have ventricular dilation, decreased myocardial

trabeculation and abnormal outflow tract development. These

data support the conclusion that cardiac primary cilia are

crucial in early heart development, where they partly coordinate

hedgehog signaling.

Supplementary material available online at

http://jcs.biologists.org/cgi/content/full/122/17/3070/DC1

Key words: Primary cilia, P19.CL6 cells, Cardiac development,

Mouse, Heart, Hedgehog signaling, siRNA, Ift88, Ift20, Cyclopamine

Summary

The primary cilium coordinates early cardiogenesisand hedgehog signaling in cardiomyocytedifferentiationChristian A. Clement1, Stine G. Kristensen1, Kjeld Møllgård2, Gregory J. Pazour3, Bradley K. Yoder4,Lars A. Larsen5 and Søren T. Christensen1,*1Department of Biology, University of Copenhagen, Universitetsparken 13, DK-2100 Copenhagen, Denmark2Department of Cellular and Molecular Medicine and 5Wilhelm Johannsen Centre for Functional Genome Research, University of Copenhagen,Blegdamsvej 3, DK-2200 Copenhagen, Denmark3University of Massachusetts Medical School Worcester, Worcester, MA 01655, USA4The University of Alabama at Birmingham, Birmingham, AL 35294, USA*Author for correspondence ([email protected])

Accepted 29 May 2009Journal of Cell Science 122, 3070-3082 Published by The Company of Biologists 2009doi:10.1242/jcs.049676

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JCS ePress online publication date 4 August 2009

3071The primary cilium in cardiogenesis

cardiogenesis. Therefore, P19 EC cells offer a unique model cell

system to investigate the mechanisms by which Hh signaling is

coordinated by the cells during differentiation in early cardiogenesis.

Recent reports have indicated that primary cilia have an important

role in an array of vertebrate developmental processes. Primary cilia

are microtubule-based organelles, organized in a 9+0 axonemal

ultrastructure, which are assembled and maintained via a process

termed intraflagellar transport (IFT) in most mammalian cells during

growth arrest (Rosenbaum and Witman, 2002; Pedersen et al., 2008).

Primary cilia are thought to function as mechano- and chemosensory

organelles that specifically coordinate a series of cellular signal

transduction pathways during development and in tissue

homeostasis, including Hedgehog (Hh), PDGFRα and Wnt

signaling (reviewed by Christensen et al., 2007; Christensen et al.,

2008; Wong and Reiter, 2008; Gerdes and Katsanis, 2008).

Consequently, defects in assembly of the primary cilium or

mutations in ciliary signaling components lead to severe

developmental diseases and disorders, now referred to as ciliopathies

(reviewed by Pan, 2008; Lehmann et al., 2008; Davenport and

Yoder, 2005). One of the first diseases to be related to dysfunctional

primary cilia, was polycystic kidney disease (PKD), which was

originally identified in mice mutated in the gene encoding

Ift88/Polaris in the Oak Ridge Polycystic Kidney mouse (ORPK

mouse, Ift88orpk or Ift88Tg737NRpw) (Moyer et al., 1994). Ift88 is a

subunit of the IFT particle complex B required for functional IFT

and assembly of the primary cilium (Pazour et al., 2000; Murcia et

al., 2000; Haycraft et al., 2001; Taulman et al., 2001; Yoder et al.,

2002; Lucker et al., 2005). No other function of Ift88 is known,

and genes encoding IFT are found only in organisms that possess

cilia.

Many of the essential Hh signaling components, such as Gli2,

Gli3, Smo and Ptc, localize to primary cilia in a number of cell

types, including fibroblasts (Haycraft et al., 2005; Rohatgi et al.,

2007), epithelial cells in renal tubules (Harris and Torres, 2009)

and the exocrine duct of the pancreas (Nielsen et al., 2008), as well

as in human embryonic stem cells (Kiprilov et al., 2008; Breunig

et al., 2008). It was suggested that the concerted movement of Smo

and Ptc into and out of the cilium creates a switch by which cells

can turn Hh signaling on and off during development and tissue

homeostasis (Corbit et al., 2005; Rohatgi et al., 2007; Christensen

and Ott, 2007). In this scenario, binding of ligands to Ptc in the

cilium might activate the Hh pathway by removal of Ptc from the

cilium in a process that is associated with ciliary enrichment of

Smo (Rohatgi et al., 2007). In vitro activation of Smo in cells

exposed to Shh is blocked in mouse embryonic fibroblasts (MEFs)

lacking Ift172 or the dynein retrograde motor, Dync2h1 (Ocbina

and Anderson, 2008). The heterotrimeric kinesin complex

comprising the motor subunits Kif3a and Kif3b and the nonmotor

protein KAP is responsible for microtubule-based anterograde

translocation destined for membranous organelles, as well as for

ciliogenesis (Yamazaki et al., 1995; Haraquchi et al., 2006).

Furthermore, in mice lacking Kif3a and Smo there is a failure

in the maturation of radial astrocytes that would normally develop

into the dentate gyrus, which is responsible for maintenance of adult

neurogenesis (Han et al., 2008). Disruption of Kif3a results in severe

developmental abnormalities in the neural tube, cardiovascular

insufficiencies and randomized left-right development (Takeda et

al., 1999; Nonaka et al., 1998). Consequently, mutations in IFT

proteins and other proteins associated with the cilium and the

centrosome might result in dysfunctional Hh signaling and/or Gli

processing with severe developmental disorders in mammals,

including skeletogenesis (Gouttenoire et al., 2007), limb

development (Haycraft et al., 2007), neural tube formation

(Gorivodsky et al., 2008), cerebellar development (Chizhikov et

al., 2007; Spassky et al., 2008), mammary gland development and

defects in ovarian function (Johnson et al., 2008). Recently,

Brueckner and co-workers (Slough et al., 2008) showed that the

embryo heart at embryonic day 9.5 (E9.5) in Kif3a–/– mice has

abnormal development of endocardial cushions (ECCs) and reduced

trabeculation, indicating that primary cilium could coordinate

processes in cardiac morphogenesis.

Since Kif3 family proteins regulate cellular processes in

mammalian cells that are not necessarily related to the primary

cilium (Teng et al., 2005; Haraguchi et al., 2006; Corbit et al., 2008),

there is a need for a more thorough investigation on the role of the

primary cilium in early cardiogenesis and Hh signaling, which is

crucial for cardiomyocyte differentiation. In this study, we

investigated the role of the primary cilium in Hh signaling and early

cardiogenesis by: (1) characterization of primary cilia and their role

in Hh signaling and differentiation of the pluripotent P19.CL6 cell

line, an isolated subclone from the P19 cell line, into cardiomyocytes

and (2) microscopy analysis of defects in heart development in E11.5

embryos from wild-type (WT) and Ift88-null (Ift88tm1Rpw) mice.

P19.CL6 cells have no requirement for being cultured in suspension

and form embryoid bodies before differentiation (Uchida et al.,

2007). This allowed us to follow the function of the primary cilium

in the initial phases of differentiation from day 1 over a 2 week

period, until beating cardiomyocytes formed. By analyzing the

protein and mRNA levels of Gata4, Nkx2-5 and α-actinin, we can

determine the effect on cardiogenesis when shutting down Hh

signaling with cyclopamine treatment, a Smo-specific antagonist

(Chen et al., 2002), or knocking down primary cilia by Ift88 and

Ift20 siRNA. Ift20 is associated with the Golgi complex, and

knockdown of this IFT particle reduces ciliary assembly without

affecting Golgi structure (Follit et al., 2006). We show here that

P19.CL6 cells form primary cilia and that Hh signaling components

such as Ptc1, Smo and Gli2 localize to the cilia in these cells.

Furthermore, cyclopamine halts Hh signaling, preventing P19.CL6

cells from forming beating clusters of cardiomyocytes. Ift88 and

Ift20 siRNA nucleofection strongly reduced the assembly of primary

cilia, mRNA and protein levels of Gata4, Nkx2-5 and α-actinin,

expression of Ptc1 and Gli1, and nuclear localization of Gli1.

Furthermore, the induced loss of primary cilia with Ift88 and Ift20

siRNA reduced and delayed the number of beating clusters of

cardiomyocytes. In E11.5 embryos of Ift88-null (Ift88–/–) mice,

which lack primary cilia, we saw ventricular dilation, abnormal

outflow tract development and abnormal myocardial trabeculae

morphology compared with the wild type. These data support the

conclusion that primary cilia are crucial for differentiation of

P19.CL6 cells into cardiomyocytes and in early heart development

in the mouse, partly via coordination of Hh signaling.

ResultsP19.CL6 cell morphology and the effect of cyclopamine oncardiomyocyte differentiationTo investigate the cellular events during early heart development,

we studied the morphology and changes in expression of cardiac

transcription factors in P19.CL6 cell cultures during their

differentiation into cardiomyocytes over a 2 week period. We first

analyzed the effect of cyclopamine on Hh signaling and

cardiomyogenesis (Fig. 1). Addition of 1% DMSO to the growth

medium of P19.CL6 cells led to the formation of 15-20 beating

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clusters of cardiomyocytes at day 12 on average in a 9.6 cm2 culture

dish (Fig. 1A,L). At this time point, the individual clusters had a

diameter of 0.2-0.6 mm and the clusters could be observed in small

Journal of Cell Science 122 (17)

networks beating synchronously at a frequency of about 60 rhythmic

contractions per minute. Comparable structures of cardiogenic cell

clusters were observed in differentiating P19.SI cells, another

Fig. 1. Morphology of P19.CL6 cells during cardiomyocyte differentiation and the effect of cyclopamine on heart transcription factors. (A) Light microscopeimages of P19.CL6 cell morphology during differentiation at day 1, 5, 8 and 12. Arrow indicates a cluster of beating cardiomyocytes. (B) Immunofluorescencemicroscopy (IF) analysis of Gata4 localization at day 1, 5, 8, 12 (Gata4, green; DAPI, blue). (C) IF analysis of Nkx2-5 localization at day 1, 5, 8 and 12 (Nkx2-5,red) DF. (D) IF analysis of α-actinin and glutaminated tubulin at day 1, 5, 8, and 12 (α-actinin, red; Glu tb, green). (E) IF analysis of α-actinin localization mergedwith light microscope image of beating cardiomyocyte cluster at day 12 (α-actinin, red; dotted line marks edge of cluster). (F) High magnification ofcardiomyocyte cell at day 12 (arrow indicates primary cilium; open arrow, Z-line in α-cardiac muscle stress fibers). (G) Quantitative RT-PCR analysis of Gata4,Nkx2-5 and α-actinin relative mRNA levels (data from one representative experiment of n>3). (H) Western blot analysis (WB) of anti-Gata4 (~53 kDa), Nkx2-5(~40 kDa), α-actinin (~100 kDa) and β-actin (~43 kDa) reactivity on P19.CL6 cells at day (D) indicated. (I) Light microscope images of P19.CL6 cell morphologyduring differentiation at day 1, 5, and 12 with added 0.3, 1, 3 or 10 μM cyclopamine. Arrow indicates beating cluster of cardiomyocytes. (J) Quantitative RT-PCRanalysis on Gata4, Nkx2-5 and α-actinin relative mRNA levels at day 1 and 9 in control and 3 μM cyclopamine-treated P19.CL6 cells. **P<0.01; ***P<0.001.(K) WB analysis using anti-Gata4 (~53 kDa), Nkx2-5 (~40 kDa), α-actinin (~100 kDa) and β-actin (~43 kDa) on P19.CL6 cells at day 12 in control and 3 μMcyclopamine treated P19.CL6 cells. (L) Bar graph showing number of beating cardiomyocyte clusters at day 1, 5, 8, 11 and 12. Data from one representativeexperiment are shown. (M,N) IF analysis of Gata4 and Nkx2-5 localization (M) at day 12 in control and 3 μM cyclopamine-treated P19.CL6 cells (Gata4, green;Nkx2-5, red; DAPI, blue). (N)α-actinin localization at day 12 in control and 5 μM cyclopamine-treated P19.CL6 cells (α-actinin, red; DAPI, blue).

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subclone of P19 cells, which contracted synchronically by about 2

weeks (Angello et al., 2007). The network between clusters in

differentiated P19.CL6 cells later develop into a thick unified layer

of cardiac muscle epithelium (data not shown). Analysis of Gata4

and Nkx2-5 by immunofluorescence microscopy (Fig. 1B-C),

showed that Gata4-positive cells were present from day 2 onwards

during cardiomyocyte differentiation. Nkx2-5-positive cells

appeared at a later stage (~day 9) during cardiomyogenesis. To verify

that the P19.CL6 cells form cardiomyocytes, we stained with an

antibody against the cardiomyocyte marker α-actinin, which

localizes in the Z-line on α-cardiac muscle stress fibers (Fig. 1D-

F). The structural organization of P19.CL6 cardiomyocyte muscle

fibers takes place at approximately day 12. Primary cilia, localized

with anti-detyrosinated tubulin (Glu-tub), label cells that have

entered growth arrest (Satir and Christensen, 2007) and were present

at all stages of heart development (Fig. 1D-F). This was particularly

prominent in cells that had formed contact with other cells, either

as confluent monolayers before the beginning of differentiation or

as multilayered cell clusters during the subsequent phases of

differentiation and formation of the beating cardiomyocyte. mRNA

levels of Gata4, Nkx2-5 and α-actinin were analyzed by quantitative

RT-PCR (Q-PCR) (Fig. 1G) and showed an increase over the 2

week growth period that matched the appearance of positive cells

observed by immunofluorescence microscopy. As a control, the

protein levels of Gata4, Nkx2-5 and α-actinin in western blot (WB)

analysis (Fig. 1H), followed the increase in mRNA levels and

localization intensities of the proteins upon immunofluorescence

analysis; Gata4 expression was upregulated after day 2 and Nkx2-

5 and α-actinin at around day 9. The effect of cyclopamine on

P19.CL6 morphology indicated that a concentration of 0.3 μM of

this Smo inhibitor was too low to suppress cardiomyocyte

development (Fig. 1I). However, at cyclopamine concentrations

above 1 μM, there were no beating clusters of cardiomyocytes, and

at 10 μM or more, the cyclopamine became toxic and affected cell

viability. Furthermore, cyclopamine concentrations equal to or

greater than 1 μM altered the cell morphology in the culture and

the cells aggregated in disorganized clusters that adhered poorly to

the culture dish. The addition of 3 μM cyclopamine to the culture

medium had a significant negative effect on the mRNA levels of

Gata4, Nkx2-5 and α-actinin at day 9, compared with levels in

control cells (Fig. 1J). A similar reduction in mRNA expression of

the cardiomyocyte markers was observed at day 12 (data not shown).

Furthermore, cyclopamine significantly reduced the protein levels

of Gata4, Nkx2-5 and α-actinin in cells analyzed at day 12 (Fig.

1K), indicating an inhibition of cardiomyogenesis. Normal

cardiomyocyte formation took place on average at day 12, whereas

upon addition of 3 μM cyclopamine, we never observed any beating

clusters of cardiomyocytes (Fig. 1L). There were no Gata4- and

Nkx2-5 positive cells when cells were treated with 3 μM

cyclopamine (Fig. 1M). Furthermore, a diffuse α-actinin staining

pattern was observed (Fig. 1N), with no α-cardiac muscle stress

fibers, suggesting that the cardiomyocyte sarcomeres failed to

develop in the presence of cyclopamine by inhibition of the Hh

pathway.

Stem cell markers during P19.CL6 differentiationTo verify that DMSO promotes the differentiation of P19.CL6 cells

into cardiomyocytes and not other cell lineages, we followed the

shift in cellular expression of stem cell markers into Gata4-positive

cells. Confirmation that all P19.CL6 cells remained undifferentiated

at day 1 was made by immunofluorescence analysis using the

transcription factors Sox2 and Oct4, both markers for

undifferentiated cells (Pesce and Scholer, 2001; Masui et al., 2007).

Both markers colocalized to the nuclei of all cells at day 1 (Fig.

2A), at which time no cells were positive for Gata4 (Fig. 1B; Fig.

2B, left panels). During differentiation at day 5 after DMSO

Fig. 2. Characterization of P19.CL6 stem cell lineage during cardiogenesis. (A) Immunofluorescence microscopy (IF) analysis of Sox2 and Oct4 localization at day1 (Oct4, green; Sox2, red; DAPI, blue). (B) IF analysis of Sox2 and Gata4 localization at day 1, 5, 12 during differentiation in the presence of 1% DMSO (Gata4,green; Sox2, red; DAPI, blue). (C) IF analysis of Sox2 and Gata4 localization at day 12 in growth medium without 1% DMSO (Gata4, green; Sox2, red; DAPI,blue).

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treatment, we observed a shift in the cellular expression of Sox2

to Gata4 such that cells were either Sox2 or Gata4 positive (Fig.

2B, middle panels). About 40% of the cells were Gata4 positive at

this time point of differentiation. At day 12, the number of Gata4-

positive cells increased to about 80%, whereas the remaining cells

expressed Sox2 (Fig. 2B, right panels). As a control, all cells left

in growth medium without DMSO for 12 days remained

undifferentiated (Fig. 2C). These results show that DMSO primarily

promotes the differentiation of P19.CL6 cells into cardiomyocytes

and not other cell lineages.

Cyclopamine inhibits the Hedgehog signaling pathway andheart development in P19.CL6 cellsThe results of cyclopamine treatment in P19.CL6 cells presented

in Fig. 1 indicate that the Hh signaling pathway is turned off (Fig.

3) and that normal cardiogenesis requires the Hh signaling pathway

to become activated, as previously suggested for P19 EC cells

(Gianakopoulos and Skerjanc, 2005). Elevated mRNA levels of Gli1

and Ptc1 during P19.CL6 differentiation at day 9 were no longer

apparent upon 3 μM cyclopamine treatment (Fig. 3A). The Hh

transcription factor Gli2 exists in a full-length activator form (Gli2-

A) and in repressor forms (Gli2-R), which are formed after C-

terminal degradation of the protein (Pan et al., 2006). We here show

that 3 μM cyclopamine at day 5 reduces the level of the full-length

form of Gli2 (~160 kDa), as judged by WB analysis with an antibody

directed against the internal region of Gli2 (Gli2-G20) that

recognizes only the full-length form of Gli2 (Nielsen et al., 2008)

(Fig. 3B). WB analysis was also performed at day 9 using an

antibody directed against the N-terminal region of Gli2 (Gli2-N20),

which recognizes processed forms of Gli2 (Nielsen et al., 2008).


In this analysis Gli2-N20 detected a protein band at ~60 kDa, which

increased in intensity in the presence of cyclopamine (Fig. 3C).

Both protein bands were eliminated by addition of blocking peptide

to the antibodies. These results might indicate that inhibition of Hh

signaling by cyclopamine treatment is associated with processing

of Gli2 into repressor forms, although further analysis will be

required to identify their function in Hh signaling.

The effect of cyclopamine on Gli1 expression was also

investigated by immunofluorescence analysis at day 2, 5 and 9 of

differentiation. The level of Gli1 expression greatly increased in

the nucleus at day 5 and 9 (Fig. 3D-F), and this increase was largely

abolished in the presence of 3 μM cyclopamine. Confirmation that

Gli1 increased in cells that differentiate into cardiomyocytes was

made by immunofluorescence analysis, which showed

colocalization of Gli1 and Nkx2-5 in cells at day 9 (Fig. 3G).

Similarly, Gli2 and Gli3 expression levels at day 5 in the absence

and in the presence of 3 μM cyclopamine was observed (Fig. 3H-

I). We used Gli2 (G-20) and Gli3 (C-20) antibodies, which recognize

the full-length forms of each transcription factor. In both cases,

cyclopamine strongly reduced the nuclear localization of Gli2 and

Gli3. These results support the conclusion that cyclopamine prevents

P19.CL6 cells from differentiating into cardiomyocytes by shutting

down Hh signaling in the early phases of differentiation.

Hedgehog signaling components localize to primary cilia inP19.CL6 cellsHh signaling was previously suggested to be coordinated by

primary cilia (Liu et al., 2005). To confirm formation of primary

cilia in cultures of P19.CL6 cells, we initially performed

immunofluorescence analysis using anti-pericentrin (Pctn), which

Fig. 3. Effect of cyclopamine on the Hh-signaling pathway in P19.CL6 cells. (A) Quantitative RT-PCR analysis of Ptc1 and Gli1. Relative mRNA levels on day 1and 9 in control and 3 μM cyclopamine-treated P19.CL6 cells. *P<0.05; **P<0.01. (B,C) Western blot analysis using (B) anti-Gli2 (G-20 ~160 kDa) full-lengthand β-actin (~43 kDa) reactivity on P19.CL6 cells on day 5 (markers: 70, 80, 90, 100, 120, 160, 220 kDa), with a blocking peptide control (BP, 60� Abconcentration) and (C) anti-Gli2 (N-20 ~55 kDa) repressor and β-actin (~43 kDa) on day 9 (markers: 50, 60, 70, 80, 90, 100 kDa) with BP control. (D-F) Immunofluorescence microscopy (IF) analysis of Gli1 localization in control and 3 μM cyclopamine-treated P19.CL6 cells on day 2 (D), day 5 (E) and day 12(F). Gli1, green; DAPI, blue. (G) IF analysis of Gli1 and Nkx2-5 localization on day 9. Gli1, green; Nkx2-5, red. (H,I) IF analysis of Gli2(G-20) (H) andGli3(C-20) (I) localization on day 5 in control and 3 μM cyclopamine-treated P19.CL6 cells.

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labels the centrosome, and anti-acetylated α-tubulin, which labels

primary cilia in growth-arrested cells (Schneider et al., 2005). As

shown in Fig. 4A, primary cilia were clearly observed in cells either

in confluent monolayers or in multilayered cell clusters, as also

indicated with Glu-tub in Fig. 1D-F. Co-localization studies with

α-tubulin and antibodies directed against Ptc1, Smo and Gli2

showed that all three Hh components localize to the primary cilium

as indicated in Fig. 4B-D. Immunofluorescence analysis showed

the ciliary localization of Ptc-1 and Gli2 in cells expressing Gata4

and Nkx2-5, respectively (Fig. 4E-F), which confirms that Hh

components localize to cilia in cells differentiating into

cardiomyocytes. We also observed that the intensity of ciliary Ptc1

and Smo fluctuated in the cell cultures during cardiogenesis. This

was particularly evident around the forming cell clusters, which

tended to have more Smo and less Ptc1 in the cilium (data not

shown). At the end stage of differentiation, however, we also

observed that ciliary Ptc1 often increased in intensity in the cilium.

These observations could indicate that the primary cilium in

P19.CL6 cells is part of the signaling machinery that coordinates

activation of the Hh pathway during differentiation, and that Ptc1

participates in a negative regulatory feedback inhibition in the

developed cardiomyocytes. In this scenario, either newly expressed

and/or pre-existing Ptc-1 might translocate to the primary cilium

at the end stage of differentiation, although further analysis is

required to examine the importance of changes in ciliary

localizations of Hh components during differentiation of P19.CL6

cells into beating cardiomyocytes.

Knocking down the primary cilium with Ift88 and Ift20 siRNAprevents cardiogenesisTo investigate more directly the importance of primary cilia in early

cardiogenesis, we investigated the effects of knockdown of

Ift88/Polaris and Ift20 on differentiation of P19.CL6 cells into

cardiomyocytes. Using antibodies against Ift88 and Ift20 (Fig. 5A)

we initially observed that Ift88 predominantly localizes to the ciliary

base and tip, whereas Ift20 had a centrosome and Golgi localization

at the primary cilium in P19.CL6 cells. This localization is identical

to that observed in other cell types (Pazour et al., 2000; Taulman

et al., 2001; Follit et al., 2006; Haycraft et al., 2005). Confirmation

that Ift88 and Ift20 localized to the cilia, centrosome and Golgi in

cells that differentiate into cardiomyocytes was made by

immunofluorescence analysis, which showed localization of both

IFT proteins in cells that express Nkx2-5 at day 9 (Fig. 5B).

For knockdown studies, we nucleofected cells either with a

siRNA construct targeting Ift88 alone or in combination with a

siRNA construct targeting Ift20 (Ift88+Ift20); the Ift88 construct

expresses GFP. Then, we followed the effect of the knockdown

constructs on expression rates of Ift88 and Ift20, assembly rates of

primary cilia, expression rates of Gata4, Nkx2-5 and α-actinin, and

rates of beating cardiomyocytes. Using Q-PCR analysis, we found

that Ift88 and Ift88+Ift20 knockdown reduced the level of Ift88

mRNA to about 50% compared with mock-transfected cells under

both conditions 3 days after nucleofection (Fig. 5C). Ift20 mRNA

was reduced to about 25% when transfected with Ift88+Ift20 siRNA

but it was not affected by Ift88 nucleofection alone. These results

indicate that Ift88 siRNA does not affect the expression of Ift20

and vice versa. Furthermore, western blot analysis showed that the

protein levels of Ift88 (~95 kDa) and Ift20 (~16 kDa) are

significantly reduced by siRNA nucleofection (Fig. 5D). As a control

for the Ift88 antibody, we performed western blot analysis on

NIH3T3 cells, wt MEFs and Ift88Tg737NRpw MEFs (Tg737orpk MEF)

showing that the Ift88 protein band at 95 kDa is absent in mutant

fibroblasts (Fig. 5E). This has also been reported in prx1cre;Ift88fl/n

conditional mutants (Haycraft et al., 2007). Immunofluorescence

analysis was performed to show that cells nucleofected with the

Ift88 siRNA construct expressing GFP grew no or very short cilia

(<1 μm) compared with mock-transfected cells, which formed cilia

of ~4 μm in length (Fig. 5F). Furthermore, in mock-transfected

cultures, about 70% of the cells formed primary cilia, which was

reduced to about 30% in Ift88-transfected cells (Fig. 5G). A

combination of both Ift88 and Ift20 siRNA reduced the frequency

of ciliated cells further to about 20%.

Q-PCR analysis demonstrated that knockdown of the primary

cilium was associated with a reduction of mRNA expression rate of

Gata4 to about 40% and 25% relative to mock-transfected cells

with Ift88 and Ift88+Ift20 siRNA, respectively (Fig. 6A).

Immunofluorescence analysis confirmed that cells nucleofected with

the Ift88 siRNA construct expressing GFP were Gata4 negative (Fig.

6B) and Sox2 positive (Fig. 6C), indicating that knockdown of the

primary cilium maintains cells in their undifferentiated state. In

Fig. 4. Ciliary localization of Hh-signaling components in P19.CL6 cells. (A) Immunofluorescence microscopy (IF) analysis of acetylated tubulin (tb) on primarycilia after 24 hours (tb, red; pericentrin, green; DAPI, blue). Asterisk indicates centrosome region and arrows show primary cilium. (B) IF analysis of Ptc1localization to the primary cilium. (C) IF analysis of Smo localization to the primary cilium. (D) IF analysis of Gli2 localization to the primary cilium. (E) IFanalysis of Ptc-1 and Gata4 localization at day 12. (F) IF analysis of Gli2(H-300) and Nkx2-5 localization at day 12. Arrows in B-F indicate the primary cilium.

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addition, Ift88+Ift20 siRNA effectively blocked expression and

nuclear localization of Gata4 (Fig. 6D). Similarly, Q-PCR and

western blot analysis showed that Ift88+Ift20 siRNA reduced the

mRNA and/or protein levels of Nkx2-5 and α-actinin at day 9 (Fig.

6E,F). Finally, we analyzed the number of clusters of beating

cardiomyocytes after addition of siRNA (Fig. 6G). The cultures were

scanned daily for beating clusters; day [X] marking the day of the

first observed beating cluster in a 9.6 cm2 culture dish. The cells were

then recounted for beating clusters the day after [X+1]. The time

point for formation of beating clusters could vary a few days from

one experiment to another, although all experiments showed beating

clusters, on average, at day 12. In a screen of more than eight

independent experiments, we observed no major differences in the

time point for appearance of beating clusters in mock-treated versus

non-treated cells. As shown in Fig. 6G, both Ift88 and Ift88+Ift20

siRNA reduced the number of beating cardiomyocytes from about

12 beating clusters in mock-transfected cells to about 3 and 1.5 beating

clusters on average in Ift88 and Ift88+Ift20 siRNA nucleofected cells,

respectively, at day X+1. The size of the beating clusters in siRNA

nucleofected cells was reduced to about 20% of the size of clusters

in mock-transfected cells, and no networks were observed between

individual clusters. These results support the conclusion that primary

cilia are crucial regulators of P19.CL6 cardiogenesis.

Knockdown of Ift88 and Ift20 blocks Hedgehog signalingTo further investigate the significance of primary cilia in

coordination of Hh signaling in P19.CL6 cardiogenesis, we initially


analyzed the effect of Ift88+Ift20 siRNA on the expression levels

of Ptc1 mRNA (Fig. 7A) and Gli1 mRNA (Fig. 7B) at day 5 of

differentiation by Q-PCR analysis. In both cases, knockdown of

the cilium largely reduced the expression levels of both Hh signaling

markers to about 12% and 4%, respectively, relative to mock-

transfected cells. Immunofluorescence analysis demonstrated that

Ift88+Ift20 siRNA blocked the expression and nuclear localization

of Gli1 at day 5 (Fig. 7C), which was comparable with that observed

in the presence of cyclopamine (Fig. 3E). Therefore, the primary

cilium might control P19.CL6 cardiogenesis partly by coordinating

the Hh signaling machinery.

Heart defects in E11.5 Ift88-null miceThe Ift88-null (Ift88tm1Rpw, Ift88–/–) mouse has multiple

developmental phenotypes including random left-right axis

specification, neural tube closure and patterning abnormalities,

hepatic and pancreatic ductal defects, polydactyly, cerebellar

hypoplasia and retinal degeneration because of malfunctioning or

loss of primary cilia (Lehman et al., 2008). Since knockdown of

Ift88 severely inhibits cardiomyogenesis in P19.CL6 cells (Fig. 6),

we hypothesized that the loss of Ift88 in Ift88tm1Rpw (Ift88–/–) mice

would cause heart defects. This hypothesis was tested by

investigating cardiac tissue sections from WT and Ift88–/– embryos

at day E11.5 (Fig. 8). In the embryonic hearts of homozygous Ift88–/–

mice (Fig. 8A,B), we observed malformations of the cardiac

outflow tract (OFT) and the ventricles. The length of the distal

truncus in Ift88–/– mice was significantly shorter compared with

Fig. 5. Ift88 and Ift20 knockdown by siRNA nucleofection in P19.CL6 cells. (A) Immunofluorescence microscopy (IF) analysis of Ift88 (top panel) and Ift20protein (bottom panel) localization in P19.CL6 cells [Ift88+Ift20, green; acetylated tubulin (tb), red; DAPI, blue]. (B) Localization of Ift88 and Nkx2-5 (top panel)and Ift20 and Nkx2-5 (bottom panel) in P19.CL6 cells (Ift88 and Ift20, green; Nkx2-5, red; DAPI, blue). Arrows in A,B indicate primary cilium. (C) QuantitativeRT-PCR analysis on Ift88 and Ift20 relative mRNA levels after Ift88 and Ift88+Ift20 siRNA nucleofection vs mock treatment after 72 hours. (D) Western Blot(WB) analysis on day 5 of Ift88 (~95 kDa), Ift20 (~16 kDa) and β-actin (~43 kDa) reactivity on P19.CL6 cells after Ift88+Ift20 siRNA nucleofection vs mocktreatment. (E) WB analysis of Ift88 (~95 kDa) and β-actin (~43 kDa) reactivity on NIH3T3, wt and ORPK (Tg737orpk, Ift88Tg737Rpw) MEF cells after 24 hours ofserum starvation. (F) IF analysis of acetylated tubulin localization combined with GFP-Ift88-siRNA nucleofection (GFP-Ift88 siRNA, green; tb, red; DAPI, blue).Arrows indicate primary cilium; open arrow indicates missing primary cilium. (G) Bar graph showing the percentage of ciliated cells in mock, Ift88 and Ift88/Ift20siRNA nucleofected cells after 24 hours. **P<0.01; ***P<0.001.

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that of WT mice, and the OFT cushions seemed to be malformed,

with a thinner appearance (Fig. 8A). The WT mice had expanded

cardiac cushion tissue and a distinct transformation of endocardial

cells into mesenchyme, whereas the epithelial-mesenchymal

transformation (EMT) was virtually absent in the Ift88–/– mice. The

ventricles of Ift88–/– mice appeared dilated and empty because of

greatly reduced ventricular trabeculation compared with that in WT

mice (Fig. 8B). Furthermore, there was an increased volume of the

pericardial space in the Ift88–/– embryo compared with that of the

WT embryo (arrowheads in Fig. 7B). We verified that the mouse

embryos were of the correct genotype (Fig. 8C) and

immunohistochemical analysis showed that the Ift88-null mice had

no or very short primary cilia, as expected (Fig. 8D).

Immunohistochemical analysis also showed that Gli2 localizes to

primary cilia in the developing heart of WT embryos, such as in

the ventricular body wall (Fig. 8E, top panel). This localization is

absent in Ift88–/– embryos (Fig. 8E, bottom panel). These in vivo

findings support the conclusion that primary cilia have an important

role in cardiogenesis by coordinating Hh signaling.

DiscussionPrimary cilia have a critical role in the coordination of a number

of developmental processes in mammals, such as embryonic

left/right determination, skeletal patterning, limb formation and

neurogenesis (Nonaka et al., 1998; Gouttenoire et al., 2007;

Haycraft et al., 2007; Breunig et al., 2008). Brueckner and co-

workers (Slough et al., 2008) demonstrated that E9.5 Kif3a–/– mouse

embryos have abnormal development of ECCs and reduced

trabeculation, indicating that primary cilia in the heart could

regulate processes in cardiac morphogenesis, because Kif3a is

required for ciliary assembly amongst other cellular functions

(Slough et al., 2008). Here, we studied the function of the primary

cilium in early cardiogenesis by examining differentiated clusters

of cardiomyocytes, and by analysis of defects in heart development

in Ift88-null E11.5 mouse embryos, which form no or very short

primary cilia.

Primary cilia in P19.CL6 stem cell differentiationP19.CL6 stem cells spontaneously differentiate into clusters of

beating cardiomyocytes in the presence of DMSO (Habara-Ohkubo,

1996). We show that P19.CL6 stem cell form primary cilia and that

essential components of the Hh pathway, Ptc1, Smo and Gli2,

localize to P19.CL6 cilia. This is the first discovery of primary cilia

in this cell line. Before cardiomyocyte differentiation, the cells are

kept in their pluripotent state, as evidenced by expression of the

stem cell markers Sox2 and Oct4. Inhibition of the Hh pathway by

cyclopamine blocks DMSO-induced differentiation of P19.CL6

cells into clusters of beating cardiomyocytes by obstructing the

expression and nuclear localization of the heart transcription factors

Gata4 and Nkx2-5, which mark early cardiomyocyte differentiation

(Grepin et al., 1997; Lints et al., 1993). Further, cyclopamine altered

the cell morphology; cells aggregated in disorganized clusters

associated with a decreased expression of α-actinin and a lack of

α-actinin organized into the Z-line on α-cardiac muscle stress fibers,

which indicate fully differentiated cardiomyocytes. The inhibitory

effect of cyclopamine on Hh signaling in P19.CL6 cells was

Fig. 6. Effect of Ift88 and Ift20 knockdown by siRNA nucleofection on cardiomyocyte formation in P19.CL6 cells. (A) Quantitative RT-PCR analysis on Gata4relative mRNA levels after nucleofection with Ift88 siRNA alone or with Ift88+Ift20 siRNA vs mock control on day 5. (B) IF analysis of Gata4 expression in Ift88nucleofected P19.CL6 cells, 72 hours after nucleofection (GFP-Ift88 siRNA, green; Gata4, red; DAPI, blue). (C) IF analysis of Sox2 expression in Ift88nucleofected P19.CL6 cells 72 hours after nucleofection (GFP-Ift88 siRNA, green; Sox2, red; DAPI, blue). (D) IF analysis of Gata4 expression in Ift88+Ift20nucleofected cells vs mock control at day 5 (Gata4, red; DAPI, blue). (E) Quantitative RT-PCR analysis on Nkx2-5 relative mRNA levels after Ift88 and Ift88+Ift20siRNA nucleofection vs mock control at day 5. (F) WB analysis of Gata4 (~53 kDa, day5), Nkx2-5 (~40 kDa, day 12), α-actinin (~100 kDa, day 12) and β-actin(~43 kDa) reactivity on P19.CL6 cells after Ift88+Ift20 siRNA nucleofection vs mock control. (G) Number of beating cardiomyocyte clusters in mock, Ift88 andIft88+Ift20 nucleofected P19.CL6 cells. Day [X] symbolizes the first day where beating clusters were observed in a culture, the cultures were recounted on thefollowing day (Day [X+1]). *P<0.05; **P<0.01; ***P<0.001.

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confirmed by inhibition of DMSO-induced upregulation of Gli1

and Ptc1 mRNA expression and nuclear localization of Gli1, Gli2

and Gli3. In support of the conclusion that cyclopamine inhibits

Hh signaling by processing of Gli2, we used western blot analysis

to show that the level of the full-length form of Gli2, which might

function as an activator form of Gli2, is reduced in cyclopamine-

treated cells. Concomitantly, in the presence of cyclopamine, we

observed an increase in a processed form of Gli2, which might

represent an inhibitory form of Gli2 (Pan et al., 2006). These results

confirm that Hh signaling is required for differentiation of P19.CL6

cells into cardiomyocytes, and that Hh signaling may be associated

with primary cilia in P19.CL6 cells.

To determine the function of the primary cilium in regulation of

Hh signaling and in cardiomyogenesis, we carried out experiments

in which the primary cilium was knocked down by Ift88 and Ift20

siRNA. Initially, we showed that Ift88 uniquely localizes to primary

cilia and that Ift20 localizes to the centrosome and Golgi region in

P19.CL6 cells, as previously shown in differentiated cells (Pazour

et al., 2000; Follit et al., 2006; Haycraft et al., 2005). Ift88 is a

subunit of the IFT particle complex B and is required for functional

IFT and ciliary assembly (Pazour et al., 2000; Lucker et al., 2005).

Ift20 functions in the delivery of ciliary membrane proteins from

the Golgi complex to the cilium and strong knockdown of this IFT

particle blocks ciliary assembly without affecting Golgi structure

(Follit et al., 2006). Ift20 is anchored to the Golgi complex by the

golgin protein GMAP210, and mice lacking GMAP210 die at birth

with a pleiotropic phenotype that includes ventricular septal defects

of the heart, although cells have normal Golgi structure (Follit et

al., 2008). Knockdown of Ift88 in P19.CL6 cells reduced the


frequency of ciliated cells by more than 50% and inhibited the

expression levels of both Gata4 and Nkx2-5 to about 40% at day

5 of transfection, and this was associated with a decrease in nuclear

localization of Gata4 followed by a reduced number of beating

cardiomyocytes at around day 12. We also observed that cells

transfected with Ift88 siRNA were Sox2 positive, indicating that

knockdown of the cilium maintains cells in their undifferentiated

state and the cells then do not undergo apoptosis or differentiate

into other cell lineages. We next performed double knockout of Ift88

and Ift20 to show that an augmented reduction of primary cilia is

associated with an additional decrease in mRNA expression levels

of Gata4 and Nkx2-5 and numbers of clusters of beating

cardiomyocytes in accordance with a strong reduction in protein

levels of Gata4, Nkx2-5 and α-actinin. Since knockdown of Ift88

and Ift20 produces an inhibitory response on Hh signaling that is

similar to that of cyclopamine in P19.CL6 cells at day 5 of

differentiation (i.e. strongly reduces the expression levels of Ptc1

and Gli1 as well as the nuclear localization of Gli1), we conclude

that differentiation of P19.CL6 cells into cardiomyocytes is

governed by the primary cilium, partly by regulation of Hh

signaling.

A number of observations have indicated a crucial function of

primary cilia in differentiation processes in mammalian stem cells.

Human embryonic stem cells (hESCs) possess primary cilia

(Kiprilov et al., 2008) that contain a series of signal transduction

components, including PDGFRα (Awan et al., 2009) and members

of the Hh and Wnt signaling systems (Awan et al., 2009; Kiprilov

et al., 2008), which are important in stem cell maintenance,

differentiation and proliferation. It was further shown that primary

cilia are crucial for the development of neural stem cells needed

for proper development of the hippocampal region (Han et al., 2008)

and in development of the neocortex and cerebellum (Spassky et

al., 2008). These results imply that stem cell primary cilia per se

might coordinate cellular processes in early development, including

cardiogenesis. The mechanism by which Ptc1, Smo and Gli2

coordinate Hh signaling in the cilium of P19.CL6 is presently not

understood. Our preliminary data suggest that Ptc1 and Smo

become differentially localized to the cilium during the

differentiation stages towards formation of beating cardiomyocytes,

as previously described for cilia after stimulation of the Hh pathway

in hESCs, fibroblasts, MDCK cells and epithelial cells from the

exocrine ducts of the human pancreas (Haycraft et al., 2005;

Huangfu and Anderson, 2005; Liu et al., 2005; May et al., 2005;

Rohatgi et al., 2007; Nielsen et al., 2008). The regulated movement

of Ptc1 out of the cilium and Smo into the cilium might create a

switch by which cells can regulate Gli processing and turn Hh

signaling on (reviewed by Christensen and Ott, 2007). Further

experiments are required to investigate this in detail to understand

how the primary cilium might function as a specialized organelle

that integrates positive and negative inputs on Hh signaling in

P19.CL6 cell differentiation.

Primary cilia are important for cardiogenesis in vivoThe cardiac phenotype of E11.5 Ift88–/– embryos, where ciliogenesis

is inhibited, resembles in part, the cardiac phenotypes of Pkd2–/–

and Kif3a–/– mice (Slough et al., 2008), which have malformed

endocardial cushions, decreased trabeculation and increased

pericardial space. Slough et al. (Slough et al., 2008) showed, by

comparison to lrd–/– embryos, that decreased trabeculation and

increased pericardial space are not related to abnormal development

of the left-right asymmetry, which is initiated at the node of the

Fig. 7. Effect of knockdown of Ift88 and Ift20 on Hh signaling in P19.CL6cells. (A,B) Quantitative RT-PCR analysis of Ptc1 (A) and Gli1 (B) relativemRNA levels after Ift88+Ift20 siRNA nucleofection vs mock control on day 5.(C) Immunofluorescence microscopy analysis of Gli1 expression in Ift88-Ift20nucleofected P19.CL6 cells compared with mock control on day 5 (Gata4,green; DAPI, blue). **P<0.01.

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mouse at E7.75 and coordinated by nodal cilia. Therefore, our results

on the Ift88–/– mouse support these findings (Slough et al., 2008)

and suggest that these defects in cardiac morphogenesis are not

caused by defects in nodal cilia, but in cardiac primary cilia in the

developing heart. We also show that that Ift88–/– E11.5 embryos

have malformations of the cardiac outflow tract (OFT), indicating

that primary cilia also have an essential role in formation of the

OFT. Brueckner and colleagues (Slough et al., 2008) did not

investigate OFT malformations.

A series of signal transduction pathways have been implicated

in the morphogenesis of the embryonic heart after establishment of

the left-right asymmetry, some of which could be coordinated by

the cardiac primary cilium. Our in vitro data on P19.CL6 cells show

that Hh signaling is one of the ciliary pathways necessary for

cardiomyocyte differentiation, and we surmise that ciliary Hh

signaling in a similar manner coordinates in vivo morphogenesis

of the heart. As an example, Gli2 localizes to primary cilia in both

P19.CL6 cells and in the developing heart of WT embryos, and this

localization is disrupted in Ift88–/– embryos. Interestingly, the OFT

phenotype of Ift88–/– embryos resembles that of Shh–/– mice

(Washington Smoak et al., 2005; Goddeeris et al., 2007), suggesting

that aberrant Hh signaling due to defects in assembly of primary

cilia is involved in shortening of the OFT and potentially in other

cardiac structures in Ift88–/– embryos. Indeed, cardiac expression

of Nkx2-5 during heart development is blocked in the Smo–/– mouse

embryo at the 2- to 3-somite stage (which corresponds to E9),

suggesting that Nkx2-5 is a specific cardiac target of Hh signaling

and when blocked, underlies the defective heart morphogenesis

observed in Smo mutants (Zhang et al., 2001). Similarly, our results

show that knockdown of the primary cilium in P19.CL6 cells

inhibits Hh signaling and blocks the expression of Nkx2-5,

supporting the conclusion that primary cilia have an important role

in heart development, in part by coordinating Hh signaling. Whether

the primary cilium regulates cardiogenesis by coordinating other

signaling pathways, such as Wnt, BMP and PDGF signaling, is

presently unknown. Wnt and PDGF signaling are regulated by the

cilium in a series of other cell types involved in growth control,

migration and differentiation (reviewed by Christensen et al., 2008;

Gerdes and Katsanis, 2008). BMP promotes the induction of

cardiomyocytes from the mouse stem cell line P19.SI (Angello et

al., 2007) and human embryonic stem cells (Takei et al., 2009).

Based on their findings on Pkd2–/– mouse embryos, Slough and

colleagues (Slough et al., 2008) also hypothesized that primary cilia

in the heart and/or vasculature can function as mechanosensors to

Fig. 8. Cardiac morphology of WT and Ift88tm1Rpw-null embryos. (A) Comparable 3-μm-thick longitudinal mid-saggital sections of E11.5 WT and Ift88tm1Rpw

(Ift88–/–) embryos. BW, body wall; L, liver; LA, left atrium; OFT, outflow tract; PS, pericardial space (arrowhead); SV, sinus venosus; T, tongue; V, ventricle.(B) Comparable 3-μm-thick longitudinal parasagittal sections of day E11.5 WT and Ift88-null mice. Arrowheads indicate the pericardial space (PS), which isextended in Ift88-null mouse. The black bars have identical dimensions in A and B. The distal end of the OFT are marked with a dotted line. (C) Genotyping ofE11.5 WT and Ift88-null mice. Wild-type samples have a 120 bp band whereas Ift88-null has a 270 bp band. (D) Immunofluorescence microscopy analysis ofacetylated tubulin (tb) on primary cilia in the heart of E11.5 WT and Ift88-null mice (tb, red; DAPI, blue). Arrow indicates primary cilium and open arrow showsmissing primary cilium. (E) Immunofluorescence microscopy analysis of Gli2(H-300) and acetylated tubulin (tb) on primary cilia in the heart of WT and Ift88-nullmice (tb, red; DAPI, blue). Arrow indicates primary cilium.

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3080 Journal of Cell Science 122 (17)

detect blood flow essential for cardiac morphogenesis. In this regard,

GMAP210 and Ift20 might function together at the Golgi in the

sorting or transport of polycystin-2 to the ciliary membrane (Follit

et al., 2008). Furthermore, primary cilia on endothelial cells (ECs)

in blood vessels might function as laminar shear stress sensors to

maintain heart homeostasis (Iomini et al., 2004). Cilia in cultures

of embryonic ECs contain polycystin-1, which when subjected to

shear stress, is cleaved, leading to changes in cellular signaling

processes (Nauli et al., 2008). These results support the conclusion

that primary cilia have a major role during development and in

homeostasis of the heart.

We show here that primary cilia are crucial for coordination of

early cardiogenesis. Knockdown of Ift88 and Ift20 blocks assembly

of primary cilia and differentiation of P19.CL6 stem cells into

clusters of beating cardiomoycytes. Knockout of Ift88 and defective

assembly of primary cilia in the E11.5 Ift88–/– mouse lead to cardiac

malformation, most probably as a consequence of defective ciliary

Hh signaling. The fact that other signaling pathways, such as Wnt,

BMP and PDGFR signaling, are crucial in cardiogenesis warrants

further investigation on the signaling systems coordinated by the

primary cilium, and whether these signaling systems impinge on

ciliary Hh signaling in heart development.

Materials and MethodsAnimals and tissue sectioningWild-type and Ift88-null (Ift88tm1Rpw, Ift88–/–) embryos were isolated from timed

matings at E11.5. The Ift88-null embryos were generated from by intercrossing two

Ift88tm1Rpw heterozygotes congenic on the FVB/N genetic background. Tissue

sectioning was performed as described (Nielsen et al., 2008). Serial sections, 3 to 5

μm thick, were cut sagittally and the amniotic sac was used for genotyping.

Cell cultureThe P19.CL6 cell line is of mouse origin isolated from embryonal carcinoma tissue.

The originator is Habara, Akemi and registered with Murofushi, Kimiko, Japan (ref.

2406 3467). The cells were grown in T75 cell culture flasks (Cellstar) at 37°C, 5%

CO2 and 95% humidity. Cells were cultured in MEM Alpha medium (Gibco),

containing 1% penicillin-streptomycin (Gibco) and 10% foetal bovine serum (FBS,

Gibco). The medium was supplemented with 1% dimethyl sulfoxide (DMSO, Merck)

to induce cardiomyocyte differentiation. The cells were passaged every 2-3 days by

trypsination (Trypsin-EDTA, Gibco). Swiss NIH3T3 mouse fibroblasts and primary

cultures of embryonic fibroblasts (MEFs) from WT and ORPK (Ift88Tg737RPW,

Tg737orpk) mice were cultured as described (Schneider et al., 2005). Experimental

cells were seeded at a confluency of about 30%.

Primary antibodiesAntibodies from Santa Cruz: rabbit anti-Gli3 (H-280) (Cat. no. SC-20688), goat anti-

Gli2 (G-20) (SC-20291), goat anti-Gli3 (N-19) (SC-6155), goat anti-Gli2 (N-20) (SC-

20290), goat anti-Gli3 (C-20) (SC-6154), rabbit anti-Gli2 (H-300) (SC-28674), goat

anti-patched (G-19) (SC-6144), rabbit anti-Gata4 (H-112) (SC-9053), goat anti-Nkx-

2,5 (N-19) (SC-8697); goat anti-Oct3/4 (C-20) (SC-8629); rabbit anti-Gli1 (Abcam

AB14149); rabbit anti-Smo (MBL International LS2666); mouse anti-α-actinin

(Sarcomeric) Sigma (A-7811); mouse anti-acetylated tubulin, Sigma (T7451); rabbit

anti-detyrosinated α-tubulin (Glu-tubulin, Chemicon (AB3201); goat anti-Sox2

(R&D systems) (AF2018), mouse anti-Sox2 (R&D systems) (MAB2018). Secondary

antibodies for immunofluorescence from Molecular Probes: Alexa Fluor® 568

Donkey IgG, anti-mouse (A10037), Alexa Fluor® 488 Donkey IgG, anti-goat

(A11055), Alexa Fluor® 568 Donkey IgG, anti-rabbit (A21206). Secondary antibodies

for western blot analysis: goat anti-mouse F(ab�)2 specific alkaline phosphatase

conjugated, Sigma (A1293), Goat anti-rabbit F(ab�)2 specific alkaline phosphatase

conjugated, Sigma (A3937), Rabbit anti-goat whole molecule alkaline phosphatase-

conjugated, Sigma (A4187). DAPI: 4�,6-diamidino-2-phenylindole, dihydrochloride

(Molecular Probes, D1306). Blocking peptides (BPs) (Santa Cruz): BP for goat anti-

Gli2 (G-20) (SC-20291-P); BP for goat anti-Gli2 (N-20) (SC-20290-P).

siRNA constructs and transfectionThe cells were transfected with the plasmids pGP678.12 and pJAF135.45 encoding

siRNA targeted at mouse Ift88 and Ift20, respectively, by nucleofection with the

Nucleofector device II from Amaxa Biosystems. We followed the recommended

protocol for P19 cell transfection and used a Nucleofector Kit V. The recommended

amount of cells for transfection was 2�106 and with 2 μg plasmid. Complementary

oligonucleotides corresponding to the coding region of mouse Ift20 and mouse Ift88

were annealed and cloned into pGP676.13 digested with BglII and HindIII to produce

pGP678.12 and pJAF135.45, respectively (Follit et al., 2006).

ImmunofluorescenceThe cells were grown on coverslips in six-well trays (TTP) and subjected to

immunofluorescence microscopy analysis as described (Schneider et al., 2005).

Pictures were captured with a cooled CCD Optronics camera on a Nikon-Japan,

Eclipse E600 epifluorescence microscope and the digital images processed with

Photoshop 6.0.

HistochemistryRepresentative sections of WT and Ift88tm1Rpw mice were stained with hematoxylin

and eosin (H&E). In preparation for immunohistochemistry, tissue sections were

heated for 1 hour at 60°C. Then, sections were deparaffinized for 10 minutes in xylene,

2 minutes in 99% ethanol, twice for 15 minutes in 99% ethanol, 15 minutes in 96%

ethanol, 15 minutes in 70% ethanol and a final 15 minutes in H2O. The sections were

then placed in a tub with running tap water for 10 minutes and in PBS for 10 minutes.

Sections were circled with a PAP pen and incubated in blocking buffer (PBS with

5% BSA) for 20 minutes. Immunohistochemistry was performed as described (Nielsen

et al., 2008).

SDS-PAGE and western blot analysisCells were grown in Petri dishes and were rapidly washed once in PBS and spun

down at 500 g for 5 minutes, after which 350 μl lysis buffer with 1% β-

mercaptoethanol was added. Proteins were purified using Nucleospin kit protocol

(Macherey-Nagel) for RNA or protein analysis. SDS-PAGE and western blotting were

performed as described (Schneider et al., 2005).

Quantitative RT-PCRRNA was purified using a Nucleospin kit (Macherey-Nagel). For cDNA synthesis

we used DNase I Amplification grade (Invitrogen) and SuperScriptTM II reverse

transcriptase (Invitrogen). The Q-RT-PCR reactions were performed on a 7500 fast

Realtime PCR-system from Applied Biosystems with Lightcycler Fast Start DNA

Master plus SYBR Green 1 (Roche). Primers used are listed in supplementary material

Table S1.

StatisticsWe used one-way analysis of variance (ANOVA) test on n=3 or more. Significance

levels were divided into three categories (*P<0.05, significant; **P<0.01, highly

significant; ***P<0.001, extremely significant).

This work was supported by the Lundbeck Foundation, the Danish

Science Research Council no. 272-07-0530 (S.T.C.), the Danish Heart

Association (L.A.L.), funds from the Department of Biology, University

of Copenhagen, Denmark (C.A.C.), by NIH RO1 HD056030 (B.K.Y.)

and by GM60992 (G.J.P.). Wilhelm Johannsen Centre for Functional

Genome Research is established by the Danish National Research

Foundation. The authors would like to thank Lillian Rasmussen,

Kirsten Winther, Venus Childress and Laura Smedegaard Kruuse for

excellent technical assistance. Deposited in PMC for release after 12

months.

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Journal of Cell Science 122 (17)3082

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CHAPTER 9

Using Nucleofection of siRNA Constructsfor Knockdown of Primary Cilia inP19.CL6 Cancer Stem Cell Differentiationinto Cardiomyocytes

Christian A. Clement*, Lars A. Larsen†, and Søren T. Christensen**Department of Biology, Section of Cell and Developmental Biology, University of Copenhagen, DK-2100Copenhagen OE, Denmark

†Wilhelm AU1Johannsen Centre for Functional Genome Research, University of Copenhagen, Copenhagen,Denmark

AbstractI. IntroductionII. RationaleIII. Materials

A. Cell Line and Cell Culture ReagentsB. Reagents and Solutions for NucleofectionC. Reagents and Solutions for IFM AnalysisD. Western BlotE. Quantitative RT-PCR

IV. MethodsA. Introductory Remarks and Experimental OutlineB. Cell Culturing and PassagingC. Nucleofection of P19.CL6 Cells with IFT80 and IFT20 siRNA (Plasmid DNA)D. Methods for Optimization with IFT88 siRNA Plasmid for High Transfection RateE. Nucleofection for Light and Immunofluorescence Microscopy AnalysisF. Nucleofection for Western Blot AnalysisG. Nucleofection for Quantitative Real-Time RT-PCR (qPCR) Analysis

V. Results and DiscussionA. Timetable and Markers of Differentiation in P19.CL6 CellsB. Nucleofection with IFT88 and IFT20 siRNA and Its Consequences on Ciliary

Formation in P19.CL6

Font: Bembo & Times Size:10/12pt Margins:Top:5pc Gutter:5pc2 T.Area:34pc8�43pc6.5 1 Color Lines: 44 Fresh Recto

Els US MCB-94 Chapter 9 15-10-2009 18:28 Page: 175 Trim: 190mm�235mm Floats: Top/Bottom TS: Integra, India

METHODS IN CELL BIOLOGY, VOL. 94 978-0-12-375024-2Copyright � 2009 Elsevier Inc. All rights reserved. 175 DOI: 10.1016/S0091-679X(08)94009-7

C. Nucleofection with IFT88 and IFT20 siRNA and Its Consequences on Differen-tiation and Hh Signaling in P19.CL6 Cells

VI. SummaryAcknowledgmentsReferences

Abstract

Primary cilia assemble as solitary organelles in most mammalian cells during growtharrest and are thought to coordinate a series of signal transduction pathways requiredfor cell cycle control, cell migration, and cell differentiation during development and intissue homeostasis. Recently, primary cilia were suggested to control pluripotency,proliferation, and/or differentiation of stem cells, which may comprise an importantsource in regenerative biology. We here provide a method using a P19.CL6 embryoniccarcinoma (EC) stem cell line to study the function of the primary cilium in earlycardiogenesis. By knocking down the formation of the primary cilium by nucleofectionof plasmid DNAwith siRNA sequences against genes essential in ciliogenesis (IFT88and IFT20) we block hedgehog (Hh) signaling in P19.CL6 cells as well as thedifferentiation of the cells into beating cardiomyocytes (Clement et al., 2009). Immu-nofluorescence microscopy, western blotting, and quantitative PCR analysis wereemployed to delineate the molecular and cellular events in cilia-dependent cardiogen-esis. We optimized the nucleofection procedure to generate strong reduction in thefrequency of ciliated cells in the P19.CL6 culture.

I. Introduction

Primary cilia are organelles that emanate from the surface of most growth-arrestedmammalian cells. They consist of a microtubule (MT)-based axoneme organized in a9þ 0 axonemal ultrastructure ensheathed by a bilayer lipid membrane continuous withthe plasma membrane, but which contains a distinct subset of receptors and otherproteins engaged in signaling pathways in developmental processes and tissue home-ostasis. Primary cilia are formed via a process termed intraflagellar transport (IFT),which is essential for the assembly and maintenance of almost all eukaryotic cilia andflagella (Cole and Snell, 2009; Pedersen et al., 2008). Separating the two membranecompartments at the ciliary base is a region known as the “ciliary necklace” (Gilula andSatir, 1972), which is connected by fibers to the transition zone of the basal body,which may function as a pore where ciliary precursors and IFT proteins accumulateprior to entering the ciliary compartment via IFT, a process essential for assembly ofvirtually all cilia and flagella. IFT is a bidirectional transport system that tracks alongthe polarized MTs of the ciliary axoneme. IFT is composed of large protein complexes,known as IFT particles, and the motor proteins heterotrimeric kinesin-2 (kinesin-2) for

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176 Christian A. Clement et al.



anterograde (base to tip) transport of ciliary building blocks, and cytoplasmic dynein 2for retrograde (tip to base) transport of ciliary turnover products (Pedersen et al., 2008).The signaling pathways being coordinated by the developed primary cilium includeHh, Wingless (Wnt), platelet-derived growth factor receptor (PDGFR)a, Ca2þ, neuro-nal and purinergic receptor signaling, and communication with the ECM (Satir et al.,2009 AU2). Accordingly, defects in assembly or function of the primary cilium are a majorcause of human diseases and developmental abnormalities and disorders nowcommonly referred to as ciliopathies (reviewed in Lehman et al., 2008).

Recent observations indicate that primary cilia in stem cells coordinate signalingpathways, including Hh signaling, in cell differentiation during embryonic develop-ment and potentially in regulation of stem cell maintenance and/or pluripotency(reviewed in Veland et al., 2009). Stem cells hold great promises for their possibletherapeutical abilities since they can give rise to all three germinal layers and differ-entiate to form specific cell types dependent on the environment and specific factorspresent. Stem cells may also be important targets against cancer. Hh regulates cellproliferation and differentiation in numerous embryonic tissues and Hh ligands areexpressed in the notochord, the floorplate of the neural tube, the brain, the limb budzone of polarizing activity, and the gut (Odent et al., 1999). Hh signaling is furtherrequired in homeostasis of mature tissues and is implicated in human cancers (Beachyet al., 2004) and neurodegenerative disorders (Bak et al., 2003). A screen for embryo-nic patterning mutations characteristic of defective Hh signaling first indicated a linkbetween IFT proteins, Hh signaling, and nervous system development (Huangfu et al.,2003). Subsequent studies confirmed that Hh signaling is coordinated by the primarycilium to control targets of the Hh pathway by Gli transcription factors (Corbit et al.,2005; Huangfu and Anderson, 2005; Liu et al., 2005; reviewed in Wong and Reiter,2008). Functioning Hh components, including Ptc-1, Smo, and Gli transcriptionfactors are localized in primary cilia of human embryonic stem cells (Kiprilov et al.,2008) and neuronal development proceeds ciliary Hh signaling in adult neural stemcell formation, specification of neural cell fate, hippocampal neurogenesis and devel-opment of cerebellum and neocortex (Breunig et al., 2008; Han et al., 2008; Komadaet al., 2008; Spassky et al., 2008). Similarly, primary cilia are involved in thecoordination of Hh signaling, for example, in limb bud formation (Haycraft et al.,2007), skeletogenesis (Gouttenoire et al., 2007), mammary gland development andovarian function (Johnson et al., 2008), molar tooth number (Ohazama et al., 2009),and development of the pancreas (Nielsen et al., 2008).

Primary cilia and Hh signaling are also implicated in early cardiogenesis as evi-denced by defective heart development in knockout mice with defects in ciliaryassembly, including decreased trabeculation, increased pericardial space, and malfor-mations of the cardiac outflow tract (Clement et al., 2009). Further, knock down of theprimary cilium in the pluripotent P19.CL6 EC stem cell line blocked Hh signaling anddifferentiation of cells into beating cardiomyocytes in vitro (Clement et al., 2009). TheP19.CL6 cell line is a subclone from the P19 cell line that spontaneously differentiateinto clusters of beating cardiomyocytes in the presence of dimethyl sulfoxide (DMSO)(Habara-Ohkubo, 1996). Further, P19.CL6 cells have no requirement for being

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9. Knockdown of Primary Cilia in P19.CL6 Cells 177



cultured in suspension and form embryoid bodies before carrying out the analysis oncardiac differentiation (Uchida et al., 2007). This allows the investigator to follow thefunction of the primary cilium in the initial phases of differentiation from day 1through a 2-week period until the formation of beating cardiomyocytes.

II. Rationale

Here we provide a detailed and optimized method for nucleofecting P19.CL6 ECcells with IFT88 and IFT20 siRNA plasmid DNA to produce a high transfectionpercentage to knockdown primary cilia in cultures of P19.CL6 cells during theirdifferentiation into cardiomyocytes. IFT88 is a subunit of the IFT particle complex Brequired for functional IFT and assembly of the primary cilium (Pedersen andRosenbaum, 2008). IFT20 is associated with the Golgi apparatus, and knockdown ofthis IFT particle reduces ciliary assembly without affecting Golgi structure (Follitet al., 2006). Following knockdown of the cilium cell differentiation can be assessedby light microscopy (LM), immunofluorescence microscopy (IFM), SDS-PAGE,western blotting (WB), and quantitative PCR (qPCR) analysis in order to followchanges in DMSO-induced formation of beating cardiomyocytes, expression andlocalization of stem cell and cardiomyocyte markers, and activation of Hh signaling.

III. Materials

A. Cell Line and Cell Culture Reagents

The P19.CL6 cell line is of mouse origin isolated from embryonal carcinoma tissue.The originator is Habara, Akemi and registered with Murofushi, Kimiko, Japan (ref nr.2406 3467).

MEM Alpha medium (Gibco AU3, Cat#22561-021)Penicillin/streptomycin (Gibco, pen/strep, Cat#15140-148)Phosphate-Buffered Saline (PBS)Fetal bovine serum (FBS, Gibco, Cat#10 106-177)Trypsin (Trypsin-EDTA, Gibco, Cat#15 400-054)T75 cell culture flasks (Cell star AU4, Cat#658 170)T25 cell culture flasks (Cell star, Cat#690 175)6-well trays (TTP AU5, Cat# 92006)Petri dishes (Cell Star, 60� 15mm, Cat# 628 160)DMSO (MERCK AU6, Cat#1.02952.1000)

B. Reagents and Solutions for Nucleofection

Nucleofector device II (Amaxa Biosystems AU7)Nucleofector Kit V (Amaxa Biosystems)2 µg of IFT88 or IFT20 siRNA plasmid DNA (high grade, high concentration)MEM Alpha medium (Gibco, Cat#22561-021)

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C. Reagents and Solutions for IFM Analysis

Microscope slides and glass coverslips (12mm diameter)Concentrated HClHumidity chamberEthanol (96% v/v and 70% v/v)Paraformaldehyde (PFA), 4% w/v solutionBlocking buffer: 2% w/v Bovine Serum Albumin (BSA) in PBSPermeabilization buffer: 0.2% v/v Triton X-100, 1% w/v BSA in PBSDAPI (40,6-diamidino-2-phenylindole, dihydrochloride)Mounting medium (PBS, 2% w/v N-propylgallate, 85% v/v glycerol in PBS)Nail polishAntibodies and fluorescent reagents (Table I)

D. Western Blot

NOVEX system (XcellSure Lock AU8)Precast NuPAGE 10% and 12% BIS-TRIS 12-well gelsNucleospin kit (Macherey-Nagel AU9, Cat#740 933.50) for RNA/protein isolationBSA protein standard (Pierce Biotechnology, Rockford, IL, USA)Protein assay (BioRAD AU10, DC based on Lowry’s method)Running buffer (Invitrogen, Cat#NP0001)Transferbuffer (Invitrogen, Cat#NP0006-1)Non-fat dry milk blocking bufferNuPAGE Antioxidant (Invitrogen, Cat#NP0005)Ethanol (96% v/v)TBST AU11

Antibodies (Table I)

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Table IUsed Antibodies and Fluorescent Reagents

Primary Antibodies Dilution

Mouse anti-a-actinin (Sarcomeric), (Sigma AldrichAU21 , Cat#A-7811) 1:100Goat anti-Nkx2 (N-19), (Santa Cruz, Cat#SC-8697) 1:100Mouse anti-b-actin (Sigma Aldrich, Cat#A-5441) 1:5000Mouse antiacetylated Tubulin, (Sigma Aldrich, Cat#T7451) 1:1500Rabbit antidetyrosinated a-Tubulin (Glu-Tubulin, ChemiconAU22 , Cat#AB3201) 1:600Rabbit anti-IFT20 (see Follit et al. 2006) 1:1000

Secondary Antibodies and Fluorescent AgentsDonkey anti-goat (DAG), Alexa Flour® 488 (Molecular Probes, Eugene, OR, USA, Cat#A11055) 1:600Donkey anti-mouse (DAM), Alexa Flour® 568 (Molecular Probes, Cat#A10037) 1:600Goat anti-rabbit (GAR), F(ab0)2-specific Alkaline phosphatase-conjugated (Sigma Aldrich, Cat#A3937) 1:5000Goat anti-mouse (GAM), F(ab0)2-specific Alkaline phosphatase-conjugated (Sigma Aldrich, Cat#A1293) 1:5000DAPI: 40,6-diamidino-2-phenylindole, dihydrochloride (Molecular Probes, Cat#D1306) 1:1000




E. Quantitative RT-PCR

Nucleospin kit (Macherey-Nagel, Cat#740 933.50) for RNA/protein isolationSuperScriptTM II reverse transcriptase (Invitrogen, Cat#18064-014)DNase I Amplification grade (Invitrogen, Cat#18068-015)PCR-grade nuclease-free waterdNTP mix, 10mM of each dNTPRNAse inhibitor (rRNasin from Promega AU12)qPCR plate (Applied Biosystems, Foster City, CA, USA Cat#4346906)Lightcycler® FastStart DNA MasterPLUS SYBR Green I (Roche AU13, Cat#030515885001)RNA purifying kit (Macherey-Nagel, Cat#740 933.50) for RNA/protein Ammonium

Buffer with 15mM MgCl2 (Ampliqon III, Cat#AMP300305)TAQ-polymerase (5 units/µl, Ampliqon III)Agarose, ethidium bromide, TAE buffer (4.84 g Tris base, 1.14ml glacial acetic acid, 2ml

0.5 M Na2 EDTA pH 8.0, ddH2O up to 1 l), nucleic acid molecular weight markerPCR primers (Table II)Mouse universal reference total RNA (Stratagene AU14, Cat#740100)E.N.Z.A. gel extraction kit (Omega Bio-Tek inc., Norcross, GA, USA, Cat#D2500-00)

IV. Methods

A. Introductory Remarks and Experimental Outline

Growth arrest and formation of primary cilia in cultures of mammalian cells canbe induced either by depletion of serum and/or by growing cells to confluency. Incultures of P19.CL6 EC stem cells primary cilia are formed in the presence ofserum as the cells leave their pluripotent stage and enter the differential steps

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Table IIPCR Primers Used in this Study

Gene Forward primer (50–30) Reverse primer (50–30) Annealing temp. (°C)

Mef2c TGCCATCAGTGAATCAAAGG ATGTTATGTAGGTGCTGCTGC 58Myh6 CAAAGGAGGCAAGAAGAAAGG GTCCCCATAGAGAATGCGG 56Myh7 TGAGGGAACAGTATGAGGAGG TCGATCTCATTCTGCAGCC 60

Gene Forward primer (50–30) Reverse primer (50–30) Annealing temp. (°C)Ptc-1 CCTGCCCACCAAGTGATTGT CGTTGGGTTCCGAGGGTT 52Gli1 GCGGAAGGAATTCGTGTGCC CGACCGAAGGTGCGTCTTGA 62

Gene Forward primer (50–30) Reverse primer (50–30) Annealing temp. (°C)Gapdh AACAGCAACTCCCACTCTTC TGGTCCAGGGTTTCTTACTC 58B2m ATTTTCAGTGGCTGCTACTCG ATTTTTTTCCCGTTCTTCAGC 58Hprt CAAAATGGTTAAGGTGCAAGC TTTTACTGGCAACATCAACAGG 57Psmd4 GCAAGATGGTGTTGGAGAGC TTTGGGTTGGACAGTGTGG 59Rp13a GGAGAAACGGAAGGAAAAGG CTCTATCCACAGGAGCAGTGC 57Alas1 GGATCGGTGATCGGGATGGCGTCA AGGTGGTGAAGATGAAGCCCGCAGCGTA 60Pbgd2 CGTTTGCAGATGGCTCCAATAGTAAAG TGGCATACAGTTTGAAATCATTGCTATGT 60




induced by DMSO to form clusters of beating cardiomyocytes, most probablybecause physical contact between individual cells promotes the entrance into G0

(Clement et al., 2009). This allows the investigation of the function of primary ciliain stem cell differentiation and cardiomyogenesis in vitro by knocking down thecilium. Cultures of P19.CL6 cells in 9.6 cm2 Petri dishes may form up to about 20clusters of cardiomyocytes with a diameter of 0.2–0.6mm that beat synchronouslyat a frequency of about 60 rhythmic contractions per minute around day 12 in thepresence of DMSO (Clement et al., 2009). Prior to differentiation cells are positivefor stem cell markers Sox2 and Oct4, which are replaced by Gata4 positive cells atday 2, Nkx2–5 positive cells around day 8, and a-actinin positive cells marking theZ-line on a-cardiac muscle stress fibers around day 12 concomitantly with the onsetof contractions of the mini hearts. During these steps of cell proliferation anddifferentiation, the expression of the transcriptional target genes for Hh signaling,Gli1 and Ptc-1 are highly upregulated: both cardiomyocytes differentiation and Hhsignaling being blocked in the presence of the Smo antagonist and Hh-signalinginhibitor, cyclopamine (Clement et al., 2009).

In this protocol we performed siRNA knockdown of the primary cilium in P19.CL6 cells by nucleofection, which is a transfection method that enables efficientand reproducible transfer of nucleic acids such as DNA, RNA, and siRNA intocells. Nucleofection is also known as Nucleofector Technology, which wasinvented by Amaxa (http://www.amaxa.com). Nucleofection uses a combinationof optimized parameters generated by a device termed Nucleofector with cell-typespecific reagents and buffers. The substrate is transferred directly into the cellnucleus and cytosol. Here we nucleofected P19.CL6 cells with IFT88 and IFT20siRNA plasmid DNAs in order to knock down the cilium and follow its conse-quences in stem cell maintenance, cell differentiation, and Hh signaling. The IFT88plasmid DNA is expressing GFP, which enables quantification of nucleofectionrates and direct visualization of its consequences on ciliary formation and celldifferentiation. Here we present a detailed protocol on the nucleofection procedure,followed by qualitative and quantitative analysis on cell differentiation and Hhsignaling by IFM, SDS-PAGE, WB, and qPCR analysis. The protocol for qPCRanalysis is presented comprehensively, while IFM, SDS-PAGE, and WB analysesare portrayed in less detail.

B. Cell Culturing and Passaging

The cells were grown for passaging in T25 cell culture flasks at 37°C, 5% CO2, and95% humidity. The cells were passaged every 2–3 days by trypsination, and grown inMEM Alpha cell culture media, containing 1% penicillin/streptomycin and 10% FBS.Prior to trypsination the cells were washed once in 37°C PBS and reseeded at 10–15%confluency (avoid growing cells into 100% confluency). To induce cardiomyocytedifferentiation, the medium was supplemented with 1% DMSO and grown undernormal incubator conditions. Experimental cells were seeded at a confluency ofabout 30% in T75 cell culture flasks, 6-well trays, or Petri dishes.

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C. Nucleofection of P19.CL6 Cells with IFT80 and IFT20 siRNA (Plasmid DNA)

Sufficient cells were cultivated in T75 flasks, enough to provide each sample with2� 106 cells. Preferably, the cells should be passaged the day before nucleofection butif the experiments require the cells to enter AU15a stage of differentiation (e.g., DMSO-induced differentiation that needs siRNA knockdown at day 2–3), it is still possible tocarry out the nucleofection with a high transfection rate. This will disrupt cell culturemorphology since the cells will need to be resuspended in nuclofection solution andreseeded.Before the actual nucleofection step, prepare the following:

• 6-well trays/dishes with the appropriate working volume of differentiation media areput inside the incubator (37°C/5% CO2) and prewarmed 20min beforenucleofection.

• Prewarm the supplemental Cell Line Nucleofector Solution V to room temperature.• Thaw up your highly purified IFT88 and IFT20 siRNA plasmids. Amount of µg is

determined by optimization (e.g., 2 µg DNA per sample AU16, see Section IV.D).

The medium was removed from the T75 culture flasks and washed once in 37°CPBS. The cells were then trypsinated in 1�Trypsin-EDTA for 5min in 37°C/5% CO2

and resuspended in an appropriate volume of 37°C growth medium (preferably freshlymade). The volume was adjusted so that the cell density was high enough to collect2� 106 cells in a 1.5ml Eppendorf tube. The cells were counted in a hemocytometer todetermine the actual number of cells. The cells were then spun down at 90� g for10min at room temperature. The cell pellet was resuspended in 100-µl Cell LineNucleofector Solution V per 2� 106 cells per sample. Do not keep the cells in theNucleofector Solution V for more than 15min.Each sample is nucleofected in the following steps:

• Add 2 µg of siRNA plasmid DNA into the 100-µl solution containing the cell pellet.• Gently mix the DNA, cells, and Cell Line Nucleofector Solution V three times in a

1000-µl pipette and transfer the sample to an Amaxa-certified cuvette (make sure thesample covers the cuvette bottom with no air bubbles).

• Close the cuvette with the lid and insert into the Nucleofector device II and runprogram C-020.

• Transfer 500 µl media from the prewarmed 6-well tray/dish using the suppliedplastic pipettes into the cuvette and transfer the whole sample back into the 6-welltray/dish. Avoid transferring white foam “popcorn” to the 6-well tray/dish since thiscontains only cell debris

The 6-well trays/dishes are put into the incubator as soon as possible in 37°C/5%CO2 to provide as little stress as possible. Analysis of the samples and the effect ofIFT88 and IFT20 siRNA knockdown can be performed 24 h after nucleofection. Atthis time point the maximal effect of the IFT88 and IFT20 knockdown can be found.The optimal time for siRNA plasmid expression can be determined by selectingdifferent time points after nucleofection and verifying the level of mRNA knockdownwith IF analysis of a GFP-reporter inserted in the siRNA plasmid.

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D. Methods for Optimization with IFT88 siRNA Plasmid for High Transfection Rate

To find the optimal transfection rate, we used the pmaxGFPTM positive controlvector supplied by Amaxa biosystems. An experimental setup with variable amountsof pmaxGFPTM (1 µg, 1.5 µg, 2 µg, 2.5 µg, 3 µg, and 4 µg) was nucleofected into2� 106 cells. The cells were grown on coverslips in 6-well trays at 37°C/5% CO2 for24 h. The cells were fixed in 4% paraformaldehyde for 15min at RT, followed by twowashes in PBS, and then permeabilized in 0.2% Triton X-100, 1% BSA for 12min.Cells were then stained with DAPI to visualize total cell numbers on the coverslips andthe total-cell/transfected-cell ratio could hereby be determined. Also variable timepoints after nucleofection (12, 24, 36, 48, and 72 h) were tested to determine atwhich time point the transfection rate was the highest. The recommended DNA/cellnumber ratio supplied by Amaxa biosystems was 2� 106 cells with AU172-µg DNAplasmid followed by analysis after 24 h. The optimal transfection rate for our experi-ments was found under the recommended conditions and was close to 80%. Cellviability was good—estimated to ~70%. Furthermore, it is highly recommendable toperform both positive and negative nucleofection controls (cellsþ solutionþDNA –

program) (cells þ solution – DNA þ program) to assess influences of nucleofection orpurity of DNA on cell viability.

E. Nucleofection for Light and Immunofluorescence Microscopy Analysis

After nucleofection the cells were transferred to 6-well trays containing coverslipsand grown in 37°C/5% CO2 for at least 24 h to visualize the optimal effect of thetransfection. After 1, 5, 8, and 12 days of culturing in differentiation medium the cellswere subjected to IFM in order to analyze the frequency of ciliated cells. The protocolused for IFM and detection of primary cilia is identical to that described in Chapter 3by Thorsteinsson et al. (this volume). For detection of primary cilia we used primaryantibodies against acetylated a-tubulin (1:1500) and detyrosinated a-tubulin (1:600).Similarly, one can use antibodies directed against specific markers of cardiomyogen-esis in order to follow the time course for differentiation, and how nucleofection withIFT88 and IFT20 siRNA constructs impinge on stem cell maintenance and cardio-myocte differentiation with antibodies directed against Oct4 and Sox2 (stem cellmarkers) and Gata4, Nkx2–5, and a-actinin (cardiomyocytes markers) (Clement etal., 2009). Further, expression and localization of Hh signaling components such as Glitranscription factors are assessed by IFM and the formation of beating clusters ofcardiomyocytes is easily detected by light microscopy.

F. Nucleofection for Western Blot Analysis

After nucleofection the cells were transferred to Petri dishes and grown in 37°C/5%CO2 for at least 24 hours. Hereafter the cells were cultured for 1–12 days in differentia-tion medium. For WB analysis cells were washed in PBS, spun down at 500� g for5min, and then added with 350-µl lysis buffer with 1% b-mercaptoethanol. Proteinswere purified using Nucleospin kit protocol (Macherey-Nagel, Cat# 740 933.50) for

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RNA/protein. After protein precipitation the cells were added with 2% SDS, 1%Glycerol lysis buffer and sonicated, centrifuged at 20,000� g to precipitate nonsolublematerial. The protein concentration was compared with a BCA protein standard (PierceBiotechnology) and measured with a Protein assay so that an equal amount of proteincould be loaded in each lane. For SDS-PAGE a NOVEX system was used with precastNuPAGE 10% and 12% BIS-TRIS 12-well gels (Schneider et al., 2005).

G. Nucleofection for Quantitative Real-Time RT-PCR (qPCR) Analysis

After nucleofection the cells were treated as in the above for WB analysis in order toisolate total RNA accordingly to the Nucleospin kit protocol for RNA/protein and toassess transcriptional processes in Hh signaling and cell differentiation. RNA wastreated with DNase I Amplification grade and cDNA was produced from 1µg totalRNA using SuperScriptTM II reverse transcriptase. PCR primers for amplification ofhousekeeping genes (Gapdh, B2m, Hprt, Psmd4, Rp13a, Alas1, Pbgd2), cardiomyo-cyte markers (Mef2C, Myh6, Myh7) and Hh signaling genes (Smo, Ptc-1) weredesigned using Oligo version 6.23 (Table II). PCR annealing temperature was opti-mized in a QuatroCycler temperature gradient thermocycler (VWR, West Chester, PA,USA) using universal mouse cDNA as template. For each primer pair PCR fragmentswere excised from an agarose gel and extracted using an E.N.Z.A. gel extraction kit.Standard curves were generated by 10-fold serial dilutions of the PCR fragments. Thequantitative real-time RT-PCR (qPCR) reactions were performed in a 7500 fast Real-time PCR system (Applied Biosystems,) using a Lightcycler Fast Start DNA Masterplus SYBR Green 1 kit (Roche, Copenhagen, Denmark).For each gene the cycle threshold (Ct) value was converted to a relative expression (Er)

value using the standard curve. Due to the high sensitivity of qPCR, differences in theefficiency of the cDNA synthesis and differences in PCR reaction kinetics betweensamples may lead to incorrect quantitation of the expression. To correct for intersamplevariation, samples are normalized by dividing the Er value of the gene of interest with theEr value of an endogenous control. The ideal endogenous control for qPCR analysis is agene that displays similar reaction kinetics and expression profiles in all samples(VanGuilder et al., 2008). Usually, housekeeping genes are used as endogenous controls.However, not all housekeeping genes are resistant to experimental conditions; thus toensure a robust expression profile of the endogenous control in all samples, we normalizedusing the average Er value of at least three housekeeping genes with similar expressionprofiles across the samples. Only normalized Er values were compared in the experiments.

V. Results and Discussion

A. Timetable and Markers of Differentiation in P19.CL6 Cells

In order to delineate the onset for DMSO-induced P19.CL6 stem cell differentiationwe initially used light microscopy analysis to evaluate the time point for formation ofbeating clusters of cardiomyocytes. As depicted in Fig. 1A most cell cultures form

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beating clusters around day 12, clusters appearing in small networks that beat at afrequency of about 60 rhythmic contractions per minute. These results are consistentwith the findings that clusters of cells at this time point are positive for a-actinin thatmarks the Z-line on a-cardiac muscle stress fibers (Fig. 1C, upper panels) (Clementet al., 2009). To follow differentiation of P19.CL6 cells in more detail we thenmeasured the transcription of the cardiomyocyte markers Gata4, Nkx2-5, Actc2,

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Day 1

Day 1 Day 12

Day 12(A)

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glu-tbα-actininDAPI


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Fig. 1 Morphology and expressionAU23 profile of cardiomyocyte markers in P19.CL6 cells duringcardiomyocyte differentiation induced by 1% DMSO. (A) Light microscope images of P19.CL6 cellmorphology at day 1 prior to differentiation and at day 12 where cells have formed beating clusters ofcardiomyocytes (open arrow). (B) Quantitative RT-PCR analysis on Myh6, Myh7, and Mef2c mRNA levelsrelative to expression of housekeeping genes during days 1–10 of P19.CL6 differentiation.(C) Immunofluorescence microscopy analysis at days 1 and 12 of localization of a-actinin and primarycilia (lower panels) and a-actinin and Nkx2–5 (upper panels). Upper panels: Nkx2–5: green; a-actinin: red;DAPI: blue. Lower panels: primary cilia (arrows, detyrosinated a-tubulin, glu-tb): green; a-actinin: red,DAPI: blue. (See Plate no. xx in the Color Plate Section.)




Mef2C, Myh6, and Myh7 using qPCR. As exemplified in Fig. 1B cardiomyocytemarker genes are transcriptionally upregulated around day 4 after DMSO addition,their expression levels peaking around day 10. These results are in agreement withprevious data, showing that cardiomyocyte markers are upregulated at around day 2and 6 in P19.CL6 cells stimulated with DMSO (Clement et al., 2009). Finally, weperformed IFM analysis to show that clusters of cells either at day 1 in undifferentiatedcells or day 12 in differentiated cells (positive for Nkx2–5 and a-actinin; Fig. 1C,upper panels) formed primary cilia (Fig. 1C, lower panels) as evidenced by stainingwith antidetyrosinated a-tubulin (Glu-tb) that is highly enriched in primary cilia ofvertebrate cells (Gundersen and Bulinski, 1986). This allows the investigator toexamine the role of the primary cilium by RNAi methods in differentiation of P19.CL6 cells using qPCR analysis on the expression of cardiomyocyte genes, Myh6,Myh7, and Mef2c.

B. Nucleofection with IFT88 and IFT20 siRNA and Its Consequences on Ciliary Formation inP19.CL6

Nucleofection was performed with IFT88 and IFT20 siRNA plasmid constructs inorder to inhibit the formation of primary cilia in cultures of P19.CL6 cells and thensubsequently analyze the effect of ciliary knockdown on cardiomyocyte differentia-tion. In this regard there are a number of analyses to perform to ensure optimalknockdown efficiency and minimize unwanted effects on cell behavior, such as cellviability caused by the transfection method. As described in Section IV.D, transfectionrates can initially be determined with a pmaxGFPTM positive control vector, in whichthe rate of transfection can be monitored and estimated by green fluorescence innucleofected cells. In this analysis we found that the optimal transfection rate of80% was achieved by nucleofection of 2� 106 P19.CL6 cells with 2-µg plasmidDNA 2 days after the addition of DMSO to the cell cultures, followed by analysisafter 24 h. In this setup cell viability was estimated to about 70%. We then used theseparameters to analyze cell viability and transfection rate after nucleofection with IFT88siRNA GFP plasmid. The cells were brought into suspension and nucleofected with theplasmid, and after 24 h of subsequent cell culture the cells were fixed, stained withDAPI, and the transfection rate was calculated to be about 70–80% as judged by IFM(Fig. 2A). In total, the cells had 3 days of differentiation with 1% DMSO in the culturemedia, hence the day 3 in Fig. 2A.In order to examine the effect of IFT88 knockdown on formation of primary cilia,

we performed IFM analysis with antibodies directed against either detyrosinateda-tubulin or acetylated a-tubulin, the latter also marking primary cilia and other stablecellular MTs (Piperno and Fuller, 1985). As indicated in Fig. 2B, the number ofprimary cilia at day 6 was markedly reduced in cells nucleofected with IFT88siRNA plasmid as compared to mock transfected cells. Further, primary cilia emergingfrom cells in IFT88 siRNA-nucleofected cells were often shorter than in the controlcells, supporting the conclusion that IFT88 is required for ciliary assembly. Thepercentage of ciliated cells at day 3 was further enumerated by IFM analysis as

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shown in Fig. 2C. In controls, the percentage of ciliated cells was calculated to beabout 75%, whereas this number was reduced to about 30% in IFT88 siRNA-nucleofected cells. As a further control, we found no differences in cell viabilitybetween mock and IFT88 siRNA-nucleofected cells, which was estimated to beabout 70% in both cases (Clement et al., 2009). We then went on to perform a doubleknockdown of the primary cilium in P19.CL6 cells using both IFT88 and IFT20siRNA plasmid DNA. As indicated in Fig. 2C, the number of ciliated cells was furtherreduced to about 20%, showing that a reduction in both IFT88 and IFT20 has astronger inhibitory effect on ciliary formation than IFT88 alone.

The level of IFT20 knockdown by nucleofection can also be verified on protein levelswith WB analysis. We show here (Fig. 2D) an example of such an analysis of IFT20protein levels in cells subjected to IFT20 siRNA knockdown compared to mocktransfected cells on day 3, 6, and 10. Initially protein levels were greatly reduced onday 3 and 6 compared to control cells. At day 10, however, reduction in the protein levelwas less pronounced, indicating a loss in siRNA of IFT20. A similar phenomenon wasobserved with IFT88 siRNA (not shown). A plausible explanation for this is that aportion of the transfected cells may lose their siRNA plasmid over time as a consequenceof continuous cell divisions such that the number of plasmid-containing cells in the

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Fig. 2 NucleofectioAU24 n of IFT88 and IFT20 plasmid siRNA in P19.CL6 cells. (A) Immunofluorescencemicroscopy analysis of cells nucleofected with GFP-expressing IFT88 siRNA plasmid at day 3. Transfectionrate efficiency: ~70–80% (IFT88 siRNA GFP: green; DAPI: blue). (B) Immunofluorescence microscopyanalysis of primary cilia at day 6 of differentiation after nucleofection with mock and IFT88 siRNA (primarycilia were localized with antiacetylated a-tubulin, tb: red; DAPI: blue). (C) Bar graph showing the percentageof ciliated cells in mock, IFT88, and IFT88þ IFT20 siRNA-nucleofected P19.CL6 cells at day 3.Reproduced with permission from Clement et al. (2009). (D) Western blot analysis of IFT20 (~20 kDa)and b-actin (~43 kDa) protein levels in P19.CL6 cells after 3, 6, and 10 days of IFT20 siRNA nucleofectionversus mock transfected cells. (See Plate no. xx in the Color Plate Section.)




culture is reduced. This also means that the percentage of ciliated cells in nucleofectedcultures may increase over time around day 10. Since nucleofection requires that cellsare kept in suspension, it was not an option to perform a second round of nucleofectionof P19.CL6 cells at day 10 without disrupting cell morphology and clusters ofcardiomyocytes, which begin to form at this time point after DMSO stimulation.

C. Nucleofection with IFT88 and IFT20 siRNA and Its Consequences on Differentiation andHh Signaling in P19.CL6 Cells

To analyze the effects of IFT88 and IFT20 knockdown on cardiomyocte differentiationand Hh signaling we initially performed qPCR analysis on the mRNA levels of myocyteenchancer factor 2c (Mef2c) and myosin heavy chain 7 (Myh7). As shown in Fig. 3A the

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Fig. 3 Knockdown of IFT88 and IFT20 by siRNA nucleofection inhibits cardiomyocyte differentiationand Hh signaling in P19.CL6 cells. (A) Bar graph showing quantitative RT-PCR analysis on Mef2c andMyh7 mRNA levels relative to expression of housekeeping genes after siRNA nucleofection versus mock atday 5. (B) Number of beating cardiomyocyte clusters in mock, IFT88, and IFT88þ IFT20 siRNA-nucleofected P19.CL6 cells at day 12. (C) Bar graph showing quantitative RT-PCR analysis on Ptc1 andGli1 mRNA levels relative to expression of housekeeping genes siRNA nucleofection versus mock at day 5.Reproduced with permission from Clement et al. (2009).




mRNA levels of both markers of cardiomyocytes differentiation were largely reduced byabout 75 and 60%, respectively, compared to mock transfected cells. Similarly, it wasshown that IFT88 and IFT20 knockdown reduced the mRNA and/or protein levels ofGata4, Nkx2–5, and a-actinin as judged by qPCR, WB, and IFM analysis (Clement et al.,2009), supporting the conclusion that the primary cilium is critical in the regulation ofP19.CL6 cell differentiation into cardiomyocytes. This was further sustained by theobservation that knock down of IFT proteins reduced the number of clusters of beatingcardiomyocytes at day 12. Figure 3B presents data from a single and representativeexperiment in which 15 beating clusters in mock transfected cells were reduced to 3 and 1clusters in IFT88 and IFT88þ 20 siRNA-nucleofected cells, respectively. Moreover, theclusters of cardiomyocytes in IFT siRNA-nucleofected cells were abnormally small withno or very little networks between the individual clusters. It is likely that these irregularclusters of cardiomyocytes are formed as a consequence of a partial loss of plasmid DNAin IFT88 and IFT88þ 20 siRNA-nucleofected cells.

Early cardiogenesis is regulated by a number of different signal transduction pathways,including Hh, Wnt, bone morphogenetic protein (BMP), and PDGFR signaling (Hirata etal., 2007; Kwon et al., 2008; van Wijk et al., 2007; Washington Smoak et al., 2005). Inconjunction with defects in cardiomyogenesis qPCR analysis showed that IFT88þ 20siRNA inhibited Hh signaling in P19.CL6 cells. Under normal cardiomyocyte formationthe key elements in Hh signaling, Gli1 and Ptc-1, are upregulated throughout differentia-tion from day 1 to 9 (Clement et al., 2009). As shown in Fig. 3C, the relative mRNAlevels of Gli1 and Ptc-1 at day 5 were reduced to about 5 and 10%, respectively, of thelevel in mock transfected cells (Clement et al., 2009). These results indicate that Hhsignaling in P19.CL6 cell differentiation is coordinated by the primary cilium. Additionalexperiments will be required to investigate whether IFT88þ 20 siRNA and ciliaryknockdown in P19.CL6 cells also affects other signaling pathways, including Wnt,PDGFR, and BMP signaling, which may provide further insight into the function ofthe primary cilium in cardiomyogenesis and heart development.

VI. Summary

Wehavedescribed a detailed siRNA-basednucleofectionprotocol for examining the roleof the primary cilium in differentiation of P19.CL6 cancer stem cells into cardiomyocytes.Knockdown of IFT88 and IFT20 with their corresponding siRNA plasmid DNA inhibitsciliary formation, Hh signaling, and differentiation of cells into cardiomyocytes as judgedby qPCR, IFM, SDS-PAGE, and WB analysis. In addition to identifying cilia-relatedsignaling pathways, the nucleofection method can be extended to identify other genesthat are involved in P19.CL6 stem cell maintenance, proliferation, and differentiation.

AcknowledgmentsThis work was supported by the Lundbeck Foundation, the Danish Science Research Council (STC), the

Danish Heart Association (LAL), and funds from the Department of Biology, University of Copenhagen,

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Denmark (CAC). Wilhelm Johannsen Centre for Functional Genome Research is established by the DanishNational Research Foundation. The authors would like to thank Stine Gry Kristensen for excellent help onqPCR analysis. The authors would also like to thank Lillian Rasmussen, Kirsten Winther, and LauraSmedegaard Kruuse for technical assistance.

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