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.
Christian Alexandro Clement, PhD thesis 2009 2/45
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.
Christian Alexandro Clement, PhD thesis 2009 3/45
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
Christian Alexandro Clement, PhD thesis 2009 4/45
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
Christian Alexandro Clement, PhD thesis 2009 5/45
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.
Christian Alexandro Clement, PhD thesis 2009 6/45
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.
Christian Alexandro Clement, PhD thesis 2009 7/45
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.
Christian Alexandro Clement, PhD thesis 2009 8/45
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]).
Christian Alexandro Clement, PhD thesis 2009 9/45
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,
Christian Alexandro Clement, PhD thesis 2009 10/45
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).
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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”).
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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).
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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
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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
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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.
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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).
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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
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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
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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
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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
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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
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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).
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CHAPTER 3 – PRIMARY CILIA IN STEM CELL DIFFERENTIATION AND CARDIOGENESIS
3.1 INTRODUCTORY REMARKS
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
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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
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[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‐/‐
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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.
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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
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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.
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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.
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CHAPTER 4 – SENSORY CILIA IN THE PANCREAS AND REPRODUCTIVE ORGANS
4.1 INTRODUCTORY REMARKS
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
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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
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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).
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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.
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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 6 – REFERENCES
International Stem Cell Initiative, Adewumi O, Aflatoonian B, Ahrlund‐Richter L, Amit M, Andrews PW, Beighton G, Bello PA, Benvenisty N, Berry LS, Bevan S, Blum B, Brooking J, Chen KG, Choo AB, Churchill GA, Corbel M, Damjanov I, Draper JS, Dvorak P, Emanuelsson K, Fleck RA, Ford A, Gertow K, Gertsenstein M, Gokhale PJ, Hamilton RS, Hampl A, Healy LE, Hovatta O, Hyllner J, Imreh MP, Itskovitz‐Eldor J, Jackson J, Johnson JL, Jones M, Kee K, King BL, Knowles BB, Lako M, Lebrin F, Mallon BS, Manning D, Mayshar Y, McKay RD, Michalska AE, Mikkola M, Mileikovsky M, Minger SL, Moore HD, Mummery CL, Nagy A, Nakatsuji N, O'Brien CM, Oh SK, Olsson C, Otonkoski T, Park KY, Passier R, Patel H, Patel M, Pedersen R, Pera MF, Piekarczyk MS, Pera RA, Reubinoff BE, Robins AJ, Rossant J, Rugg‐Gunn P, Schulz TC, Semb H, Sherrer ES, Siemen H, Stacey GN, Stojkovic M, Suemori H, Szatkiewicz J, Turetsky T, Tuuri T, van den Brink S, Vintersten K, Vuoristo S, Ward D, Weaver TA, Young LA, Zhang W. (2007). Characterization of human embryonic stem cell lines by the International Stem Cell Initiative. Nat Biotechnol. Jul;25(7):803‐16. Epub 2007 Jun 17. Amit M, Carpenter MK, Inokuma MS, Chiu CP, Harris CP, Waknitz MA, Itskovitz‐Eldor J, Thomson JA. (2000). Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev. Biol. 227: 271–278. Apelqvist A, Ahlgren U, Edlund H. (1997). Sonic hedgehog directs specialised mesoderm differentiation in the intestine and pancreas. Curr. Biol. 7: 801–804 Ashizawa N, Endoh H, Hidaka K, Watanabe M, Fukumoto S. (1997). Three‐dimensional structure of the rat pancreatic duct in normal and inflammated pancreas. Microsc Res Tech 37:543–556. Atkin NB, Baker MC, Robinson R, Gaze SE. (1974). Chromosome studies on 14 near‐diploid carcinomas of the ovary. Eur. J. Cancer 10: 144–146. Aubin J, Dery U, Lemieux M, Chaliller P, Jeannotte L. (2002). Stomach regional specification requires Hoxa5‐driven mesenchymal‐epithelial signaling. Development 129: 4075–4087 Avilion AA, Nicolis SK, Pevny LH, Perez L, Vivian N, Lovell‐Badge R. (2003). Multipotent cell lineages in early mouse development depend on SOX2 function. Genes Dev. Jan 1;17(1):126‐40. Awan A, Oliveri RS, Jensen PL, Christensen ST, Andersen, CY. (2009). Chapter 11: Characterization of Human Embryonic Stem Cells (hESCs) grown under feeder‐free conditions. Human Embryonic Stem Cell Protocols. Methods in Molecular Biology (Ed. Turksen, K.), pp. 195‐210. Humana Press.
Bailey JM, Mohr AM, Hollingsworth MA. (2009). Sonic hedgehog paracrine signaling regulates metastasis and lymphangiogenesis in pancreatic cancer. Oncogene. Oct 8;28(40):3513‐25. Banizs B, Pike MM, Millican CL, Ferguson WB, Komlosi P, Sheetz J, Bell PD, Schwiebert EM, Yoder BK. (2005). Dysfunctional cilia lead to altered ependyma and choroid plexus function, and result in the formation of hydrocephalus. Development. 132(23):5329‐39. Bastida MF, Sheth R, Ros MA. (2009). A BMP‐Shh negative‐feedback loop restricts Shh expression during limb development. Development. 136(22):3779‐89. Basu B, Brueckner M. (2008). Cilia multifunctional organelles at the center of vertebrate left‐right asymmetry. Curr Top Dev Biol.;85:151‐74. Bellusci S, Furuta Y, Rush MG, Henderson R, Winnier G, Hogan BL. (1997). Involvement of Sonic hedgehog (Shh) in mouse embryonic lung growth and morphogenesis. Development. 124(1):53‐63. Berbari NF, Lewis JS, Bishop GA, Askwith CC, Mykytyn K. (2008). Bardet‐Biedl syndrome proteins are required for the localization of G protein‐coupled receptors to primary cilia. Proc Natl Acad Sci U S A 105, 4242‐4246. Berbari NF, O'Connor AK, Haycraft CJ, Yoder BK. (2009). The primary cilium as a complex signaling center. Curr Biol. Jul 14;19(13):R526‐35. Berman DM, Karhadkar SS, Maitra A, Montes De OR, Gerstenblith MR, Briggs K, Parker AR, Shimada Y, Eshleman JR, Watkins DN, Beachy PA. (2003). Widespread requirement for Hedgehog ligand stimulation in growth of digestive tract tumours. Nature;425:846–851. Bitgood MJ, McMahon AP. (1995). Hedgehog and BMP genes are coexpressed at many diverse sites of cell‐cell interaction in the mouse embryo. Dev. Biol. 172: 126–138. Boisvieux‐Ulrich E, Laine MC, Sandoz D. (1989). In vitro effects of taxol on ciliogenesis in quail oviduct. J. Cell Sci. 92, 9–20. Bongso A, Fong CY, Ng SC, Ratnam S. (1994). Isolation and culture of inner cell mass cells from human blastocysts. Hum. Reprod. 9: 2110–2117. Bradley A, Evans M, Kaufman MH, Robertson E. (1984). Formation of germ‐line chimaeras from embryo‐derived teratocarcinoma cell lines. Nature 309: 255–256.
Christian Alexandro Clement, PhD thesis 2009 36/45
Brand T. (2003). Heart development: molecular insights into cardiac specification and early morphogenesis. Dev Biol. 258(1):1‐19.
Bruneau BG. (2008). The developmental genetics of congenital heart disease. Nature. Feb 21;451(7181):943‐8. Breunig JJ, Sarkisian MR, Arellano JI, Morozov YM, Ayoub AE, Sojitra S, Wang B, Flavell RA, Rakic P, Town T. (2008). Primary cilia regulate hippocampal neurogenesis by mediating sonic hedgehog signaling. Proc Natl Acad Sci U S A. 105(35):13127‐32. Brokaw CJ, Kamiya R. (1987). Bending patterns of Chlamydomonas flagella: IV. Mutants with defects in inner and outer dynein arms indicate differences in dynein arm function. Cell Motil Cytoskeleton. 8(1):68‐75. Brown CB, Wenning JM, Lu MM, Epstein DJ, Meyers EN, Epstein JA. (2004). Cre‐mediated excision of Fgf8 in the Tbx1 expression domain reveals a critical role for Fgf8 in cardiovascular development in the mouse. Dev Biol. Mar 1;267(1):190‐202. Buckingham M, Meilhac S, Zaffran S. (2005). Building the mammalian heart from two sources of myocardial cells. Nature Rev. Genet. 6, 826–835. Campione‐Piccardo J, Sun J‐J, Craig J, McBurney MW. (1985). Cell‐cell interaction can influence drug‐induced differentiation of murine embryonal carcinoma cells. Dev. BioL, 109: 25‐31. Cano DA, Murcia NS, Pazour GJ, Hebrok M. (2004). Orpk mouse model of polycystic kidney disease reveals essential role of primary cilia in pancreatic tissue organization. Development 131:3457–3467. Cano DA, Hebrok M, Zenker M. (2007). Pancreatic development and disease. Gastroenterology 132:745–762. Cano DA, Sekine S, Hebrok M. (2006). Primary cilia deletion in pancreatic epithelial cells results in cyst formation and pancreatitis. Gastroenterology 131:1856–1869. Caspary T, Larkins CE, Anderson KV. (2007). The graded response to Sonic Hedgehog depends on cilia architecture. Dev Cell 12:767–778. Cathcart RS, Worthington WC. (1964). Ciliary movement in the rat cerebral ventricles: clearing action and directions of currents. J. Neuropath. Exp. Neurol. 23, 609–618.
Chandross KJ, Mezey, E. (2001). Plasticity of adult bone marrow stem cells. Mattson, M.P. and Van Zant, G. eds. (Greenwich, CT: JAI Press).
Chen MH, Wilson CW, Li YJ, Law KK, Lu CS, Gacayan R, Zhang X, Hui CC, Chuang PT. (2009). Cilium‐independent regulation of Gli protein function by Sufu in Hedgehog signaling is evolutionarily conserved. Genes Dev. 15;23(16):1910‐28. Chizhikov VV, Davenport J, Zhang Q, Shih EK, Cabello OA, Fuchs JL, Yoder BK, Millen KJ. (2007). Cilia proteins control cerebellar morphogenesis by promoting expansion of the granule progenitor pool. J Neurosci. 27(36):9780‐9.
Christensen ST, Guerra C, Wada Y, Valentin T, Angeletti RH, Satir P, Hamasaki T. (2001). A regulatory light chain of ciliary outer arm dynein in Tetrahymena thermophila. J Biol Chem. Jun 8;276(23):20048‐54.. Christensen ST, Ott CM. (2007). Cell signaling. A ciliary signaling switch. Science. 317(5836):330‐1. Christensen ST, Pedersen LB, Schneider L, Satir P. (2007). Sensory cilia and integration of signal transduction in human health and disease. Traffic. Feb;8(2):97‐109. Christensen ST, Pedersen SF, Satir P, Veland IR, Schneider L. (2008). Chapter 10 the primary cilium coordinates signaling pathways in cell cycle control and migration during development and tissue repair. Curr Top Dev Biol. 85:261‐301. Cooper AF, Yu KP, Brueckner M, Brailey LL, Johnson L, McGrath JM, Bale AE. (2005). Cardiac and CNS defects in a mouse with targeted disruption of suppressor of fused. Development 132: 4407–4417. Corbit KC, Aanstad P, Singla V, Norman AR, Stainier DY, Reiter JF. (2005). Vertebrate Smoothened functions at the primary cilium. Nature. 437(7061):1018‐21. Corbit KC, Shyer AE, Dowdle WE, Gaulden J, Singla V, Chen MH, Chuang PT, Reiter JF. (2008). Kif3a constrains beta‐catenin‐dependent Wnt signalling through dual ciliary and non‐ciliary mechanisms. Nat Cell Biol. 10(1):70‐6.
Davenport JR, Yoder BK. (2005). An incredible decade for the primary cilium: a look at a once‐forgotten organelle. Am J Physiol Renal Physiol. 289(6):F1159‐69.
DeHaan RL. (1965). Development of pacemaker tissue in the embryonic heart. Ann. NY. Acad.. Sct.. 127:7‐18
Doetschman T, Maeda N, Smithies O. (1988). Targeted mutation of the Hprt gene in mouse embryonic stem cells. Proc. Natl. Acad. Sci. USA 85, 8583‐8587.
Dunaeva M, Michelson P, Kogerman P, Toftgard R. (2003). Characterization of the physical interaction of Gli proteins with SUFU proteins. J Biol Chem 278: 5116–5122.
Edwards MKS, Harris JF, McBurney MW. (1983). Induced muscle differentiation in an embryonal carcinoma cell line. Mol. Cell. Biol., 3: 2280‐2286. Eggenschwiler JT, Anderson KV. (2007). Cilia and developmental signaling. Annu Rev Cell Dev Biol.;23:345‐73. Evans MJ, Kaufman MH. (1981). Establishment in culture of pluripotential cells from mouse embryos. Nature 292: 154–156. Flaherty MP, Dawn B. (2008). Noncanonical Wnt11 signaling and cardiomyogenic differentiation. Trends Cardiovasc Med. Oct;18(7):260‐8. Follit JA, Tuft RA, Fogarty KE, Pazour GJ. (2006). The intraflagellar transport protein Ift20 is associated with the golgi complex and is required for cilia assembly. Mol Biol of the Cell. 17:3781‐3792.
Christian Alexandro Clement, PhD thesis 2009 37/45
Follit JA, San Agustin JT, Xu F, Jonassen JA, Samtani R, Lo CW, Pazour GJ. (2008). The Golgin GMAP210/TRIP11 anchors Ift20 to the Golgi complex. PLoS Genet. 4(12):e1000315. Follit JA, Xu F, Keady BT, Pazour GJ. (2009). Characterization of mouse IFT complex B. Cell Motil Cytoskeleton. Aug;66(8):457‐68. Gerdes JM, Davis EE, Katsanis N. (2009). The vertebrate primary cilium in development, homeostasis, and disease. Cell 137, 32‐45. Gerdes JM, Katsanis N. (2008). Chapter 7: Ciliary function and wnt signal modulation. Curr Top Dev Biol. 85:175‐95. Gerdes JM, Liu Y, Zaghloul NA, Leitch CC, Lawson SS, et al. (2007). Disruption of the basal body compromises proteasomal function and perturbs intracellular Wnt response. Nat Genet 39: 1350–1360. Gianakopoulos PJ, Skerjanc IS. (2005). Hedgehog signaling induces cardiomyogenesis in P19 cells. J Biol Chem. 280(22):21022‐8. Gianakopoulos PJ, Skerjanc IS. (2009). Cross talk between hedgehog and bone morphogenetic proteins occurs during cardiomyogenesis in P19 cells. In Vitro Cell Dev Biol Anim. Oct;45(9):566‐72. Epub 2009 Jul 8. Gilula NB, Satir P. (1972). The ciliary necklace. A ciliary membrane specialization. J Cell Biol 53, 494‐509. Goddeeris MM, Schwartz R, Klingensmith J, Meyers EN. (2007). Independent requirements for Hedgehog signaling by both the anterior heart field and neural crest cells for outflow tract development. Development. 1593‐604. Gorivodsky M, Mukhopadhyay M, Wilsch‐Braeuninger M, Phillips M, Teufel A, Kim C, Malik N, Huttner W, Westphal H. (2008). Intraflagellar transport protein 172 is essential for primary cilia formation and plays a vital role in patterning the mammalian brain. Dev Biol. 325(1):24‐32. Gouttenoire J, Valcourt U, Bougault C, Aubert‐Foucher E, Arnaud E, Giraud L, Mallein‐Gerin F. (2007). Knockdown of the intraflagellar transport protein Ift46 stimulates selective gene expression in mouse chondrocytes and affects early development in zebrafish. J Biol Chem. 282(42):30960‐73. Grepin C, Dagnino L, Robitaille L, Haberstroh L, Antakly T, Nemer M. (1994). A hormone‐encoding gene identifies a pathway for cardiac but not skeletal muscle gene transcription. Mol Cell Biol. 14(5):3115‐29. Grépin C, Nemer G, Nemer M. (1997). Enhanced cardiogenesis in embryonic stem cells overexpressing the GATA‐4 transcription factor. Development. 124(12):2387‐95. Guillaume DJ, Zhang SC. (2008). Human embryonic stem cells: a potential source of transplantable neural progenitor cells. Neurosurg Focus.;24(3‐4):E3.
Habara‐Ohkubo A. (1996). Differentiation of beating cardiac muscle cells from a derivative of P19 embryonal carcinoma cells. Cell Struct Funct. 21(2):101‐10. Han YG, Kim HJ, Dlugosz AA, Ellison DW, Gilbertson RJ, Alvarez‐Buylla A. (2009). Dual and opposing roles of primary cilia in medulloblastoma development. Nat Med. Sep;15(9):1062‐5. Handel M, Schulz S, Stanarius A, Schreff M, Erdtmann‐Vourliotis M, Schmidt H, Wolf G, Hollt V. (1999). Selective targeting of somatostatin receptor 3 to neuronal cilia. Neuroscience 89, 909‐926. Haraguchi K, Hayashi T, Jimbo T, Yamamoto T, Akiyama T. (2006). Role of the kinesin‐2 family protein, KIF3, during mitosis. J. Biol. Chem. 281:4094–4099. Harris PC, Torres VE. (2008). Polycystic Kidney Disease. Annu Rev Med. Oct 23. Haycraft CJ, Swoboda P, Taulman PD, Thomas JH, Yoder BK. (2001). The C. elegans homolog of the murine cystic kidney disease gene Tg737 functions in a ciliogenic pathway and is disrupted in osm‐5 mutant worms. Development. 128(9):1493‐505. Haycraft CJ, Banizs B, Aydin‐Son Y, Zhang Q, Michaud EJ, Yoder BK. (2005). Gli2 and Gli3 localize to cilia and require the intraflagellar transport protein polaris for processing and function. PLoS Genet. 1(4):e53. Haycraft CJ, Zhang Q, Song B, Jackson WS, Detloff PJ, Serra R, Yoder BK. (2007). Intraflagellar transport is essential for endochondral bone formation. Development. 134(2):307‐16. Hebrok M, Kim SK, St Jacques B, McMahon AP, Melton DA. (2000). Regulation of pancreas development by Hedgehog signaling. Development 127: 4905–4913 Hebrok M, Kim SK, Melton DA. (1999). Screening for novel pancreatic genes expressed during embryogenesis. Diabetes. Aug;48(8):1550‐6.
Hildebrandt F, Otto E. (2005). Cilia and centrosomes: a unifying pathogenic concept for cystic kidney disease? Nat. Rev. Genet. 6, 928–940 Hildebrandt F, Zhou W. (2007). Nephronophthisis‐associated ciliopathies. J. Am. Soc. Nephrol. 18, 1855–1871
Hirata H, Kawamata S, Murakami Y, Inoue K, Nagahashi A, Tosaka M, Yoshimura N, Miyamoto Y, Iwasaki H, Asahara T, Sawa Y. (2007). Coexpression of platelet‐derived growth factor receptor alpha and fetal liver kinase 1 enhances cardiogenic potential in embryonic stem cell differentiation in vitro. J Biosci Bioeng. 103(5):412‐9. Hirokawa N. (1998). Kinesin and dynein superfamily proteins and the mechanism of organelle transport. Science;279: 519–526.
Christian Alexandro Clement, PhD thesis 2009 38/45
Hirokawa N, Tanaka Y, Okada Y, Takeda S. (2006). Nodal flow and the generation of left–right asymmetry. Cell 125, 33–45 Huang P, Schier AF. (2009). Dampened Hedgehog signaling but normal Wnt signaling in zebrafish without cilia. Development. Sep;136(18):3089‐98. Huangfu D, Anderson KV. (2006). Signaling from Smo to Ci/Gli: conservation and divergence of Hedgehog pathways from Drosophila to vertebrates. Development. 133(1):3‐14. Hutson MR, Kirby ML. (2007). Model systems for the study of heart development and disease. Cardiac neural crest and conotruncal malformations. Semin Cell Dev Biol. 18(1):101‐10. Hynes M, Stone DM, Dowd M, Pitts‐Meek S, Goddard A, Gurney A, Rosenthal A. (1997). Control of cell pattern in the neural tube by the zinc finger transcription factor and oncogene Gli‐1. Neuron. 19(1):15‐26. Ingham PW, McMahon AP. (2001). Hedgehog signaling in animal development: paradigms and principles. Genes Dev. 15, 3059–3087 Ivanova N, Dobrin R, Lu R, Kotenko I, Levorse J, DeCoste C, Schafer X, Lun Y, Lemischka IR. (2006). Dissecting self‐renewal in stem cells with RNA interference. Nature. Aug 3;442(7102):533‐8. Epub 2006 Jun 11. Jay PY, Harris BS, Maguire CT, Buerger A, Wakimoto H, Tanaka M, Kupershmidt S, Roden DM, Schultheiss TM, O'Brien TX, Gourdie RG, Berul CI, Izumo S. (2004). Nkx2‐5 mutation causes anatomic hypoplasia of the cardiac conduction system. J Clin Invest. 113(8):1130‐7. Jeffery PK, Reid L. (1975). New observations of rat airway epithelium: a quantitative and electron microscopic study. J Anat. Nov;120(Pt 2):295‐320. Jensen CG, Poole CA, McGlashan SR, Marko M, Issa ZI, Vujcich KV, Bowser SS. (2004). Ultrastructural, tomographic and confocal imaging of the chondrocyte primary cilium in situ. Cell Biol Int 28, 101‐110. Jia J, Kolterud A, Zeng H, Hoover A, Teglund S, Toftgård R, Liu A. (2009). Suppressor of Fused inhibits mammalian Hedgehog signaling in the absence of cilia. Dev Biol. Jun 15;330(2):452‐60. Johnson ET, Nicola T, Roarty K, Yoder BK, Haycraft CJ, Serra R. (2008). Role for primary cilia in the regulation of mouse ovarian function. Dev Dyn. 237(8):2053‐60. Johnson RL, Riddle RD, Laufer E, Tabin C. (1994). Sonic hedgehog: a key mediator of anterior‐posterior patterning of the limb and dorso‐ventral patterning of axial embryonic structures. Biochem Soc Trans. 22(3):569‐74. Jonassen JA, San Agustin J, Follit JA, Pazour GJ. (2008). Deletion of IFT20 in the mouse kidney causes misorientation of the mitotic spindle and cystic kidney disease. J Cell Biol. Nov 3;183(3):377‐84.
Kahan BW, Ephrussi B. (1970). Developmental potentialities of clonal in vitro cultures of mouse testicular teratoma. J. Natl. Cancer Inst. 44: 1015–1036. Kalderon D. (2005). The mechanism of hedgehog signal transduction. Biochem Soc Trans. 33(Pt 6):1509‐12. Kawahira H, Ma NH, Tzanakakis ES, McMahon AP, Chuang P‐T, Hebrok M. (2003). Combined activities of 650 J. Lau, H. Kawahira and M. Hebrok Hedgehog signaling in pancreas Hedgehog signaling inhibitors regulate pancreas development. Development 130: 4871–4879 Kawahira H, Scheel DW, Smith SB, German MS, Hebrok M. (2005). Hedgehog signaling regulates expansion of pancreatic epithelial cells. Dev Biol 280:111–121. Kelly RG, Brown NA, Buckingham ME. (2001).The arterial pole of the mouse heart forms from Fgf10‐expressing cells in pharyngeal mesoderm. Dev. Cell 1, 435–440. Kennedy KA, Porter T, Mehta V, Ryan SD, Price F, Peshdary V, Karamboulas C, Savage J, Drysdale TA, Li SC, Bennett SA, Skerjanc IS. (2009). Retinoic acid enhances skeletal muscle progenitor formation and bypasses inhibition by bone morphogenetic protein 4 but not dominant negative beta‐catenin. BMC Biol. Oct 8;7:67. Kennedy MP, Omran H, Leigh MW, Dell S, Morgan L, Molina PL, Robinson BV, Minnix SL, Olbrich H, Severin T, Ahrens P, Lange L, Morillas HN, Noone PG, Zariwala MA, Knowles MR. (2007). Congenital heart disease and other heterotaxic defects in a large cohort of patients with primary ciliary dyskinesia. Circulation. Jun 5;115(22):2814‐21. Kestler HA, Kuhl M. (2008). From individual Wnt pathways towards a Wnt signalling network. Philos Trans R Soc Lond B Biol Sci; 363: 1333–1347. Kim SK, Melton DA. (1998). Pancreas development is promoted by cyclopamine, a hedgehog signaling inhibitor. Proc. Natl. Acad. Sci. USA 95: 13036–13041. Kiprilov EN, Awan A, Desprat R, Velho M, Clement CA, Byskov AG, Andersen CY, Satir P, Bouhassira EE, Christensen ST, Hirsch RE. (2008). Human embryonic stem cells in culture possess primary cilia with hedgehog signaling machinery. J Cell Biol. 180(5):897‐904. Kleinsmith LJ, Pierce Jr GB. (1964). Multipotentiality of single embryonal carcinoma cells. Cancer Res. 24: 1544–1551. Klimanskaya I, Chung Y, Becker S, Lu SJ, Lanza R. (2006). Human embryonic stem cell lines derived from single blastomeres. Nature 444: 481–485. Knight MM, McGlashan SR, Garcia M, Jensen CG, Poole CA. (2009). Articular chondrocytes express connexin 43 hemichannels and P2 receptors ‐ a putative mechanoreceptor complex involving the primary cilium? J Anat 214, 275‐283.
Christian Alexandro Clement, PhD thesis 2009 39/45
Kodama T. (1983). A light and electron microscopic study on the pancreatic ductal system. Acta Pathol Jpn 33:297–321. Koorstra JB, Feldmann G, Habbe N, Maitra A. (2008). Morphogenesis of pancreatic cancer: role of pancreatic intraepithelial neoplasia (PanINs). Langenbecks Arch Surg. 393(4):561‐70. Kovacs JJ, Whalen EJ, Liu R, Xiao K, Kim J, Chen M, Wang J, Chen W, Lefkowitz RJ. (2008). β‐Arrestin‐mediated localization of smoothened to the primary cilium. Science;320: 1777–1781. Kozminski KG, Johnson KA, Forscher P, Rosenbaum JL. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc Natl Acad Sci USA 90, 5519‐5523. Kuehn EW, Walz G, Benzing T. (2007). Von hippel‐lindau: a tumor suppressor links microtubules to ciliogenesis and cancer development. Cancer Res 67, 4537‐4540. Kuo CT, Morrisey EE, Anandappa R, Sigrist K, Lu MM, Parmacek MS, Soudais C, Leiden JM. (1997). GATA4 transcription factor is required for ventral morphogenesis and heart tube formation. Genes Dev. 11(8):1048–60. Kwon C, Cordes KR, Srivastava D. (2008). Wnt/beta‐catenin signaling acts at multiple developmental stages to promote mammalian cardiogenesis. Cell Cycle. 7(24):3815‐8. Lau J, Kawahira H, Hebrok M. (2006). Hedgehog signaling in pancreas development and disease. Cell Mol Life Sci 63: 642–652. Laugwitz KL, Moretti A, Caron L, Nakano A, Chien KR. (2008). Islet1 cardiovascular progenitors: a single source for heart lineages? Development. 135(2):193‐205. Lavine KJ, Kovacs A, Ornitz DM. (2008). Hedgehog signaling is critical for maintenance of the adult coronary vasculature in mice. J Clin Invest. 118(7):2404‐14. Lehman JM, Michaud EJ, Schoeb TR, Aydin‐Son Y, Miller M, Yoder BK. (2008). The Oak Ridge Polycystic Kidney mouse: modeling ciliopathies of mice and men. Dev Dyn. 1960‐71.
Lensch MW. (2009). Cellular reprogramming and pluripotency induction. Br Med Bull. 90:19‐35. Lieber M, Mazzetta J, Nelson‐Rees W, Kaplan M, Todaro G. (1975). Establishment of a continuous tumor‐cell line (panc‐1) from a human carcinoma of the exocrine pancreas. Int J Cancer 15:741–747.
Lin G, Ou Yang Q, Zhou X, Gu Y, Yuan D, Li W, Liu G, Liu T, Lu G. (2007). A highly homozygous and parthenogenetic human embryonic stem cell line derived from a one‐pronuclear oocyte following in vitro fertilization procedure. Cell Res. 17: 999–1007. Lints TJ, Parsons LM, Hartley L, Lyons I, Harvey RP. (1993). Nkx2‐5: a novel murine homeobox gene expressed in early heart progenitor cells and their myogenic descendants. Development 119, 419‐431.
Liu A, Wang B, Niswander LA. (2005). Mouse intraflagellar transport proteins regulate both the activator and repressor functions of Gli transcription factors. Development. 132(13):3103‐11. Lucker BF, Behal RH, Qin H, Siron LC, Taggart WD, Rosenbaum JL, Cole DG. (2005). Characterization of the intraflagellar transport complex B core: direct interaction of the Ift81 and Ift74/72 subunits. J Biol Chem. 280(30):27688‐96. Lyons GE. (1994). In situ analysis of the cardiac muscle gene program during embryogenesis. Trends. Cardiovasc. Med. 4:70–77. Lyons I, Parsons LM, Hartley L, Li R, Andrews JE, Robb L, Harvey RP. (1995). Myogenic and morphogenetic defects in the heart tubes of murine embryos lacking the homeo box gene Nkx2–5. Genes Dev. 9:1654–1666. Mai Q, Yu Y, Li T, Wang L, Chen MJ, Huang SZ, Zhou C, Zhou Q. (2007). Derivation of human embryonic stem cell lines from parthenogenetic blastocysts. Cell Res. 17: 1008–1019. Mans DA, Voest EE, Giles RH. (2008). All along the watchtower: is the cilium a tumor suppressor organelle? Biochim Biophys Acta. 1786(2):114‐25. Martin GR. (1980). Teratocarcinomas and mammalian embryogenesis. Science, 209: 768‐776. Martin GR. (1981). Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinomas stem cells. Proc. Natl. Acad. Sci. 78: 7634–7638. Masui S, Nakatake Y, Toyooka Y, Shimosato D, Yagi R, Takahashi K, Okochi H, Okuda A, Matoba R, Sharov AA, Ko MS, Niwa H. (2007). Pluripotency governed by Sox2 via regulation of Oct3/4 expression in mouse embryonic stem cells. Nat Cell Biol. 9(6):625‐35. Masyuk AI, Masyuk TV, LaRusso NF. (2008). Cholangiocyte primary cilia in liver health and disease. Dev Dyn 237, 2007‐2012. McBurney MW, Jones‐Villeneuve EMV, Edwards MKS, Anderson PJ. (1982). Control of muscle and neuronal differentiation in a cultured embryonal carcinoma cell line. Nature, 299: 165‐167. McGrath J, Somlo S, Makova S, Tian X, Brueckner M. (2003). Two populations of node monocilia initiate left‐right asymmetry in the mouse. Cell 114:61–73. Meilhac SM, Esner M, Kelly RG, Nicolas JF, Buckingham ME. (2004). The clonal origin of myocardial cells in different regions of the embryonic mouse heart. Dev. Cell 6, 685–698. Mellgren AM, Smith CL, Olsen GS, Eskiocak B, Zhou B, Kazi MN, Ruiz FR, Pu WT, Tallquist MD. (2008). Platelet‐derived growth factor receptor beta signaling is required for efficient epicardial cell migration and development of two distinct coronary vascular smooth muscle cell populations. Circ Res. Dec 5;103(12):1393‐401.
Christian Alexandro Clement, PhD thesis 2009 40/45
Michaud EJ, Yoder BK. (2006). The primary cilium in cell signaling and cancer. Cancer Res 66, 6463‐6467. Mitsui K, Tokuzawa Y, Itoh H, Segawa K, Murakami M, Takahashi K, Maruyama M, Maeda M, Yamanaka S. (2003).The homeoprotein Nanog is required for maintenance of pluripotency in mouse epiblast and ES cells. Cell. May 30;113(5):631‐42. Miyamoto Y, Maitra A, Ghosh B, Zechner U, Argani P, Iacobuzio‐Donahue CA, Sriuranpong V, Iso T, Meszoely IM, Wolfe MS, Hruban RH, Ball DW, Schmid RM, Leach SD. (2003). Notch mediates TGF alpha‐induced changes in epithelial differentiation during pancreatic tumorigenesis. Cancer Cell 3:565–576. Mjaatvedt CH, Nakaoka T, Moreno‐Rodriguez R, Norris RA, Kern MJ, Eisenberg CA, Turner D, Markwald RR. (2001). The outflow tract of the heart is recruited from a novel heart‐forming field. Dev Biol. Oct 1;238(1):97‐109. Molkentin JD. (2000). The zinc finger‐containing transcription factors GATA‐4, ‐5, and ‐6. Ubiquitously expressed regulators of tissue‐specific gene expression. J. Biol. Chem. 275, 38949–38952. Molkentin JD, Antos C, Mercer B, Taigen T, Miano JM, Olson EN, (2000a). Direct activation of a GATA6 cardiac enhancer by Nkx2.5: evidence for a reinforcing regulatory network of Nkx2.5 and GATA transcription factors in the developing heart. Dev. Biol. 217, 301–309. Molkentin JD, Lin Q, Duncan SA, Olson EN. (1997). Requirement of the transcription factor GATA4 for heart tube formation and ventral morphogenesis. Genes Dev. 11, 1061–1072. Molkentin JD, Tymitz KM, Richardson JA, Olson EN, (2000b). Abnormalities of the genitourinary tract in female mice lacking GATA5. Mol. Cell. Biol. 20, 5256–5260. Morgan D, Eley L, Sayer J, Strachan T, Yates LM, Craighead AS, Goodship JA. (2002). Expression analyses and interaction with the anaphase promoting complex protein Apc2 suggest a role for inversin in primary cilia and involvement in the cell cycle. Hum Mol Genet 11: 3345–3350. Morin S, Charron F, Robitaille L, Nemer M. (2000). GATA‐dependent recruitment of MEF2 proteins to target promoters. EMBO J. 19(9):2046‐2055.
Moyer JH, Lee‐Tischler MJ, Kwon HY, Schrick JJ, Avner ED, Sweeney WE, Godfrey VL, Cacheiro NL, Wilkinson JE, Woychik RP. (1994). Candidate gene associated with a mutation causing recessive polycystic kidney disease in mice.Science. 264(5163):1329‐33.
Mulac‐Jericevic B, Mullinax RA, DeMayo FJ, Lydon JP, Conneely OM. (2000). Subgroup of reproductive functions of progesterone mediated by progesterone receptor‐B isoform. Science. 289(5485):1751‐4.
Murcia NS, Richards WG, Yoder BK, Mucenski ML, Dunlap JR, Woychik RP. (2000). The Oak Ridge Polycystic Kidney (orpk) disease gene is required for left‐right axis determination. Development. 127(11):2347‐55.
Nanot J, Le Douarin G. (1977). Ultrastructure of the embryonic mouse sinus node at the early stages of development. J Embryol Exp Morphol. Feb;37(1):133‐47. Narita N, Bielinska M, Wilson DB. (1997). Wild‐type endoderm abrogates the ventral developmental defects associated with GATA‐4 deficiency in the mouse. Dev. Biol. 189, 270–274. Nemer M. (2008). Genetic insights into normal and abnormal heart development. Cardiovasc Pathol. 17(1):48‐54. Neugebauer JM, Amack JD, Peterson AG, Bisgrove BW, Yost HJ. (2009). FGF signalling during embryo development regulates cilia length in diverse epithelia. Nature. Apr 2;458(7238):651‐4. Nichols J, Zevnik B, Anastassiadis K, Niwa H, Klewe‐Nebenius D, Chambers I, Schöler H, Smith A. (1998). Formation of pluripotent stem cells in the mammalian embryo depends on the POU transcription factor Oct4. Cell. Oct 30;95(3):379‐91. Nielsen SK, Møllgård K, Clement CA, Veland IR, Awan A, Yoder BK, Novak I, Christensen ST. (2008). Characterization of primary cilia and Hedgehog signaling during development of the human pancreas and in human pancreatic duct cancer cell lines. Dev Dyn. 237(8):2039‐52. Nilsson O, Reinius S. (1969). Light and electron microscopic structure of the oviduct. In the mammalian oviduct, edn 1, pp 57‐83. Eds ESE Hafez & RJ Blandau. Chicago and London: The University of Chicago press.
Niwa H, Miyazaki J, Smith AG. (2000). Quantitative expression of Oct‐3/4 defines differentiation, dedifferentiation or self‐renewal of ES cells. Nature Genet 24, 372–376.
Niwa H, Toyooka Y, Shimosato D, Strumpf D, Takahashi K, Yagi R, Rossant J. (2005). Interaction between Oct3/4 and Cdx2 determines trophectoderm differentiation. Cell. Dec 2;123(5):917‐29.
Niwa H. (2007). How is pluripotency determined and maintained? Development 134, 635–646.
Ocbina PJ, Tuson M, Anderson KV. (2009). Primary cilia are not required for normal canonical Wnt signaling in the mouse embryo. PLoS One. Aug 31;4(8):e6839.
O'Toole SA, Swarbrick A, Sutherland RL. (2009). The Hedgehog signalling pathway as a therapeutic target in early breast cancer development. Expert Opin Ther Targets. Sep;13(9):1095‐103. Pan J. (2008). Cilia and ciliopathies: from Chlamydomonas and beyond. Sci China C Life Sci. 51(6):479‐86.
Pan Y, Bai CB, Joyner AL, Wang B. (2006). Sonic hedgehog signaling regulates Gli2 transcriptional activity by suppressing its processing and degradation. Mol Cell Biol. 26(9):3365‐77.
Park OK, Mayo KE. (1991). Transient expression of progesterone receptor messenger RNA in ovarian granulosa cells after the preovulatory luteinizing hormone surge. Mol Endocrinol. Jul;5(7):967‐78.
Christian Alexandro Clement, PhD thesis 2009 41/45
Parker JF, Florell SR, Alexander A, DiSario JA, Shami PJ, Leachman SA. (2003). Pancreatic carcinoma surveillance in patients with familial melanoma. Arch Dermatol 139:1019–1025. Pazour GJ, Dickert BL, Vucica Y, Seeley ES, Rosenbaum JL, Witman GB, Cole DG. (2000). Chlamydomonas Ift88 and its mouse homologue, polycystic kidney disease gene tg737, are required for assembly of cilia and flagella. J Cell Biol. 151:709‐718. Pazour GJ, San Agustin JT, Follit JA, Rosenbaum JL, Witman GB. (2002). Polycystin‐2 localizes to kidney cilia and the ciliary level is elevated in orpk mice with polycystic kidney disease. Curr Biol. Jun 4;12(11):R378‐80. Pazour GJ. (2004). Intraflagellar transport and cilia‐dependent renal disease: the ciliary hypothesis of polycystic kidney disease. J. Am. Soc. Nephrol. 15, 2528–2536. Pedersen T, Peters H. (1968). Proposal for a classification of oocytes and follicles in the mouse ovary. J Reprod Fertil. Dec;17(3):555‐7. Pedersen LB, Rosenbaum JL. (2008). Intraflagellar transport (IFT) role in ciliary assembly, resorption and signalling. Curr Top Dev Biol.;85:23‐61. Pera MF, Reubinoff B, Trounson A. (2000). Human embryonic stem cells. J Cell Sci. Jan;113 ( Pt 1):5‐10. Pesce M, Scholer HR. (2001). Oct‐4: Gatekeeper in the beginnings of mammalian development. Stem Cells. Stem Cells.19(4):271‐8. Peters H, Byskov AG, Himelstein‐Braw R, Faber M. (1975). Follicular growth: the basic event in the mouse and human ovary. J Reprod Fertil. Dec;45(3):559‐66. Petropoulos H, Skerjanc IS. (2002). Beta‐catenin is essential and sufficient for skeletal myogenesis in P19 cells. J Biol Chem. May 3;277(18):15393‐9.
Plotnikova OV, Golemis EA, Pugacheva EN. (2008). Cell cycle‐dependent ciliogenesis and cancer. Cancer Res 68, 2058‐2061.
Praetorius H, Leipziger J. (2009). Released nucleotides amplify the cilium‐dependent, flow‐induced [Ca(2)] response in MDCK cells. Acta Physiol (Oxf).
Prasad NB, Biankin AV, Fukushima N, Maitra A, Dhara S, Elkahloun AG, Hruban RH, Goggins M, Leach SD. (2005). Gene expression profiles in pancreatic intraepithelial neoplasia reflect the effects of Hedgehog signaling on pancreatic ductal epithelial cells. Cancer Res;65:1619–1626.
Revazova ES, Turovets NA, Kochetkova OD, Kindarova LB, Kuzmichev LN, Janus JD, Pryzhkova MV. (2007). Patient‐specific stem cell lines derived from human parthenogenetic blastocysts. Cloning Stem Cells 9: 432–449.
Robertson EJ. (1986). Pluripotential stem cell lines as a route into the mouse germ line. Trends Genet. 2, 9‐13.
Rochais F, Mesbah K, Kelly RG. (2009). Signaling pathways controlling second heart field development. Circ Res. Apr 24;104(8):933‐42.
Rohatgi R, Milenkovic L, Scott MP. (2007). Patched1 regulates hedgehog signaling at the primary cilium. Science. 317(5836):372‐6.
Rohatgi R, Milenkovic L, Corcoran RB, Scott MP. (2009). Hedgehog signal transduction by Smoothened: pharmacologic evidence for a 2‐step activation process. Proc Natl Acad Sci U S A. Mar 3;106(9):3196‐201. Roose J, Clevers H. (1999). TCF transcription factors: molecular switches in carcinogenesis. Biochim Biophys Acta 1424:M23–M37. Rosenbaum JL, Witman GB. (2002). Intraflagellar transport. Nat Rev Mol Cell Biol 813‐825. Ross AJ, May‐Simera H, Eichers ER, Kai M, Hill J, Jagger DJ, Leitch CC, Chapple JP, Munro PM, Fisher S, Tan PL, Phillips HM, Leroux MR, Henderson DJ, Murdoch JN, Copp AJ, Eliot MM, Lupski JR, Kemp DT, Dollfus H, Tada M, Katsanis N, Forge A, Beales PL. (2005). Disruption of Bardet‐Biedl syndrome ciliary proteins perturbs planar cell polarity in vertebrates. Nat Genet. Oct;37(10):1135‐40. Ruel L, Thérond PP. (2009). Variations in Hedgehog signaling: divergence and perpetuation in Sufu regulation of Gli. Genes Dev. Aug 15;23(16):1843‐8. Ruiz I Altaba A. (1999). Gli proteins encode context‐dependent positive and negative functions: implications for development and disease. Development. 126(14):3205‐16. Rutter M, Wang J, Huang Z, Kuliszewski M, Post M. (2009). Gli2 Influences Proliferation in the Developing Lung through Regulation of Cyclin Expression. Am J Respir Cell Mol Biol. Jul 2. Sadowski SL. (2009). Congenital cardiac disease in the newborn infant: past, present, and future. Crit Care Nurs Clin North Am. 21(1):37‐48, vi. Satir P. (1998). Eur. J. Protistol. 34, 267–272.
Satir P, Christensen ST. (2007). Overview of structure and function of mammalian cilia. Annu Rev Physiol. 69:377‐400.
Satir P, Christensen ST. (2008). Structure and function of mammalian cilia. Histochem Cell Biol. Jun;129(6):687‐93.
Schneider, L., Clement, C. A., Teilmann, S. C., Pazour, G. J., Hoffmann, E. K., Satir, P., Christensen, S. T. (2005). PDGFRαα signaling is regulated through the primary cilium in fibroblasts. Curr Biol 15:1861–6.
Schneider L, Cammer M, Lehman J, Nielsen SK, Guerra CF, Veland, IR, Stock C, Hoffmann EK, Yoder BK, Schwab A, Satir P, Christensen ST (2009a) Directional cell migration and chemotaxis in wound healing response to PDGF‐AA are coordinated by the primary cilium in fibroblasts. Cell. Physiol Biochem. In press.
Christian Alexandro Clement, PhD thesis 2009 42/45
Schneider L, Stock C, Dieterich P, Satir P, Schwab A, Christensen ST, Pedersen SF. (2009b). The Na+/H+ exchanger NHE1 plays a central role in directional migration stimulated via PDGFRa in the primary cilium. J. Cell Biol. 185(1):163‐76.
Scholey JM. (2008). Intraflagellar transport motors in cilia: Moving along the cell’s antenna. J. Cell Biol. 180, 23–29.
Schoumacher RA, Ram J, Iannuzzi MC, Bradbury NA, Wallace RW, Tom Hon C, Kelly DR, Schmid SM, Gelder FB, Rado TA, Frizzell RA. (1990). A cystic fibrosis pancreatic adenocarcinoma cell line. Proc Natl Acad Sci U S A 87:4012–4016. Sedmera D, McQuinn T. (2008). Embryogenesis of the heart muscle. Heart Fail Clin. (3):235‐45. Seo S, Guo DF, Bugge K, Morgan DA, Rahmouni K, Sheffield VC. (2009). Requirement of Bardet‐Biedl syndrome proteins for leptin receptor signaling. Hum Mol Genet 18, 1323‐1331. Shah AS, Ben‐Shahar Y, Moninger TO, Kline JN, Welsh MJ. (2009). Motile cilia of human airway epithelia are chemosensory. Science. 325(5944):1131‐4.
Shimada M, Terada T. (2002). FSH and LH induce progesterone production and progesterone receptor synthesis in cumulus cells: a requirement for meiotic resumption in porcine oocytes. Mol Hum Reprod. Jul;8(7):612‐8.
Simpson F, Kerr MC, Wicking C. (2009). Trafficking, development and hedgehog. Mech Dev. 126(5‐6):279‐88. Singla V, Reiter JF. (2006). The primary cilium as the cell's antenna: signaling at a sensory organelle. Science. 313(5787):629‐33. Skerjanc I S, Petropoulos H, Ridgeway AG, Wilton S. (1998). Myocyte enhancer factor 2C and Nkx2‐5 up‐regulate each other's expression and initiate cardiomyogenesis in P19 cells. J. Biol. Chem. 273(52):34904‐34910
Slack JMW. (1995) Developmental biology of the pancreas. Development 121: 1569–1580 Slough J, Cooney L, Brueckner M. (2008). Monocilia in the embryonic mouse heart suggest a direct role for cilia in cardiac morphogenesis. Dev Dyn. 237:2304‐2314. Smith AG, Heath JK, Donaldson DD, Wong GG, Moreau J, Stahl M, Rogers D. (1988). Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature 336: 688–690. Smith AG. (2001). Embryo‐derived stem cells: of mice and men. Annu Rev Cell Dev Biol 17:435‐462 Song H, Yoon C, Kattman SJ, Dengler J, Massé S, Thavaratnam T, Gewarges M, Nanthakumar K, Rubart M, Keller GM, Radisic M, Zandstra PW. (2009). Regenerative Medicine Special Feature: Interrogating functional integration between injected pluripotent stem cell‐derived cells and surrogate cardiac tissue. Proc Natl Acad Sci U S A. Oct 21.
Spassky N, Han YG, Aguilar A, Strehl L, Besse L, Laclef C, Ros MR, Garcia‐Verdugo JM, Alvarez‐Buylla A. (2008). Primary cilia are required for cerebellar development and Shh‐dependent expansion of progenitor pool. Dev Biol. 317(1):246‐59. Srivastava D, Olson EN. (2000). A genetic blueprint for cardiac development. Nature. 407:221‐26 Srivastava D. (2006). Making or breaking the heart: from lineage determination to morphogenesis. Cell. 126:1037‐1048. Stanley EG, Biben C, Elefanty A, Barnett L, Koentgen F, Robb L, Harvey RP. (2002). Efficient Cre‐mediated deletion in cardiac progenitor cells conferred by a 3'UTR‐ires‐Cre allele of the homeobox gene Nkx2‐5. Int J Dev Biol. 46(4):431‐9. Stevens LC, Little CC. (1954). Spontaneous testicular teratomas in an inbred strain of mice. Proc. Natl. Acad. ci. 40: 1080–1087. Stojkovic M, Lako M, Stojkovic P, Stewart R, Przyborski S, Armstrong L, Evans J, Herbert M, Hyslop L, Ahmad S, Murdoch A, Strachan T. (2004). Derivation of human embryonic stem cells from day‐8 blastocysts recovered after three‐step in vitro culture. Stem Cells. 22(5):790‐7. Strelchenko N, Verlinsky O, Kukharenko V, Verlinsky Y. (2004). Morula‐derived human embryonic stem cells. Reprod. Biomed. Online 9: 623–629. Sucov HM. (1998). Molecular insights into cardiac development. Annu Rev Physiol. 60:287‐308. Supp DM, Brueckner M, Kuehn MR, Witte DP, Lowe LA, McGrath J, Corrales J, Potter SS. (1999). Targeted deletion of the ATP binding domain of left‐right dynein confirms its role in specifying development of left‐right asymmetries. Development. 126(23):5495‐504. Supp DM, Witte DP, Potter SS, Brueckner M. (1997). Mutation of an axonemal dynein affects left‐right asymmetry in inversus viscerum mice. Nature 389:963–966. Svard J, Heby‐Henricson K, Persson‐Lek M, Rozell B, Lauth M, Bergstrom A, Ericson J, Toftgard R, Teglund S. (2006). Genetic elimination of Suppressor of fused reveals an essential repressor function in the mammalian Hedgehog signaling pathway. Dev Cell 10: 187–197. Tanaka M, Chen Z, Bartunkova M, Yamazaki N, Izumo S. (1999). The cardiac homeobox gene Csx/Nkx2.5 lies genetically upstream of multiple genes essential for heart development. Development.;126:1269–1280. Tam PP, Rossant J. (2003). Mouse embryonic chimeras: tools for studying mammalian development. Development. 130(25):6155‐63. Tarkowski AK. (1961). Mouse chimaeras developed from fused eggs. Nature. Jun 3;190:857‐60. Taulman PD, Haycraft CJ, Balkovetz DF, Yoder BK. (2001). Polaris, a protein involved in left‐right axis patterning, localizes to basal bodies and cilia. Mol Biol Cell. 12(3):589‐99.
Christian Alexandro Clement, PhD thesis 2009 43/45
Teilmann SC, Byskov AG, Pedersen PA, Wheatley DN, Pazour GJ, Christensen ST. (2005). Localization of transient receptor potential ion channels in primary and motile cilia of the female murine reproductive organs. Mol Reprod Dev. 71(4):444‐52. Teilmann SC, Christensen ST. (2005). Localization of the angiopoietin receptors Tie‐1 and Tie‐2 on the primary cilia in the female reproductive organs. Cell Biol Int. May;29(5):340‐6. Teilmann SC, Clement CA, Thorup J, Byskov AG, Christensen ST. (2006). Expression and localization of the progesterone receptor in mouse and human reproductive organs. J Endocrinol. 191(3):525‐35. Teng J, Rai T, Tanaka Y, Takei Y, Nakata, T, Hirasawa M, Kulkarni AB, Hirokawa N. (2005). The KIF3 motor transports N‐cadherin and organizes the developing neuroepithelium. Nat. Cell Biol. 7, 474‐482. Thayer SP, di Magliano MP, Heiser PW, Nielsen CM, Roberts DJ, Lauwers GY, Qi YP, Gysin S, Fernandez‐del CC, Yajnik V, Antoniu B, McMahon M, Warshaw AL, Hebrok M. (2003). Hedgehog is an early and late mediator of pancreatic cancer tumorigenesis. Nature 425:851–856. Theunissen JW, de Sauvage FJ. (2009). Paracrine Hedgehog signaling in cancer. Cancer Res. 69(15):6007‐10. Thomas KR, Capecchi MR. (1987). Site‐directed mutagenesis by gene targeting in mouse embryo derived stem cells. Cell 61, 503‐512. Thomson JA, Kalishman J, Golos TG, Durning M, Harris CP, Becker RA, Hearn JP. (1995). Isolation of a primate embryonic stem cell line. Proc. Natl. Acad. Sci. 92: 7844–7848. Thomson JA, Kalishman J, Golos TG, Durning M, Harris CP, Hearn JP. (1996). Pluripotent cell lines derived from common marmoset (Callithrix jacchus) blastocysts. Biol. Reprod. 55: 254–259. Thomson JA, Marshall VS. (1998). Primate embryonic stem cells. Curr. Topics Dev. Biol. 38, 133‐165. Tsukui T, Capdevila J, Tamura K, Ruiz‐Lozano P, Rodriguez‐Esteban C, Yonei‐Tamura S, Magallón J, Chandraratna RA, Chien K, Blumberg B, Evans RM, Belmonte JC. (1999). Multiple left‐right asymmetry defects in Shh(‐/‐) mutant mice unveil a convergence of the shh and retinoic acid pathways in the control of Lefty‐1. Proc Natl Acad Sci U S A. 96(20):11376‐81. Tzahor E. (2007). Wnt/beta‐catenin signaling and cardiogenesis: timing does matter. Dev Cell. 13(1):10‐3. Uchida S, Fuke S, Tsukahara T. (2007). Upregulations of Gata4 and oxytocin receptor are important in cardiomyocyte differentiation processes of P19CL6 cells. J Cell Biochem. 100(3):629‐41. Ulloa F, Itasaki N, Briscoe J. (2007). Inhibitory Gli3 activity negatively regulates Wnt/beta‐catenin signaling. Curr Biol. Mar 20;17(6):545‐50.
van den Brink GR. (2007). Hedgehog signaling in development and homeostasis of the gastrointestinal tract. Physiol Rev 87:1343–1375. van der Heyden MA, Defize LH. Twenty one years of P19 cells: what an embryonal carcinoma cell line taught us about cardiomyocyte differentiation. Cardiovasc Res. (2003). May 1;58(2):292‐302. van Wijk B, Moorman AF, van den Hoff MJ. (2007). Role of bone morphogenetic proteins in cardiac differentiation. Cardiovasc Res. 74(2):244‐255. Veland IR, Awan A, Pedersen LB, Yoder BK, Christensen ST. (2009). Primary cilia and signaling pathways in mammalian development, health and disease. Nephron Physiol. 111(3):p39‐53. Epub 2009 Mar 10. Varjosalo M, Taipale J. (2008). Hedgehog: functions and mechanisms. Genes Dev; 22: 2454–2472. Wagner M, Siddiqui MA. (2007). Signal transduction in early heart development (II): ventricular chamber specification, trabeculation, and heart valve formation. Exp Biol Med (Maywood). 232(7):866‐80. Waldo KL, Kumiski DH, Wallis KT, Stadt HA, Hutson MR, Platt DH, Kirby ML. (2001). Conotruncal myocardium arises from a secondary heart field. Development. Aug;128(16):3179‐88. Washington Smoak I, Byrd NA, Abu‐Issa R, Goddeeris MM, Anderson R, Morris J, Yamamura K, Klingensmith J, Meyers EN. (2005). Sonic hedgehog is required for cardiac outflow tract and neural crest cell development. Dev Biol. 283(2):357‐72. Willert K, Nusse R. (1998). β‐Catenin: a key mediator of Wnt signaling. Curr Opin Genet Dev; 8: 95–102. Williams RL, Hilton DJ, Pease S, Willson TA, Stewart CL, Gearing DP, Wagner EF, Metcalf D, Nicola NA, Gough NM. (1988). Myeloid leukaemia inhibitory factor maintains the developmental potential of embryonic stem cells. Nature 336: 684–687. Wilson NF, Lefebvre PA. (2004). Regulation of flagellar assembly by glycogen synthase kinase 3 in Chlamydomonas reinhardtii . Eukaryot Cell; 3: 1307–1319. Wodarczyk C, Rowe I, Chiaravalli M, Pema M, Qian F, Boletta A. (2009). A novel mouse model reveals that polycystin‐1 deficiency in ependyma and choroid plexus results in dysfunctional cilia and hydrocephalus. PLoS One. 4(9):e7137. Wong SY, Reiter JF. (2008). Chapter 9 the primary cilium at the crossroads of Mammalian hedgehog signaling. Curr Top Dev Biol. 85:225‐60. Wong SY, Seol AD, So PL, Ermilov AN, Bichakjian CK, Epstein EH Jr, Dlugosz AA, Reiter JF. (2009). Primary cilia can both mediate and suppress Hedgehog pathway‐dependent tumorigenesis. Nat Med. 15(9):1055‐61.
Christian Alexandro Clement, PhD thesis 2009 44/45
Xu H, Morishima M, Wylie JN, Schwartz RJ, Bruneau BG, Lindsay EA, Baldini A. (2004). Tbx1 has a dual role in the morphogenesis of the cardiac outflow tract. Development. 131(13):3217‐27. Yamagishi T, Ando K, Nakamura H. (2009). Roles of TGFbeta and BMP during valvulo‐septal endocardial cushion formation. Anat Sci Int. 84(3):77‐87.. Yamagishi H, Yamagishi C, Nakagawa O, Harvey R, Olson E, Srivastava D. (2002). The combinatorial activities of Nkx2.5 and dHAND are essential for cardiac ventricle formation. Dev. Biol. 239, 190–203. Yamazaki H, Nakata T, Okada Y, Hirokawa N. (1995). KIF3A/B: a heterodimeric kinesin superfamily protein that works as a microtubule plus end‐directed motor for membrane organelle transport. J Cell Biol. 130(6):1387‐99. Yoder BK, Hou X, Guay‐Woodford LM. (2002a). The polycystic kidney disease proteins, polycystin‐1, polycystin‐2, polaris, and cystin, are co‐localized in renal cilia., J Am Soc Nephrol. 2002 Oct;13(10):2508‐16. Yoder BK, Tousson A, Millican L, Wu JH, Bugg CE. Jr Schafer JA, Balkovetz DF. (2002b). Polaris, a protein disrupted in orpk mutant mice, is required for assembly of renal cilium. Am J Physiol Renal Physiol. 282(3):F541‐52.
Yu J, Thomson JA. (2008). Pluripotent stem cell lines. Genes Dev. Aug 1;22(15):1987‐97. Zeng G, Germinaro M, Micsenyi A, Monga NK, Bell A, Sood A, Malhotra V, Sood N, Midda V, Monga DK, Kokkinakis DM, Monga SP. (2006). Aberrant Wnt/beta‐catenin signaling in pancreatic adenocarcinoma. Neoplasia 8:279–289. Zhang Q, Davenport JR, Croyle MJ, Haycraft CJ, Yoder BK. (2005). Disruption of IFT results in both exocrine and endocrine abnormalities in the pancreas of Tg737(orpk) mutant mice. Lab Invest 85: 45–64. Zhang XM, Ramalho‐Santos M, McMahon AP. (2001). Smoothened mutants reveal redundant roles for Shh and Ihh signaling including regulation of L/R symmetry by the mouse node. Cell. 106(2):781‐92. Zhu D, Shi S, Wang H, Liao K. (2009). Growth arrest induces primary cilium formation and sensitizes IGF‐1‐receptor signaling during differentiation induction of 3T3‐L1 preadipocytes. J Cell Sci. 122, 2760‐2768.
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CHAPTER 7 – ARTICLES [1‐5]
525
Expression and localization of the
progesterone receptor in mouseand human reproductive organsStefan 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
Journal of Endocrinology (2006) 191, 525–535
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
www.endocrinology-journals.org
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).
Journal of Endocrinology (2006) 191, 525–535
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.
S C TEILMANN and others . PR expression and localization528
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
Journal of Endocrinology (2006) 191, 525–535
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
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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.
PR expression and localization . S C TEILMANN and others 529
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S C TEILMANN and others . PR expression and localization530
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.
Journal of Endocrinology (2006) 191, 525–535
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
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PR expression and localization . S C TEILMANN and others 531
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
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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
S C TEILMANN and others . PR expression and localization532
Journal of Endocrinology (2006) 191, 525–535 www.endocrinology-journals.org
PR expression and localization . S C TEILMANN and others 533
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
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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.
Journal of Endocrinology (2006) 191, 525–535
S C TEILMANN and others . PR expression and localization534
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.
References
Alieva IB, Gorgidze LA, Komarova YA, Chernobelskaya OA & Vorobjev IA
1999 Experimental model for studying the primary cilia in tissue culture
cells. Membrane Cell Biology 12 895–905.
Andrade YN, Fernandes J, Vazquez E, Fernandez-Fernandez JM, Arniges M,
Sanchez TM, Villalon M & Valverde MA 2005 TRPV4 channel is involved
in the coupling of fluid viscosity changes to epithelial ciliary activity.
Journal of Cell Biology 14 869–874.
Asem EK, QinW&Rane SG 2002 Effect of basal lamina of ovarian follicle on
T- and L-type Ca(2C) currents in differentiated granulosa cells. American
Journal of Physiology, Endocrinology and Metabolism 282 E184–E196.
Brenner RM 1969 The biology of oviductal cilia. In The Mammalian Oviduct,
edn 1 , pp 203–229. Eds ESE Hafez & RJ Blandau. Chicago and London:
The University of Chicago Press.
Byskov AG 1974 Cell kinetic studies of follicular atresia in the mouse ovary.
Journal of Reproduction and Fertility 37 277–285.
Christensen ST, Guerra C, Wada Y, Valentin T, Angeletti RH, Satir P &
Hamasaki T 2001 A regulatory light chain of ciliary outer arm dynein in
Tetrahymena thermophila. Journal of Biological Chemistry 276 20048–20054.
Clemm DL, Sherman L, Boonyaratanakornkit V, Schrader WT, Weigel NL &
Edwards DP 2000 Differential hormone-dependent phosphorylation of
progesterone receptor A and B forms revealed by a phosphoserine site-
specific monoclonal antibody. Molecular Endocrinology 14 52–65.
Conneely OM, Mulac-Jericevic B, Lydon JP & De Mayo FJ 2001
Reproductive functions of the progesterone receptor isoforms: lessons from
knock-out mice. Molecular and Cellular Endocrinology 179 97–103.
Conneely OM, Mulac-Jericevic B & Lydon JP 2003 Progesterone-dependent
regulation of female reproductive activity by two distinct progesterone
receptor isoforms. Steroids 68 771–778.
Denner LA, Schrader WT, O’Malley BW & Weigel NL 1990 Hormonal
regulation and identification of chicken progesterone receptor phos-
phorylation sites. Journal of Biological Chemistry 265 16548–16555.
Fliegauf M, Olbrich H, Horvath J, Wildhaber JH, Zariwala MA, Kennedy M,
Knowles MR & Omran H 2005 Mislocalization of DNAH5 and DNAH9
in respiratory cells from patients with primary ciliary dyskinesia. American
Journal of Respiratory and Critical Care Medicine 171 1343–1349.
Fuentealba B, Nieto M & Croxatto HB 1987 Ovum transport in pregnant rats
is little affected by RU486 and exogenous progesterone as compared to
cycling rats. Biology of Reproduction 37 768–774.
Gava N, Clarke CL, Byth K, Arnett-Mansfield RL & DeFAZIO A 2004
Expression of progesterone receptors A and B in the mouse ovary during
the estrous cycle. Endocrinology 45 3487–3494.
Hild-Petito S, Stouffer RL & Brenner RM 1988 Immunocytochemical
localization of estradiol and progesterone receptors in the monkey ovary
throughout the menstrual cycle. Endocrinology 123 2896–2905.
Iwai T, Nanbu Y, Iwai M, Taii S, Fujii S & Mori T 1990 Immunohisto-
chemical localization of oestrogen receptors and progesterone receptors in
the human ovary throughout the menstrual cycle. Virchows Archiv. A,
Pathological Anatomy and Histopathology 417 369–375.
Journal of Endocrinology (2006) 191, 525–535
Iwai T, Fujii S, Nanbu Y, Nonogaki H, Konishi I, Mori T &Okamura H 1991
Effect of human chorionic gonadotropin on the expression of progesterone
receptors and estrogen receptors in rabbit ovarian granulosa cells and the
uterus. Endocrinology 129 1840–1848.
Jo M, Komar CM & Fortune JE 2002 Gonadotropin surge induces two
separate increases in messenger RNA for progesterone receptor in bovine
preovulatory follicles. Biology of Reproduction 67 1981–1988.
Lange CA 2004 Making sense of cross-talk between steroid hormone
receptors and intracellular signaling pathways: who will have the last word?
Molecular Endocrinology 18 269–278.
Luconi M, Bonaccorsi L, Bini L, Liberatori S, Pallini V, Forti G & Baldi E
2002 Characterization of membrane nongenomic receptors for pro-
gesterone in human spermatozoa. Steroids 67 505–509.
Lydon JP, DeMayo FJ, Funk CR, Mani SK, Hughes AR, Montgomery CA,
Jr., Shyamala G, Conneely OM & O’Malley BW 1995 Mice lacking
progesterone receptor exhibit pleiotropic reproductive abnormalities.Genes
& Development 9 2266–2278.
Lyons RA, Djahanbakhch O, Mahmood T, Saridogan E, Sattar S, Sheaff MT,
Naftalin AA & Chenoy R 2002 Fallopian tube ciliary beat frequency in
relation to the stage of menstrual cycle and anatomical site. Human
Reproduction 17 584–588.
Machelon V, Nome F, Grosse B & Lieberherr M 1996 Progesterone triggers
rapid transmembrane calcium influx and/or calcium mobilization from
endoplasmic reticulum, via a pertussis-insensitive G-protein in granulosa
cells in relation to luteinization process. Journal of Cellular Biochemistry 61
619–628.
Mahmood T, Saridogan E, Smutna S, Habib AM & Djahanbakhch O 1998
The effect of ovarian steroids on epithelial ciliary beat frequency in the
human Fallopian tube. Human Reproduction 13 2991–2994.
Mulac-Jericevic B, Mullinax RA, DeMayo FJ, Lydon JP & Conneely OM
2000 Subgroup of reproductive functions of progesterone mediated by
progesterone receptor-B isoform. Science 289 1751–1754.
Ohta Y, Sato T & Iguchi T 1993 Immunocytochemical localization of
progesterone receptor in the reproductive tract of adult female rats. Biology
of Reproduction 48 205–213.
Okada A, Ohta Y, Inoue S, Hiroi H, Muramatsu M & Iguchi T 2003
Expression of estrogen, progesterone and androgen receptors in the oviduct
of developing, cycling and pre-implantation rats. Journal of Molecular
Endocrinology 30 301–315.
Orihuela PA & Croxatto HB 2001 Acceleration of oviductal transport of
oocytes induced by estradiol in cycling rats is mediated by nongenomic
stimulation of protein phosphorylation in the oviduct. Biology of
Reproduction 65 1238–1245.
Orihuela PA, Rios M & Croxatto HB 2001 Disparate effects of estradiol on
egg transport and oviductal protein synthesis in mated and cyclic rats.
Biology of Reproduction 65 1232–1237.
Park OK & Mayo KE 1991 Transient expression of progesterone receptor
messenger RNA in ovarian granulosa cells after the preovulatory luteinizing
hormone surge. Molecular Endocrinology 5 967–978.
Pedersen T & Peters H 1968 Proposal for a classification of oocytes and
follicles in the mouse ovary. Journal of Reproduction and Fertility 17 555–557.
Peluso JJ 2004 Rapid actions of progesterone on granulosa cells. Steroids 69
579–583.
Peluso JJ, Fernandez G, Pappalardo A &White BA 2001 Characterization of a
putative membrane receptor for progesterone in rat granulosa cells. Biology
of Reproduction 65 94–101.
Peluso JJ, Bremner T, Fernandez G, Pappalardo A & White BA 2003
Expression pattern and role of a 60-kilodalton progesterone binding protein
in regulating granulosa cell apoptosis: involvement of the mitogen-activated
protein kinase cascade. Biology of Reproduction 68 122–128.
Poletti A, Conneely OM, McDonnell DP, Schrader WT, O’Malley BW &
Weigel NL 1993 Chicken progesterone receptor expressed in Saccharomyces
cerevisiae is correctly phosphorylated at all four Ser-Pro phosphorylation
sites. Biochemistry 32 9563–9569.
Qiu M & Lange CA 2003 MAP kinases couple multiple functions of
human progesterone receptors: degradation, transcriptional synergy, and
nuclear association. Journal of Steroid Biochemistry and Molecular Biology 85
147–157.
www.endocrinology-journals.org
PR expression and localization . S C TEILMANN and others 535
Robker RL, Russell DL, Espey LL, Lydon JP, O’Malley BW & Richards JS
2000 Progesterone-regulated genes in the ovulation process: ADAMTS-1
and cathepsin L proteases. PNAS 97 4689–4694.
Satir P & Christensen ST 2006. Overview of structure and function of
mammalian cilia. Annual Review of Physiology. In press.
Shao R, Markstrom E, Friberg PA, Johansson M & Billig H 2003 Expression
of progesterone receptor (PR) A and B isoforms in mouse granulosa cells:
stage-dependent PR-mediated regulation of apoptosis and cell prolifer-
ation. Biology of Reproduction 68 914–921.
Sheridan PL, Evans RM & Horwitz KB 1989 Phosphotryptic peptide
analysis of human progesterone receptor. New phosphorylated sites
formed in nuclei after hormone treatment. Journal of Biological Chemistry
264 6520–6528.
Slomczynska M, Krok M & Pierscinski A 2000 Localization of the
progesterone receptor in the porcine ovary. Acta Histochemica 102 183–191.
SuzukiT, SasanoH,KimuraN,TamuraM,FukayaT,YajimaA&NaguraH1994
Immunohistochemical distribution of progesterone, androgen and oestrogen
receptors in the human ovary during the menstrual cycle: relationship to
expression of steroidogenic enzymes.Human Reproduction 9 1589–1595.
Takimoto GS & Horwitz KB 1993 Progesterone receptor phosphorylation,
complexities in defining a functional role. Trends in Endocrinology and
Metabolism 4 1–7.
Teilmann SC & Christensen ST 2005 Ciliary localization of angiopoetin
receptors Tie-1 and Tie-2 in the female reproductive organs. Cell Biology
International 29 340–346.
www.endocrinology-journals.org
Teilmann SC, Byskov AG, Pedersen PA, Wheatley DN, Pazour GJ &
Christensen ST 2005 Localization of transient receptor potential ion
channels in primary and motile cilia of the female murine reproductive
organs. Molecular Reproduction and Development 71 444–452.
Vanderhyden BC & Macdonald EA 1998 Mouse oocytes regulate granulosa
cell steroidogenesis throughout follicular development. Biology of Repro-
duction 59 1296–1301.
VanSlyke JK & Musil LS 2001 Biochemical analysis of connexon assembly. In
Connexin Methods and Protocols, edn 1 , pp 117–134. Eds R Bruzzone &
C Giaume. Totowa, New Jersey: Humana Press Inc.
Verdugo P 1980 Ca2C-dependent hormonal stimulation of ciliary activity.
Nature 283 764–765.
Weigel NL 1996 Steroid hormone receptors and their regulation by
phosphorylation. Biochemical Journal 319 657–667.
Wessel T, Schuchter U & Walt H 2004 Ciliary motility in bovine oviducts for
sensing rapid non-genomic reactions upon exposure to progesterone.
Hormone and Metabolic Research 36 136–141.
Received in final form 22 August 2006Accepted 23 August 2006Made available online as an Accepted Preprint25 September 2006
Journal of Endocrinology (2006) 191, 525–535
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.
PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREAS 2043
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.
PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREAS 2045
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.
PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREAS 2047
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-
PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREAS 2049
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.).
REFERENCES
Apelqvist A, Ahlgren U, Edlund H. 1997.Sonic hedgehog directs specialised meso-derm differentiation in the intestine andpancreas. Curr Biol 7:801–804.
Ashizawa N, Endoh H, Hidaka K, Wa-tanabe M, Fukumoto S. 1997. Three-di-mensional structure of the rat pancreaticduct in normal and inflammated pan-creas. Microsc Res Tech 37:543–556.
Badano JL, Mitsuma N, Beales PL, Katsa-nis N. 2006. The ciliopathies: an emerg-ing class of human genetic disorders.Annu Rev Genomics Hum Genet 7:125–148.
Bak M, Hansen C, Tommerup N, LarsenLA. 2003. The Hedgehog signaling path-way—implications for drug targets incancer and neurodegenerative disorders.Pharmacogenomics 4:411–429.
Beachy PA, Karhadkar SS, Berman DM.2004. Tissue repair and stem cell renewalin carcinogenesis. Nature 432:324–331.
Berman DM, Karhadkar SS, Maitra A,Montes De Oca R, Gerstenblith MR,Briggs K, Parker AR, Shimada Y, Eshle-man JR, Watkins DN, Beachy PA. 2003.Widespread requirement for Hedgehog
ligand stimulation in growth of digestivetract tumours. Nature 425:846–851.
Buscher D, Ruther U. 1998. Expressionprofile of Gli family members and Shh innormal and mutant mouse limb develop-ment. Dev Dyn 211:88–96.
Buttitta L, Mo R, Hui CC, Fan CM. 2003.Interplays of Gli2 and Gli3 and their re-quirement in mediating Shh-dependentsclerotome induction. Development 130:6233–6243.
Cano DA, Murcia NS, Pazour GJ, HebrokM. 2004. Orpk mouse model of polycystickidney disease reveals essential role ofprimary cilia in pancreatic tissue organi-zation. Development 131:3457–3467.
Cano DA, Sekine S, Hebrok M. 2006. Pri-mary cilia deletion in pancreatic epithelialcells results in cyst formation and pancre-atitis. Gastroenterology 131:1856–1869.
Cano DA, Hebrok M, Zenker M. 2007. Pan-creatic development and disease. Gastro-enterology 132:745–762.
Caspary T, Larkins CE, Anderson KV.2007. The graded response to SonicHedgehog depends on cilia architecture.Dev Cell 12:767–778.
Chauvet V, Tian X, Husson H, Grimm DH,Wang T, Hieseberger T, Igarashi P, Ben-nett AM, Ibraghimov-Beskrovnaya O,Somlo S, Caplan MJ. 2004. Mechanicalstimuli induce cleavage and nucleartranslocation of the polycystin-1 C termi-nus. J Clin Invest 114:1433–1443.
Chen JK, Taipale J, Young KE, Maiti T,Beachy PA. 2002. Small molecule modu-lation of Smoothened activity. Proc NatlAcad Sci U S A 99:14071–14076.
Christensen ST, Ott CM. 2007. Cell signal-ing. A ciliary signaling switch. Science20;317:330–331.
Christensen ST, Guerra C, Wada Y, Valen-tin T, Angeletti RH, Satir P, HamasakiT. 2001. A regulatory light chain of cili-ary outer arm dynein in Tetrahymenathermophila. J Biol Chem 276:20048–20054.
Christensen ST, Pedersen LB, SchneiderL, Satir P. 2007. Sensory cilia and inte-gration of signal transduction in humanhealth and disease. Traffic 8:97–109.
Corbit KC, Aanstad P, Singla V, NormanAR, Stainier DY, Reiter JF. 2005. Verte-brate Smoothened functions at the pri-mary cilium. Nature 437:1018–1021.
Corbit KC, Shyer AE, Dowdle WE, Gaul-den J, Singla V, Reiter JF. 2008. Kif3aconstrains beta-catenin-dependent Wntsignalling through dual ciliary and non-ciliary mechanisms. Nat Cell Biol 10:70–76.
Davenport JR, Watts AJ, Roper VC, CroyleMJ, van Groen T, Wyss JM, Nagy TR,Kesterson RA, Yoder BK. 2007. Disrup-tion of intraflagellar transport in adultmice leads to obesity and slow-onset cys-tic kidney disease. Curr Biol 17:1586–1594.
Delmas P, Padilla F, Osorio N, Coste B,Raoux M, Crest M. 2004. Polycystins,calcium signaling, and human dis-eases. Biochem Biophys Res Commun322:1374 –1383.
2050 NIELSEN ET AL.
Eggenschwiler JT, Anderson KV. 2007.Cilia and developmental signaling. AnnuRev Cell Dev Biol 23:345–373.
Fliegauf M, Benzing T, Omran H. 2007.When cilia go bad: cilia defects and cil-iopathies. Nat Rev Mol Cell Biol 8:880–893.
Gerdes JM, Liu Y, Zaghloul NA, Leitch CC,Lawson SS, Kato M, Beachy PA, BealesPL, DeMartino GN, Fisher S, BadanoJL, Katsanis N. 2007. Disruption of thebasal body compromises proteasomalfunction and perturbs intracellular Wntresponse. Nat Genet 39:1350–1360.
Haycraft CJ, Banizs B, Aydin-Son Y,Zhang Q, Michaud EJ, Yoder BK. 2005.Gli2 and Gli3 localize to cilia and requirethe intraflagellar transport protein po-laris for processing and function. PLoSGenet 1:e53.
Hidaka K, Ashizawa N, Endoh H, Wa-tanabe M, Fukumoto S. 1995. Fine struc-ture of the cilia in the pancreatic duct ofWBN/Kob rat. Int J Pancreatol 18:207–213.
Hollande E, Salvador-Cartier C, Alvarez L,Fanjul M. 2005. Expression of a wild-type CFTR maintains the integrity of thebiosynthetic/secretory pathway in hu-man cystic fibrosis pancreatic duct cells.J Histochem Cytochem 53:1539–1552.
Huangfu D, Anderson KV. 2005. Cilia andHedgehog responsiveness in the mouse.Proc Natl Acad Sci U S A 102:11325–11330.
Huangfu D, Anderson KV. 2006. Signalingfrom Smo to Ci/Gli: conservation and di-vergence of Hedgehog pathways fromDrosophila to vertebrates. Development133:3–14.
Huangfu D, Liu A, Rakeman AS, MurciaNS, Niswander L, Anderson KV. 2003.Hedgehog signalling in the mouse re-quires intraflagellar transport proteins.Nature 426:83–87.
Jackerott M, Lee YC, Mollgard K, Kofod H,Jensen J, Rohleder S, Neubauer N,Gaarn LW, Lykke J, Dodge R, DalgaardLT, Sostrup B, Jensen DB, Thim L, NexoE, Thams P, Bisgaard HC, Nielsen JH.2006. Trefoil factors are expressed in hu-man and rat endocrine pancreas: differ-ential regulation by growth hormone.Endocrinology 147:5752–5759.
Kawahira H, Scheel DW, Smith SB, Ger-man MS, Hebrok M. 2005. Hedgehog sig-naling regulates expansion of pancreaticepithelial cells. Dev Biol 280:111–121.
Kayed H, Kleeff J, Esposito I, Giese T, Ke-leg S, Giese N, Buchler MW. 2003. Dis-tribution of Indian hedgehog and its re-ceptors patched and smoothened inhuman chronic pancreatitis. J Endorinol178:467–478.
Kayed H, Kleeff J, Osman T, Keleg S,Buchler MW, Friess H. 2006. Hedgehogsignaling in the normal and diseasedpancreas. Pancreas 32:119–129.
Kiprilov EK, Awan A, Desprat R, Velho M,Clement CA, Byskov AG, Andersen CY,Satir P, Bouhassira EE, Christensen ST,Hirsch RE. 2008. Human embryonicstem cells in culture possess primary
cilia with hedgehog signaling machinery.J Cell Biol 180:897–904.
Kodama T. 1983. A light and electron mi-croscopic study on the pancreatic ductalsystem. Acta Pathol Jpn 33:297–321.
Kottgen M. 2007. TRPP2 and autosomaldominant polycystic kidney disease. Bio-chim Biophys Acta 1772:836–850.
Koyama E, Young B, Nagayama M, Shi-bukawa Y, Enomoto-Iwamoto M,Iwamoto M, Maeda Y, Lanske B, Song B,Serra R, Pacifici M. 2007. ConditionalKif3a ablation causes abnormal hedge-hog signaling topography, growth platedysfunction, and excessive bone and car-tilage formation during mouse skeleto-genesis. Development 134:2159–2169.
Lau J, Kawahira H, Hebrok M. 2006.Hedgehog signaling in pancreas develop-ment and disease. Cell Mol Life Sci 63:642–652.
Lieber M, Mazzetta J, Nelson-Rees W,Kaplan M, Todaro G. 1975. Establish-ment of a continuous tumor-cell line(panc-1) from a human carcinoma of theexocrine pancreas. Int J Cancer 15:741–747.
Litingtung Y, Dahn RD, Li Y, Fallon JF,Chiang C. 2002. Shh and Gli3 are dis-pensable for limb skeleton formation butregulate digit number and identity. Na-ture 418:979–983.
Liu A, Wang B, Niswander LA. 2005a.Mouse intraflagellar transport proteinsregulate both the activator and repressorfunctions of Gli transcription factors. De-velopment. 132:3103–3111.
Liu W, Murcia NS, Duan Y, Weinbaum S,Yoder BK, Schwiebert E, Satlin LM.2005b. Mechanoregulation of intracellu-lar Ca2� concentration is attenuated incollecting duct of monocilium-impairedorpk mice. Am J Physiol Renal Physiol289:F978–F988.
Low SH, Vasanth S, Larson CH, Mukher-jee S, Sharma N, Kinter MT, Kane ME,Obara T, Weimbs T. 2006. Polycystin-1,STAT6, and P100 function in a pathwaythat transduces ciliary mechanosensa-tion and is activated in polycystic kidneydisease. Dev Cell 10:57–69.
Marigo V, Johnson RL, Vortkamp A, TabinCJ. 1996. Sonic hedgehog differentiallyregulates expression of GLI and GLI3during limb development. Dev Biol 180:273–283.
May SR, Ashique AM, Karlen M, Wang B,Shen Y, Zarbalis K, Reiter J, Ericson J,Peterson AS. 2005. Loss of the retro-grade motor for IFT disrupts localizationof Smo to cilia and prevents the expres-sion of both activator and repressor func-tions of Gli. Dev Biol 287:378–389.
Merkel CE, Karner CM, Carroll TJ. 2007.Molecular regulation of kidney develop-ment: is the answer blowing in the Wnt?.Pediatr Nephrol 22:1825–1838.
Morton JP, Mongeau ME, Klimstra DS,Morris JP, Lee YC, Kawaguchi Y, WrightVE, Hebrok M, Lewis BC. 2006. Sonichedgehog acts at multiple stages duringpancreatic tumorigenesis. Proc NatlAcad Sci U S A 104:5103–5108.
Nauli SM, Alenghat FJ, Luo Y, Williams E,Vassilev P, Li X, Elia AE, Lu W, BrownEM, Quinn SJ, Ingber DE, Zhou J. 2003.Polycystins 1 and 2 mediate mech-anosensation in the primary cilium ofkidney cells. Nat Genet 33:129–137.
Novak I, Hede SE, Hansen MR. 2008.Adenosine receptors in rat and humanpancreatic ducts stimulate chloridetransport. Pflugers Arch 456:437–447.
Odent S, Atti-Bitach T, Blayau M, MathieuM, Aug J, Delezo de AL, Gall JY, LeMarec B, Munnich A, David V, Veke-mans M. 1999. Expression of the Sonichedgehog (SHH) gene during early hu-man development and phenotypic ex-pression of new mutations causing holo-prosencephaly. Hum Mol Genet 8:1683–1689.
Ostrer H, Wilson DI, Hanley NA. 2006.Human embryo and early fetus research.Clin Genet 70:98–107.
Pan Y, Bai CB, Joyner AL, Wang B. 2006.Sonic hedgehog signaling regulates Gli2transcriptional activity by suppressingits processing and degradation. Mol CellBiol 26:3365–3377.
Pazour GJ, Dickert BL, Vucica Y, SeeleyES, Rosenbaum JL, Witman GB, ColeDG. 2000. Chlamydomonas IFT88 andits mouse homologue, polycystic kidneydisease gene tg737, are required for as-sembly of cilia and flagella. J Cell Biol151:709–718.
Pazour GJ, San Agustin JT, Follit JA,Rosenbaum JL, Witman GB. 2002. Poly-cystin-2 localizes to kidney cilia and theciliary level is elevated in orpk mice withpolycystic kidney disease. Curr Biol 12:R378–R380.
Pedersen LB, Schroder JM, Veland IR,Christensen ST. 2008. Assembly of pri-mary cilia. Dev Dyn 2008 [Epub ahead ofprint].
Praetorius HA, Spring KR. 2001. Bendingthe MDCK cell primary cilium increasesintracellular calcium. J Membr Biol 184:71–79.
Praetorius HA, Spring KR. 2003. The renalcell primary cilium functions as a flowsensor. Curr Opin Nephrol Hypertens12:517–520.
Rohatgi R, Milenkovic L, Scott MP. 2007.Patched1 regulates hedgehog signalingat the primary cilium. Science 317:372–376.
Rosenbaum JL, Witman GB. 2002. In-traflagellar transport. Nat Rev Mol CellBiol 3:813–825.
Ruiz-Perez VL, Blair HJ, Rodriguez-An-dres ME, Blanco MJ, Wilson A, Liu YN,Miles C, Peters H, Goodship JA. 2007.Evc is a positive mediator of Ihh-regu-lated bone growth that localises at thebase of chondrocyte cilia. Development134:2903–2912.
Satir P, Christensen ST. 2007. Overview ofstructure and function of mammaliancilia. Annu Rev Physiol 69:377–400.
Schneider L, Clement CA, Teilmann SC,Pazour GJ, Hoffmann EK, Satir P,Christensen ST. 2005. PDGFRalphaal-pha signaling is regulated through the
PRIMARY CILIA AND HEDGEHOG SIGNALING IN PANCREAS 2051
primary cilium in fibroblasts. Curr Biol15:1861–1866.
Schoumacher RA, Ram J, Iannuzzi MC,Bradbury NA, Wallace RW, Tom Hon C,Kelly DR, Schmid SM, Gelder FB, RadoTA, Frizzell RA. 1990. A cystic fibrosispancreatic adenocarcinoma cell line.Proc Natl Acad Sci U S A 87:4012–4016.
Schweitzer R, Vogan KJ, Tabin CJ. 2000.Similar expression and regulation ofGli2 and Gli3 in the chick limb bud.Mech Dev 98:171–174.
Simons M, Gloy J, Ganner A, BullerkotteA, Bashkurov M, Kronig C, Schermer B,Benzing T, Cabello OA, Jenny A,Mlodzik M, Polok B, Driever W, Obara T,Walz G. 2005. Inversin, the gene productmutated in nephronophthisis type II,functions as a molecular switch betweenWnt signaling pathways. Nat Genet 37:537–543.
Teilmann SC, Christensen ST. 2005. Local-ization of the angiopoietin receptorsTie-1 and Tie-2 on the primary cilia inthe female reproductive organs. Cell BiolInt 29:340–346.
Thayer SP, di Magliano MP, Heiser PW,Nielsen CM, Roberts DJ, Lauwers GY,Qi YP, Gysin S, Fernandez-del CastilloC, Yajnik V, Antoniu B, McMahon M,Warshaw AL, Hebrok M. 2003. Hedge-hog is an early and late mediator of pan-creatic cancer tumorigenesis. Nature425:851–856.
van den Brink GR. 2007. Hedgehog signal-ing in development and homeostasis ofthe gastrointestinal tract. Physiol Rev87:1343–1375.
Vierkotten J, Dildrop R, Peters T, Wang B,Ruther U. 2007. Ftm is a novel basalbody protein of cilia involved in Shh sig-nalling. Development 134:2569–2577.
Wang B, Fallon JF, Beachy PA. 2000.Hedgehog-regulated processing of Gli3produces an anterior/posterior repressorgradient in the developing vertebratelimb. Cell 100:423–434.
Weimbs T. 2007. Polycystic kidney diseaseand renal injury repair: common path-ways, fluid flow, and the function of poly-cystin-1. Am J Physiol Renal Physiol293:F1423–F1432.
Wilson ME, Scheel D, German MS. 2003.Gene expression cascades in pancreaticdevelopment. Mech Dev 120:65–80.
Yoder BK. 2007. Role of primary cilia inthe pathogenesis of polycystic kidneydisease. J Am Soc Nephrol 18:1381–1388.
Yoder BK, Tousson A, Millican L, Wu JH,Bugg CE Jr, Schafer JA, Balkovetz DF.2002a. Polaris, a protein disrupted inorpk mutant mice, is required for assem-bly of renal cilium. Am J Physiol RenalPhysiol 282:F541–F552.
Yoder BK, Hou X, Guay-Woodford LM.2002b. The polycystic kidney diseaseproteins, polycystin-1, polycystin-2, po-laris, and cystin, are co-localized in re-nal cilia. J Am Soc Nephrol 13:2508 –2516.
Zhang Q, Davenport JR, Croyle MJ, Hay-craft CJ, Yoder BK. 2005. Disruption ofIFT results in both exocrine and endo-crine abnormalities in the pancreas ofTg737(orpk) mutant mice. Lab Invest 85:45–64.
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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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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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901HUMAN EMBRYONIC STEM CELLS AND PRIMARY CILIA • KIPRILOV ET AL.
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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903HUMAN EMBRYONIC STEM CELLS AND PRIMARY CILIA • KIPRILOV ET AL.
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
References Badano , J.L. , N. Mitsuma , P.L. Beales , and N. Katsanis . 2006 . The ciliopathies: an
emerging class of human genetic disorders. Annu. Rev. Genomics Hum. Genet. 7 : 125 – 148 .
Blacque , O.E. , and M.R. Leroux . 2006 . Bardet-Biedl syndrome: an emerging pathomechanism of intracellular transport. Cell. Mol. Life Sci. 63 : 2145 – 2161 .
Bottenstein , J. 1985 . Growth and differentiation of neural cells in defi ned media. In Cell Cultures in the Neurosciences. J. Bottenstein and G. Sato, editors. Plenum Press, New York. 3 – 43 .
Chen , J.K. , J. Taipale , K.E. Young , T. Maiti , and P.A. Beachy . 2002 . Small molecule modulation of Smoothened activity. Proc. Natl. Acad. Sci. USA . 99 : 14071 – 14076 .
Christensen , S.T. , L.B. Pedersen , L. Schneider , and P. Satir . 2007 . Sensory cilia and integration of signal transduction in human health and disease. Traffi c . 8 : 97 – 109 .
Corbit , K.C. , P. Aansted , V. Singla , A.R. Norman , D.Y.R. Stanier , and J.F. Reiter . 2005 . Vertebrate Smoothened functions at the primary cilium . Nature . 437 : 1018 – 1021 .
Dubreuil , V. , A.M. Marzesco , D. Corbeil , W.B. Huttner , and M. Wilsch-Br ä uninger . 2007 . Midbody and primary cilium of neural progenitors re-lease extracellular membrane particles enriched in the stem cell marker prominin-1. J. Cell Biol. 176 : 483 – 495 .
Gerdes , J.M. , Y. Liu , N.A. Zaghloul , C.C. Leitch , S.S. Lawson , M. Kato , P.A. Beachy , P.L. Beales , G.N. DeMartino , S. Fisher , et al . 2007 . Disruption of the basal body compromises proteasomal function and perturbs intra-cellular Wnt response. Nat. Genet. 39 : 1350 – 1360 .
Hirokawa , N. , Y. Tanaka , Y. Okada , and S. Takeda . 2006 . Nodal fl ow and the generation of left-right asymmetry. Cell . 125 : 33 – 45 .
Haycraft , C.J. , B. Banzs , Y. Aydin-Son , Q. Zhang , E.J. Michaud , and B.K. Yoder . 2005 . Gli2 and Gli3 localize to cilia and require the intrafl agellar trans-port protein polaris for processing and function. PLoS Genet. 1 : e53 .
Huangfu , D. , and K.V. Anderson . 2005 . Cilia and Hedgehog responsiveness in the mouse. Proc. Natl. Acad. Sci. USA . 102 : 11325 – 11330 .
Laursen , S.B. , K. M ø llg å rd , C. Olesen , R.S. Oliveri , C.B. B ø chner , A.G. Byskov , A.N. Andersen , P.E. H ø yer , N. Tommerup , and C.Y. Andersen . 2007 . Regional differences in expression of specifi c markers for human embry-onic stem cells. Reprod. Biomed. Online . 15 : 89 – 98 .
Lipinski , R.J. , J.J. Gipp , J. Zhang , J.D. Doles , and W. Bushman . 2006 . Unique and complimentary activities of the Gli transcription factors in Hedgehog signaling. Exp. Cell Res. 312 : 1925 – 1938 .
Liu , A. , B. Wang , and L.A. Niswander . 2005 . Mouse intrafl agellar transport pro-teins regulate both the activator and repressor functions of Gli transcrip-tion factors. Development . 132 : 3103 – 3111 .
May , S.R. , A.M. Ashique , M. Karien , B. Wang , Y. Shen , K. Zarbilis , J. Reiter , J. Ericson , and A.S. Peterson . 2005 . Loss of retrograde motor for IFT dis-rupts localization of Smo to cilia and prevents the expression of both acti-vator and repressor functions of Gli. Dev. Biol. 287 : 378 – 389 .
Michaud , E.J. , and B.K. Yoder . 2006 . The primary cilium in cell signaling and cancer. Cancer Res. 66 : 6463 – 6467 .
Olivier , E.N. , C. Qiu , M. Velho , R.E. Hirsch , and E.E. Bouhassira . 2006 . Large-scale production of embryonic red blood cells from human embryonic stem cells. Exp. Hematol. 34 : 1635 – 1642 .
Orozco , J.T. , K.P. Wedaman , D. Signor , H. Brown , L. Rose , and J.M. Scholey . 1999 . Movement of motor and cargo along cilia. Nature . 398 : 674 .
Pan , G. , and J.A. Thomson . 2007 . Nanog and transcriptional networks in embry-onic stem cell pluripotency. Cell Res. 17 : 42 – 49 .
Pan , J. , and W. Snell . 2007 . The primary cilium: keeper of the key to cell division. Cell . 129 : 1255 – 1257 .
Pazour , G.J. 2004 . Intrafl agellar transport and cilia-dependent renal disease: the ciliary hypothesis of polycystic kidney disease. J. Am. Soc. Nephrol. 15 : 2528 – 2536 .
Rohatgi , R. , L. Milenkovic , and M. Scott . 2007 . Patched1 regulates Hedgehog signaling at the primary cilium. Science . 317 : 372 – 376 .
Rho , J.Y. , K. Yu , J.S. Han , J.I. Chae , D.B. Koo , H.S. Yoon , S.Y. Moon , K.K. Lee , and Y.M. Han . 2006 . Transcriptional profi ling of the developmentally im-portant signaling pathways in human embryonic stem cells. Hum. Reprod. 21 : 405 – 412 .
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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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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).
Journal of Cell Science 122 (17)
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
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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
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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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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.
ReferencesAngello, J. C., Kaestner, S., Welikson, R. E., Buskin, J. N. and Hauschka, S. D. (2007).
BMP induction of cardiogenesis in P19 cells requires prior cell-cell interaction(s). Dev.Dyn. 235, 2122-2133.
Awan, A., Oliveri, R. S., Jensen, P. L., Christensen, S. T. and Andersen, C. Y. (2009).
Characterization of human embryonic stem cells (hESCs) grown under feeder-free
conditions. Methods in Molecular Biology (ed. K. Turksen). Totowa, NJ: Humana Press.
Bellusci, S., Furuta, Y., Rush, M. G., Henderson, R., Winnier, G. and Hogan, B. L.
(1997). Involvement of Sonic hedgehog (Shh) in mouse embryonic lung growth and
morphogenesis. Development 124, 53-63.
Breunig, J. J., Sarkisian, M. R., Arellano, J. I., Morozov, Y. M., Ayoub, A. E., Sojitra,
S., Wang, B., Flavell, R. A., Rakic, P. and Town, T. (2008). Primary cilia regulate
hippocampal neurogenesis by mediating sonic hedgehog signaling. Proc. Natl. Acad.Sci. USA 105, 13127-13132.
Chen, J. K., Taipale, J., Young, K. E., Maiti, T. and Beachy, P. A. (2002). Small molecule
modulation of Smoothened activity. Proc. Natl. Acad. Sci. USA 99, 14071-14076.
Chizhikov, V. V., Davenport, J., Zhang, Q., Shih, E. K., Cabello, O. A., Fuchs, J. L.,
Yoder, B. K. and Millen, K. J. (2007). Cilia proteins control cerebellar morphogenesis
by promoting expansion of the granule progenitor pool. J. Neurosci. 27, 9780-9789.
Christensen, S. T. and Ott, C. M. (2007). Cell signaling: a ciliary signaling switch. Science317, 330-331.
Christensen, S. T., Pedersen, L. B., Schneider, L. and Satir, P. (2007) Sensory cilia and
integration of signal transduction in human health and disease. Traffic 8, 87-109.
Jour
nal o
f Cel
l Sci
ence
3081The primary cilium in cardiogenesis
Christensen, S. T., Pedersen, S. F., Satir, P., Veland, I. R. and Schneider, L. (2008).
Chapter 10 the primary cilium coordinates signaling pathways in cell cycle control and
migration during development and tissue repair. Curr. Top. Dev. Biol. 85, 261-301.
Corbit, K. C., Aanstad, P., Singla, V., Norman, A. R., Stainier, D. Y. and Reiter, J. F.
(2005). Vertebrate Smoothened functions at the primary cilium. Nature 437, 1018-1021.
Corbit, K. C., Shyer, A. E., Dowdle, W. E., Gaulden, J., Singla, V., Chen, M. H., Chuang,
P. T. and Reiter, J. F. (2008). Kif3a constrains beta-catenin-dependent Wnt signalling
through dual ciliary and non-ciliary mechanisms. Nat. Cell Biol. 10, 70-76.
Davenport, J. R. and Yoder, B. K. (2005). An incredible decade for the primary cilium:
a look at a once-forgotten organelle. Am. J. Physiol. Renal Physiol. 289, F1159-F1169.
Follit, J. A., Tuft, R. A., Fogarty, K. E. and Pazour, G. J. (2006). The intraflagellar
transport protein Ift20 is associated with the golgi complex and is required for cilia
assembly. Mol. Biol. Cell 17, 3781-3792.
Follit, J. A., San Agustin, J. T., Xu, F., Jonassen, J. A., Samtani, R., Lo, C. W. and
Pazour, G. J. (2008). The Golgin GMAP210/TRIP11 anchors Ift20 to the Golgi complex.
PLoS Genet. 4, e1000315.
Gerdes, J. M. and Katsanis, N. (2008). Chapter 7, Ciliary function and wnt signal
modulation. Curr. Top. Dev. Biol. 85, 175-195.
Gianakopoulos, P. J. and Skerjanc, I. S. (2005). Hedgehog signaling induces
cardiomyogenesis in P19 cells. J. Biol. Chem. 280, 21022-21028.
Goddeeris, M. M., Schwartz, R., Klingensmith, J. and Meyers, E. N. (2007). Independent
requirements for Hedgehog signaling by both the anterior heart field and neural crest
cells for outflow tract development. Development 134, 1593-1604.
Gorivodsky, M., Mukhopadhyay, M., Wilsch-Braeuninger, M., Phillips, M., Teufel,
A., Kim, C., Malik, N., Huttner, W. and Westphal, H. (2008). Intraflagellar transport
protein 172 is essential for primary cilia formation and plays a vital role in patterning
the mammalian brain. Dev. Biol. 325, 24-32.
Grépin, C., Dagnino, L., Robitaille, L., Haberstroh, L., Antakly, T. and Nemer, M.
(1994). A hormone-encoding gene identifies a pathway for cardiac but not skeletal muscle
gene transcription. Mol. Cell. Biol. 14, 3115-3129.
Grépin, C., Nemer, G. and Nemer, M. (1997). Enhanced cardiogenesis in embryonic stem
cells overexpressing the GATA-4 transcription factor. Development 124, 2387-2395.
Gouttenoire, J., Valcourt, U., Bougault, C., Aubert-Foucher, E., Arnaud, E., Giraud,
L. and Mallein-Gerin, F. (2007). Knockdown of the intraflagellar transport protein
Ift46 stimulates selective gene expression in mouse chondrocytes and affects early
development in zebrafish. J. Biol. Chem. 282, 30960-30973.
Habara-Ohkubo, A. (1996). Differentiation of beating cardiac muscle cells from a
derivative of P19 embryonal carcinoma cells. Cell Struct. Funct. 21, 101-110.
Han, Y. G., Spassky, N., Romanguera-Ros, M., Garcia-Verdugo, J. M., Aguilar, A.,
Schneider-Maunoury, S. and Alvarez-Buylla, A. (2008). Hedgehog signaling and
primary cilia are required for the formation of adult neural stem cells. Nat. Neurosci.11, 277-284.
Haraguchi, K., Hayashi, T., Jimbo, T., Yamamoto, T. and Akiyama, T. (2006). Role
of the kinesin-2 family protein, KIF3, during mitosis. J. Biol. Chem. 281, 4094-4099.
Harris, P. C. and Torres, V. E. (2009). Polycystic kidney disease. Annu. Rev. Med. 60,
321-327.
Haycraft, C. J., Swoboda, P., Taulman, P. D., Thomas, J. H. and Yoder, B. K. (2001).
The C. elegans homolog of the murine cystic kidney disease gene Tg737 functions in
a ciliogenic pathway and is disrupted in osm-5 mutant worms. Development 128, 1493-
1505.
Haycraft, C. J., Banizs, B., Aydin-Son, Y., Zhang, Q., Michaud, E. J. and Yoder, B.
K. (2005). Gli2 and Gli3 localize to cilia and require the intraflagellar transport protein
polaris for processing and function. PLoS Genet. 1, e53.
Haycraft, C. J., Zhang, Q., Song, B., Jackson, W. S., Detloff, P. J., Serra, R. and Yoder,
B. K. (2007). Intraflagellar transport is essential for endochondral bone formation.
Development 134, 307-316.
Hirata, H., Kawamata, S., Murakami, Y., Inoue, K., Nagahashi, A., Tosaka, M.,
Yoshimura, N., Miyamoto, Y., Iwasaki, H., Asahara, T. et al. (2007). Coexpression
of platelet-derived growth factor receptor alpha and fetal liver kinase 1 enhances
cardiogenic potential in embryonic stem cell differentiation in vitro. J. Biosci. Bioeng.103, 412-419.
Huangfu, D. and Anderson, K. V. (2005). Cilia and Hedgehog responsiveness in the mouse.
Proc. Natl. Acad. Sci. USA 102, 11325-11330.
Huangfu, D. and Anderson, K. V. (2006). Signaling from Smo to Ci/Gli: conservation
and divergence of Hedgehog pathways from Drosophila to vertebrates. Development133, 3-14.
Hynes, M., Stone, D. M., Dowd, M., Pitts-Meek, S., Goddard, A., Gurney, A. and
Rosenthal, A. (1997). Control of cell pattern in the neural tube by the zinc finger
transcription factor and oncogene Gli-1. Neuron 19, 15-26.
Iomini, C., Tejada, K., Mo, W., Vaananen, H. and Piperno, G. (2004). Primary cilia of
human endothelial cells disassemble under laminar shear stress. J. Cell Biol. 164, 811-
817.
Johnson, E. T., Nicola, T., Roarty, K., Yoder, B. K., Haycraft, C. J. and Serra, R.
(2008). Role for primary cilia in the regulation of mouse ovarian function. Dev. Dyn.237, 2053-2060.
Johnson, R. L., Riddle, R. D., Laufer, E. and Tabin, C. (1994). Sonic hedgehog: a key
mediator of anterior-posterior patterning of the limb and dorso-ventral patterning of axial
embryonic structures. Biochem. Soc. Trans. 22, 569-574.
Kiprilov, E. N., Awan, A., Desprat, R., Velho, M., Clement, C. A., Byskov, A. G.,
Andersen, C. Y., Satir, P., Bouhassira, E. E., Christensen, S. T. et al. (2008). Human
embryonic stem cells in culture possess primary cilia with hedgehog signaling machinery.
J. Cell Biol. 180, 897-904.
Kuo, C. T., Morrisey, E. E., Anandappa, R., Sigrist, K., Lu, M. M., Parmacek, M. S.,
Soudais, C. and Leiden, J. M. (1997). GATA4 transcription factor is required for ventral
morphogenesis and heart tube formation. Genes Dev. 11, 1048-1060.
Kwon, C., Cordes, K. R. and Srivastava, D. (2008). Wnt/beta-catenin signaling acts at
multiple developmental stages to promote mammalian cardiogenesis. Cell Cycle 7, 3815-
3818.
Lavine, K. J., Kovacs, A. and Ornitz, D. M. (2008). Hedgehog signaling is critical for
maintenance of the adult coronary vasculature in mice. J. Clin Invest. 118, 2404-2414.
Lehman, J. M., Michaud, E. J., Schoeb, T. R., Aydin-Son, Y., Miller, M. and Yoder,
B. K. (2008). The Oak Ridge Polycystic Kidney mouse: modeling ciliopathies of mice
and men. Dev. Dyn. 1960-1971.
Lints, T. J., Parsons, L. M., Hartley, L., Lyons, I. and Harvey, R. P. (1993). Nkx2-5: a
novel murine homeobox gene expressed in early heart progenitor cells and their myogenic
descendants. Development 119, 419-431.
Liu, A., Wang, B. and Niswander, L. A. (2005). Mouse intraflagellar transport proteins
regulate both the activator and repressor functions of Gli transcription factors.
Development 132, 3103-3111.
Lucker, B. F., Behal, R. H., Qin, H., Siron, L. C., Taggart, W. D., Rosenbaum, J. L.
and Cole, D. G. (2005). Characterization of the intraflagellar transport complex B core:
direct interaction of the Ift81 and Ift74/72 subunits. J. Biol. Chem. 280, 27688-27696.
Lyons, G. E. (1994). In situ analysis of the cardiac muscle gene program during
embryogenesis. Trends. Cardiovasc. Med. 4, 70-77.
Masui, S., Nakatake, Y., Toyooka, Y., Shimosato, D., Yagi, R., Takahashi, K., Okochi,
H., Okuda, A., Matoba, R., Sharov, A. A. et al. (2007). Pluripotency governed by
Sox2 via regulation of Oct3/4 expression in mouse embryonic stem cells. Nat. Cell Biol.9, 625-635.
May, S. R., Ashique, A. M., Karlen, M., Wang, B., Shen, Y., Zarbalis, K., Reiter, J.,
Ericson, J. and Peterson, A. S. (2005). Loss of the retrograde motor for IFT disrupts
localization of Smo to cilia and prevents the expression of both activator and repressor
functions of Gli. Dev. Biol. 287, 378-389.
Moyer, J. H., Lee-Tischler, M. J., Kwon, H. Y., Schrick, J. J., Avner, E. D., Sweeney,
W. E., Godfrey, V. L., Cacheiro, N. L., Wilkinson, J. E. and Woychik, R. P. (1994).
Candidate gene associated with a mutation causing recessive polycystic kidney disease
in mice. Science 264, 1329-1333.
Murcia, N. S., Richards, W. G., Yoder, B. K., Mucenski, M. L., Dunlap, J. R. and
Woychik, R. P. (2000). The Oak Ridge Polycystic Kidney (orpk) disease gene is required
for left-right axis determination. Development 127, 2347-2355.
Nauli, S. M., Kawanabe, Y., Kaminski, J. J., Pearce, W. J., Ingber, D. E. and Zhou,
J. (2008). Endothelial cilia are fluid shear sensors that regulate calcium signaling and
nitric oxide production through polycystin-1. Circulation 117, 1161-1171.
Nielsen, S. K., Møllgård, K., Clement, C. A., Veland, I. R., Awan, A., Yoder, B. K.,
Novak, I. and Christensen, S. T. (2008). Characterization of primary cilia and Hedgehog
signaling during development of the human pancreas and in human pancreatic duct cancer
cell lines. Dev. Dyn. 237, 2039-2052.
Nonaka, S., Tanaka, Y., Okada, Y., Takeda, S., Harada, A., Kanai, Y., Kido, M. and
Hirokawa, N. (1998). Randomization of left-right asymmetry due to loss of nodal cilia
generating leftward flow of extraembryonic fluid in mice lacking KIF3B motor protein.
Cell 95, 829-837.
Ocbina, P. J. and Anderson, K. V. (2008). Intraflagellar transport, cilia, and mammalian
Hedgehog signaling: analysis in mouse embryonic fibroblasts. Dev. Dyn. 237, 2030-
2038.
Pan, J. (2008). Cilia and ciliopathies: from Chlamydomonas and beyond. Sci. China CLife Sci. 51, 479-486.
Pan, Y., Bai, C. B., Joyner, A. L. and Wang, B. (2006). Sonic hedgehog signaling regulates
Gli2 transcriptional activity by suppressing its processing and degradation. Mol. Cell.Biol. 26, 3365-3377.
Paquin, J., Danalache, B. A., Jankowski, M., McCann, S. M. and Gutkowska, J. (2002).
Oxytocin induces differentiation of P19 embryonic stem cells to cardiomyocytes. Proc.Natl. Acad. Sci. USA 99, 9550-9555.
Pazour, G. J., Dickert, B. L., Vucica, Y., Seeley, E. S., Rosenbaum, J. L., Witman, G.
B. and Cole, D. G. (2000). Chlamydomonas Ift88 and its mouse homologue, polycystic
kidney disease gene tg737, are required for assembly of cilia and flagella. J. Cell Biol.151, 709-718.
Pedersen, L. B., Veland, I. R., Schrøder, J. M. and Christensen, S. T. (2008). Assembly
of primary cilia. Dev. Dyn. 237, 1993-2006.
Pesce, M. and Scholer, H. R. (2001). Oct-4: gatekeeper in the beginnings of mammalian
development. Stem Cells 19, 271-278.
Rohatgi, R., Milenkovic, L. and Scott, M. P. (2007). Patched1 regulates hedgehog signaling
at the primary cilium. Science 317, 372-376.
Rosenbaum, J. L. and Witman, G. B. (2002). Intraflagellar transport. Nat. Rev. Mol. Cell.Biol. 3, 813-825.
Ruiz I Altaba, A. (1999). Gli proteins encode context-dependent positive and negative
functions: implications for development and disease. Development 126, 3205-3216.
Satir, P. and Christensen, S. T. (2007). Overview of structure and function of mammalian
cilia. Annu. Rev. Physiol. 69, 377-400.
Schneider, L., Clement, C. A., Teilmann, S. C., Pazour, G. J., Hoffmann, E. K., Satir,
P. and Christensen, S. T. (2005). PDGFRaa signaling is regulated through the primary
cilium in fibroblasts. Curr. Biol. 15, 1861-1866.
Skerjanc, I. S. (1999). Cardiac and skeletal muscle development in P19 embryonal
carcinoma cells. Trends Cardiovasc. Med. 9, 139-143.
Slough, J., Cooney, L. and Brueckner, M. (2008). Monocilia in the embryonic mouse
heart suggest a direct role for cilia in cardiac morphogenesis. Dev. Dyn. 237, 2304-2314.
Jour
nal o
f Cel
l Sci
ence
Spassky, N., Han, Y. G., Aguilar, A., Strehl, L., Besse, L., Laclef, C., Ros, M. R., Garcia-Verdugo, J. M. and Alvarez-Buylla, A. (2008). Primary cilia are required for cerebellar
development and Shh-dependent expansion of progenitor pool. Dev. Biol. 317, 246-259.
Sucov, H. M. (1998). Molecular insights into cardiac development. Annu. Rev. Physiol.60, 287-308.
Takeda, S., Yonekawa, Y., Tanaka, Y., Okada, Y., Nonaka, S. and Hirokawa, N. (1999).
Left-right asymmetry and kinesin superfamily protein KIF3A: new insights in
determination of laterality and mesoderm induction by kif3A–/– mice analysis. J. CellBiol. 145, 825-836.
Takei, S., Ichikawa, H., Johkura, K., Mogi, A., No, H., Yoshie, S., Tomotsune, D. andSasaki, K. (2009). Bone morphogenetic protein-4 promotes induction of cardiomyocytes
from human embryonic stem cells in serum-based embryoid body development. Am. J.Physiol. Heart Circ. Physiol. 296, H1793-H1803.
Taulman, P. D., Haycraft, C. J., Balkovetz, D. F. and Yoder, B. K. (2001). Polaris, a
protein involved in left-right axis patterning, localizes to basal bodies and cilia. Mol.Biol. Cell 12, 589-599.
Teng, J., Rai, T., Tanaka, Y., Takei, Y., Nakata, T., Hirasawa, M., Kulkarni, A. B. andHirokawa, N. (2005). The KIF3 motor transports N-cadherin and organizes the
developing neuroepithelium. Nat. Cell Biol. 7, 474-482.
Tsukui, T., Capdevila, J., Tamura, K., Ruiz-Lozano, P., Rodriguez-Esteban, C., Yonei-Tamura, S., Magallón, J., Chandraratna, R. A., Chien, K., Blumberg, B. et al. (1999).
Multiple left-right asymmetry defects in Shh(–/–) mutant mice unveil a convergence of
the shh and retinoic acid pathways in the control of Lefty-1. Proc. Natl. Acad. Sci. USA96, 11376-11381.
Uchida, S., Fuke, S. and Tsukahara, T. (2007). Upregulations of Gata4 and oxytocin
receptor are important in cardiomyocyte differentiation processes of P19CL6 cells. J.Cell Biochem. 100, 629-641.
van Wijk, B., Moorman, A. F. and van den Hoff, M. J. (2007). Role of bone
morphogenetic proteins in cardiac differentiation. Cardiovasc. Res. 74, 244-255.
Washington Smoak, I., Byrd, N. A., Abu-Issa, R., Goddeeris, M. M., Anderson, R.,Morris, J., Yamamura, K., Klingensmith, J. and Meyers, E. N. (2005). Sonic hedgehog
is required for cardiac outflow tract and neural crest cell development. Dev. Biol. 283,
357-372.
Wong, S. Y. and Reiter, J. F. (2008). Chapter 9 the primary cilium at the crossroads of
Mammalian hedgehog signaling. Curr. Top. Dev. Biol. 85, 225-260.
Yamazaki, H., Nakata, T., Okada, Y. and Hirokawa, N. (1995). KIF3A/B: a heterodimeric
kinesin superfamily protein that works as a microtubule plus end-directed motor for
membrane organelle transport. J. Cell Biol. 130, 1387-1399.
Yoder, B. K., Tousson, A., Millican, L., Wu, J. H., Bugg, C. E., Jr, Schafer, J. A. andBalkovetz, D. F. (2002). Polaris, a protein disrupted in orpk mutant mice, is required
for assembly of renal cilium. Am. J. Physiol. Renal Physiol. 282, F541-F552.
Zhang, X. M., Ramalho-Santos, M. and McMahon, A. P. (2001). Smoothened mutants
reveal redundant roles for Shh and Ihh signaling including regulation of L/R symmetry
by the mouse node. Cell 106, 781-792.
Journal of Cell Science 122 (17)3082
Jour
nal o
f Cel
l Sci
ence
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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
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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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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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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
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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
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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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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.)
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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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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).
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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.
References
Bak, M., Hansen, C., Tommerup, N., and Larsen, L. A. (2003). The Hedgehog signaling pathway—Implications for drug targets in cancer and neurodegenerative disorders. Pharmacogenomics 4, 411–429.
Beachy, P. A., Karhadkar, S. S., and Berman, D. M. (2004). Tissue repair and stem cell renewal in carcinogenesis.Nature 432, 324–331.
Breunig, J. J., Sarkisian, M. R., Arellano, J. I., Morozov, Y. M., Ayoub, A. E., Sojitra, S., Wang, B., Flavell,R. A., Rakic, P., and Town, T. (2008). Primary cilia regulate hippocampal neurogenesis by mediating sonichedgehog signaling. Proc. Natl. Acad. Sci. USA 105, 13127–13132.
Clement, C. A., Kristensen, S. G., Møllgård, K., Pazour, G. J., Yoder, B. K., Larsen, L. A., and Christensen,S. T. ( AU182009). The primary cilium coordinates early cardiogenesis and hedgehog signaling in cardiomyocytedifferentiation. J. Cell Sci. in press.
Cole, D. G., and Snell, W. J. (2009). SnapShot: Intraflagellar transport. Cell 137, 784–784.Corbit, K. C., Aanstad, P., Singla, V., Norman, A. R., Stainier, D. Y., and Reiter, J. F. (2005). VertebrateSmoothened functions at the primary cilium. Nature 437, 1018–1021.
Gilula, N. B., and Satir, P. (1972). The ciliary necklace: A ciliary membrane specialization. J. Cell Biol. 53,494–509.
Gouttenoire, J., Valcourt, U., Bougault, C., Aubert-Foucher, E., Arnaud, E., Giraud, L., and Mallein-Gerin, F. (2007). Knockdown of the intraflagellar transport protein IFT46 stimulates selective geneexpression in mouse chondrocytes and affects early development in zebrafish. J. Biol. Chem. 282,30960–30973.
Gundersen, G. G., and Bulinski, J. C. (1986). Microtubule arrays in differentiated cells contain elevatedlevels of a post-translationally modified form of tubulin. Eur. J. Cell Biol. 42, 288–294.
Follit, J. A., Tuft, R. A., Fogarty, K. E., and Pazour, G. J. (2006). The intraflagellar transport protein IFT20 isassociated with the Golgi complex and is required for cilia assembly. Mol. Biol. Cell 17, 3781–3792.
Habara-Ohkubo, A. (1996). Differentiation of beating cardiac muscle cells from a derivative of P19embryonal carcinoma cells. Cell Struct. Funct. 21, 101–110.
Han, Y. G., Spassky, N., Romaguera-Ros, M., Garcia-Verdugo, J. M., Aguilar, A., Schneider-Maunoury, S.,and Alvarez-Buylla, A. (2008). Hedgehog signaling and primary cilia are required for the formation ofadult neural stem cells. Nat. Neurosci. 11, 277–284.
Haycraft, C. J., Zhang, Q., Song, B., Jackson, W. S., Detloff, P. J., Serra, R., and Yoder, B. K. (2007).Intraflagellar transport is essential for endochondral bone formation. Development 134, 307–316.
Hirata, H., Kawamata, S., Murakami, Y., Inoue, K., Nagahashi, A., Tosaka, M., Yoshimura, N., Miyamoto,Y., Iwasaki, H., Asahara, T., and Sawa, Y. (2007). Coexpression of platelet-derived growth factor receptoralpha and fetal liver kinase 1 enhances cardiogenic potential in embryonic stem cell differentiation in vitro.J. Biosci. Bioeng. 103, 412–419.
Huangfu, D., and Anderson, K. V. (2005). Cilia and hedgehog responsiveness in the mouse. Proc. Natl.Acad. Sci. USA 102, 11325–11330.
Huangfu, D., Liu, A., Rakeman, A. S., Murcia, N. S., Niswander, L., and Anderson, K. V. (2003). Hedgehogsignalling in the mouse requires intraflagellar transport proteins. Nature 426, 83–87.
Johnson, A. L., and AU19Woods, D. C. (2008). Dynamics of avian ovarian follicle development: Cellularmechanisms of granulosa cell differentiation. Gen Comp Endocrinol. Epub 2008 Nov 27.
Kiprilov, E. N., Awan, A., Desprat, R., Velho, M., Clement, C. A., Byskov, A. G., Andersen, C. Y., Satir, P.,Bouhassira, E. E., Christensen, S. T., and Hirsch, R. E. (2008). Human embryonic stem cells in culturepossess primary cilia with hedgehog signaling machinery. J. Cell Biol. 180, 897–904.
Komada, M., Saitsu, H., Kinboshi, M., Miura, T., Shiota, K., and Ishibashi, M. (2008). Hedgehog signaling isinvolved in development of the neocortex. Development 135, 2717–2727.
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Kwon, C., Cordes, K. R., and Srivastava, D. (2008). Wnt/beta-catenin signaling acts at multiple develop-mental stages to promote mammalian cardiogenesis. Cell Cycle 7, 3815–3818.
Lehman, J. M., Michaud, E. J., Schoeb, T. R., Aydin-Son, Y., Miller, M., and Yoder, B. K. (2008). The OakRidge Polycystic Kidney mouse: Modeling ciliopathies of mice and men. Dev. Dyn. 237, 1960–1971.
Liu, A., Wang, B., and Niswander, L. A. (2005). Mouse intraflagellar transport proteins regulate both theactivator and repressor functions of Gli transcription factors. Development 132, 3103–3111.
Nielsen, S. K., Møllgård, K., Clement, C. A., Veland, I. R., Awan, A., Yoder, B. K., Novak, I., andChristensen, S. T. (2008). Characterization of primary cilia and hedgehog signaling during developmentof the human pancreas and in human pancreatic duct cancer cell lines. Dev. Dyn. 237, 2039–2052.
Odent, S., Atti-Bitach, T., Blayau, M., Mathieu, M., Aug, J., Delezo de A. L., Gall, J. Y., Le Marec, B.,Munnich, A., David, V., and Vekemans, M. (1999). Expression of the Sonic hedgehog (SHH) gene duringearly human development and phenotypic expression of new mutations causing holoprosencephaly. Hum.Mol. Genet. 8, 1683–1689.
Ohazama, A., Haycraft, C. J., Seppala, M., Blackburn, J., Ghafoor, S., Cobourne, M., Martinelli, D. C., Fan,C. M., Peterkova, R., Lesot, H., Yoder, B. K., and Sharpe, P. T. (2009). Primary cilia regulate Shh activityin the control of molar tooth number. Development 136, 897–903.
Pedersen, L. B., and Rosenbaum, J. (2008). Intraflagellar transport (IFT) role in ciliary assembly, resorptionand signalling. Curr. Top. Dev. Biol. 85, 23–61.
Pedersen, L. B., Veland, I. R., Schrøder, J. M., and Christensen, S. T. (2008). Assembly of primary cilia. Dev.Dyn. 237, 1993–2006.
Piperno, G., and Fuller, M. T. (1985). Monoclonal antibodies specific for an acetylated form of alpha-tubulinrecognize the antigen in cilia and flagella from a variety of organisms. J. Cell Biol. 101, 2085–2094.
Rohatgi, R., Milenkovic, L., and Scott, M. P. ( AU202007). Patched1 regulates hedgehog signaling at the primarycilium. Science 317, 372–376.
Schneider, L., Clement, C. A., Teilmann, S. C., Pazour, G. J., Hoffmann, E. K., Satir, P., and Christensen,S. T. (2005). PDGFRaa signaling is regulated through the primary cilium in fibroblasts. Curr. Biol. 15,1861–1866.
Spassky, N., Han, Y. G., Aguilar, A., Strehl, L., Besse, L., Laclef, C., Ros, M. R., Garcia-Verdugo, J. M., andAlvarez-Buylla, A. (2008). Primary cilia are required for cerebellar development and Shh-dependentexpansion of progenitor pool. Dev. Biol. 317, 246–259.
Uchida, S., Fuke, S., and Tsukahara, T. (2007). Upregulations of Gata4 and oxytocin receptor are importantin cardiomyocyte differentiation processes of P19CL6 cells. J. Cell Biochem. 100, 629–641.
VanGuilder, H. D., Vrana, K. E., and Freeman, W. M. (2008). Twenty-five years of quantitative PCR for geneexpression analysis. Biotechniques 44, 619–626.
van Wijk, B., Moorman, A. F., and van den Hoff, M. J. (2007). Role of bone morphogenetic proteins incardiac differentiation. Cardiovasc. Res. 74, 244–255.
Veland, I. R., Awan, A., Pedersen, L. B., Yoder, B. K., and Christensen, S. T. (2009). Primary cilia andsignaling pathways in mammalian development, health and disease. Nephron. Physiol. 111, 39–53.
Washington Smoak, I., Byrd, N. A., Abu-Issa, R., Goddeeris, M. M., Anderson, R., Morris, J., Yamamura,K., Klingensmith, J., and Meyers, E. N. (2005). Sonic hedgehog is required for cardiac outflow tract andneural crest cell development. Dev. Biol. 283, 357–372.
Wong, S. Y., and Reiter, J. F. (2008). The primary cilium at the crossroads of mammalian hedgehog signaling.Curr. Top. Dev. Biol. 85, 225–260.
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Clement et al. Figure 1
Day 1 Day 12A
▪ ▪ ▪
▪ ▪Myh6
0.001
0.01
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▪ ▪Myh7
0.001
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Nkx2-5α-actininDAPI
Nkx2-5α-actininDAPI
glu-tbα-actininDAPI
glu-tbα-actininDAPI
Day 1 Day 12C
Clement et al. Figure 2
DAPIIft88 siRNAGFP
Ift20
β-actinm
ock
Ift20
siRN
Am
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Ift20
siRN
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Day 3 Day 6 Day 10
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