Shh/Gli Signaling in Anterior Pituitary

Home , Hypoprolactinemia, Posterior pituitary, Primordium, PROP1

SHH/GLI SIGNALING IN ANTERIOR PITUITARY

AND VENTRAL TELENCEPHALON DEVELOPMENT

YIWEI WANG

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Department of Genetics

CASE WESTERN RESERVE UNIVERSITY

January, 2011

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

______

candidate for the ______degree *.

(signed)______(chair of the committee)

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(date) ______

*We also certify that written approval has been obtained for any proprietary material contained therein.

TABLE OF CONTENTS

Table of Contents ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• i

List of Figures ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• v

List of Abbreviations •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• vii

Acknowledgements •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• ix

Abstract ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• x

Chapter 1 Background and Significance ••••••••••••••••••••••••••••••••••••••••••••••••• 1

1.1 Introduction to the pituitary gland ••••••••••••••••••••••••••••••••••••••••••••••••••••• 2

1.1.1 The pituitary gland and its function ••••••••••••••••••••••••••••••••••••••••••••• 3

1.1.2 The embryonic development of pituitary •••••••••••••••••••••••••••••••••••••• 6

1.2 Shh/Gli signaling in pituitary development •••••••••••••••••••••••••••••••••••••••• 13

1.2.1 The Sonic hedgehog signal transduction pathway •••••••••••••••••••••••• 13

1.2.2 The Gli transcription factors •••••••••••••••••••••••••••••••••••••••••••••••••••• 14

1.2.3 The role of Shh signaling in holoprosencephaly •••••••••••••••••••••••••••• 17

1.2.4 The role of Shh/Gli signaling in pituitary development ••••••••••••••••••••• 19

1.3 Other signals required in pituitary development •••••••••••••••••••••••••••••••••• 22

1.3.1 Bmps •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 22

1.3.2 Fgfs •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 25

1.3.3 Wnts ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 27

1.3.4 Signaling interactions in pituitary development ••••••••••••••••••••••••••••• 28

1.4 Shh/Gli signaling in ventral telencephalon development ••••••••••••••••••••••• 29

1.4.1 Introduction to the ventral telencephalon •••••••••••••••••••••••••••••••••••• 29

1.4.2 The embryonic development of the ventral telencephalon ••••••••••••••• 30

1.4.3 Shh signaling in telencephalon development ••••••••••••••••••••••••••••••• 33

Chapter 2 Shh/Gli signaling is active in the pituitary primordium and pituitary cell progenitors •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 35

2.1 Abstract •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 36

2.2 Materials and methods •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 37

2.2.1 Mouse breeding and genotyping •••••••••••••••••••••••••••••••••••••••••••••• 37

2.2.2 Fate mapping •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 37

2.2.3 X-gal staining and Immunohistochemistry ••••••••••••••••••••••••••••••••••• 38

2.3 Results ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 40

2.3.1 Shh signaling is active in the pituitary primordium ••••••••••••••••••••••••• 40

2.3.2 Gli expression gradient in developing Rathke’s pouch •••••••••••••••••••• 45

2.3.3 Hh-responding cells contribute to ventral cell types in the anterior pituitary •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 47

2.4 Discussion ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 49

2.5 Acknowlegements •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 51

Chapter 3 Direct and indirect requirements for Shh/Gli signaling in early pituitary development •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 52

3.1 Abstract •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 53

3.2 Introduction •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 54

3.3 Materials and methods •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 57

3.4 Results ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 60

3.4.1 Loss of Gli2 leads to a selective loss of specific pituitary cell types •••• 60

3.4.2 Progenitor proliferation in the nascent pituitary requires cell

autonomous expression of Gli2 ••••••••••••••••••••••••••••••••••••••••••••••••••••••• 63

3.4.3 Removal of Gli2 function before the closure of RP causes proliferation

defects in some pituitary progenitors ••••••••••••••••••••••••••••••••••••••••••••••••• 68

3.4.4 Overlapping roles of Gli genes in pituitary development •••••••••••••••••• 72

3.4.5 Activation of Hh signaling in RP increases pituitary proliferation

without affecting patterning •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 75

3.4.6 Patterning of pituitary requires diencephalic function of Gli2 ••••••••••••• 76

3.5 Discussion ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 78

3.6 Acknowledgements •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 82

Chapter 4 Patterning of the ventral telencephalon requires positive function of

Gli transcription factors •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 83

4.1 Abstract •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 84

4.2 Introduction •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 84

4.3 Materials and methods •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 87

4.4 Results ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 90

4.4.1 A subset of ventral telencephalic progenitors receives Hh signaling

and expresses Gli1-lacZ •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 90

4.4.2 Hh-responding progenitors produce progressively superficial cortical

interneurons ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 91

4.4.3 Gli activators are required for the specification of two progenitor

groups in the telencephalic sulcus •••••••••••••••••••••••••••••••••••••••••••••••••••• 95

4.4.4 Removal of all Gli genes disrupts production and proliferation of

ventral telencephalic neurons ••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 98

4.4.5 Intermingling of different neuronal groups in Gli2/3 mutants ••••••••••• 104

4.5 Discussion ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 108

4.6 Acknowledgements •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 115

Chapter 5 Conclusions and Future Directions ••••••••••••••••••••••••••••••••••••••• 116

5.1 Summary ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 117

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5.2 Early Shh signaling in ANR may specify the pituitary primordium •••••••••• 118

5.3 Two different phases of Shh/Gli signaling in pituitary proliferation •••••••••• 119

5.4 Gli2 may be required for diencephalon-derived signals BMP4 and Fgf8 •• 124

5.5 Mouse Gli2 mutant phenotypes correlate with human HPE •••••••••••••••••• 125

5.6 Future directions •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 127 5.6.1 Shh/Gli signaling is required for early specification of the pituitary primordium ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 127 5.6.2 Shh/Gli singaling is required in the ventral diencephalon for pituitary cell proliferation ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 131 5.6.3 Shh/Gli singaling is required for cell type specification of the anterior pituitary ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 131 5.6.4 Interactions between Shh and other signaling pathways •••••••••••••••• 132 5.6.4 a Shh interacts with Bmp2 to regulate pituitary differentiation ••••• 133 5.6.4 b Gli2 regulate Bmp2 transcription by direct binding sites ••••••••• 135 5.6.5 Further study of Shh signaling in the pituitary gland •••••••••••••••••••••• 137 Appendix •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 140 References ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 151

LIST OF FIGURES

Chapter 1 Figure 1-1 Location and structure of the pituitary gland in the human brain •••••• 4 Figure 1-2 Embryonic development of the pituitary gland in the mouse ••••••••••• 7 Figure 1-3 Transcription factors in pituitary development ••••••••••••••••••••••••••• 10 Figure 1-4 The Shh/Gli signaling pathway ••••••••••••••••••••••••••••••••••••••••••••• 15 Figure 1-5 Multiple signaling pathways required for pituitary development •••••• 24 Figure 1-6 Telencephalon development •••••••••••••••••••••••••••••••••••••••••••••••• 31 Chapter 2 Figure 2-1 Fate mapping strategy ••••••••••••••••••••••••••••••••••••••••••••••••••••••• 39 Figure 2-2 X-Gal staining of E8 Gli1lacZ embryos showed active Hh signaling in the anterior neural ridge •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 41 Figure 2-3 Hh-responding cells in the ANR contribute to Rathke’s pouch ••••••• 44 Figure 2-4 Gli activity can be detected in the ventral diencephalon and Rathke’s pouch •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 46 Figure 2-5 Hh-responding cells contribute to all five ventral cell types in the anterior pituitary ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 48 Chapter 3 Figure 3-1 Gli2 mutant pituitaries have normal patterning but show defects in proliferation ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 62 Figure 3-2 Production of specific cell types is affected in Gli2 mutants at E17.5 •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 64 Figure 3-3 Gli2 mutant cells do not proliferate as well as WT cells especially in the anterior wall of the pituitary, as revealed by mosaic analysis •••••••••••••••••• 67 Figure 3-4 Early loss of Gli2 function results in proliferation defects in pituitary progenitors ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 70 Figure 3-5 Early loss of Gli2 function results in the reduction of two groups of

pituitary cell types at E17.5 ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 73 Figure 3-6 Activation of Hh signaling after the closure of RP resulted in over proliferation of the pituitary •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 74 Figure 3-7 Gli2 is required for the expression of Bmp4 and Fgf8 in the ventral diencephalon ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 77 Chapter 4 Figure 4-1 X-gal staining of Gli1-lacZ and Gli2-lacZ on telencephalic sections of E12.5 mouse embryos •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 91 Figure 4-2 Different cortical interneurons were generated from progenitors responding to Hh signaling from ~E9.5 to ~E12.5 •••••••••••••••••••••••••••••••••••• 93 Figure 4-3 Gli activator is required for the development of subgroups of ventral telencephalic progenitors •••••••••••••••••••••••••••••••••••••••••••••••••••••••• 96 Figure 4-4 Gli2 and Gli3 have overlapping function in regulating Gli1 expression in the ventral telencephalon •••••••••••••••••••••••••••••••••••••••••••••• 100 Figure 4-5 Generation of ventral telencephalic neurons is affected in Gli2/3 double mutants •••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 101 Figure 4-6 The production of post-mitotic neurons is affected in Gli2/3 double mutant telencephalon ••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••••• 103 Figure 4-7 Dorsoventral patterning in different Gli mutants ••••••••••••••••••••••• 105 Figure 4-8 Schematic of Shh/Gli signaling in the ventral telencephalon •••••••• 111 Chapter 5 Figure 5-1 Shh is required both in the ventral diencephalon and the oral ectoderm for pituitary cell proliferation •••••••••••••••••••••••••••••••••••••••••••••••• 121 Figure 5-2 In vitro culture of the ANR explants •••••••••••••••••••••••••••••••••••••• 130

LIST OF ABBREVIATIONS

ACTH adrenocorticotropic hormone

ANR anterior neural ridge

BCC basal cell carcinoma

Bmp bone morphogenetic proteins

CGE caudal ganglionic eminences

CNS central nervous system

Co-Smads common-partner Smads

CPHD combined pituitary hormone deficiency

CRH corticotropin-releasing hormone

Fgf fibroblast growth factor

FSH follicle-stimulating hormone

Fu Fused

GH growth hormone

Hh Hedgehog

Hip Hedgehog interacting protein

HPE holoprosencephaly

INF infundibulum

I-Smads inhibitory Smads

LGE lateral ganglionic eminences

LH luteinizing hormone

MGE medial ganglionic eminences

MSH melanocyte-stimulating hormone

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POMC proopiomelanocortin

PRL prolactin

PKA Protein Kinase A

Ptc Patched

RP Rathke’s pouch

R-Smads receptor-regulated Smads rT rostral tip thyrotropes

RTK receptor tyrosine kinase

Shh Sonic Hedgehog

Smo Smoothened

SOD septo-optic dysplasia

Su(Fu) Supressor of Fused

TGF-β transforming growth factor-β

TSH thyroid-stimulating hormone

viii

Acknowledgements

First, I would like to thank my mentor, Brian Bai, for introducing me into

the scientific world and providing me with this great thesis project. Brian was discreet, dutiful and particularly patient while training me. I deeply appreciate his

guidance and support. I would not survive the last five years without Brian’s

tremendous help.

I am sincerely grateful to all of my committee members, Ron, Kathy and

Radhika. Ron, as my committee chair, is like a second mentor to me. He brought

essential theories into my project and always inspired me to think. Kathy gave

me great advice on my thesis work and made it feasible to pursue. Radhika

personally taught me a lot of writing skills and better way to present data. They

all did a large amount of extra work correcting my thesis and I cannot imagine

what would have happened if my committee did not devote so much time and

energy helping me with my graduation process. I am very lucky to have them as

my committee. I truly respect their professionalism and highly admire their

scientific attitude.

Finally, I would like to thank my family and friends, especially my mother.

You are always there for me, supporting me whatever my choice. I would not

hang in without your constant encouragement. I hope to share this achievement

with you.

Shh/Gli Signaling in Anterior Pituitary and Ventral Telencephalon

Abstract

YIWEI WANG

Signaling pathways that play a significant role during development have been studied as potential causes of human diseases. Sonic Hedgehog (Shh), one of the Hedgehog family members, was discovered in 1993 and named after

Sega's video game character “Sonic the Hedgehog”. As the best studied ligand of the hedgehog signaling pathway in mammals, Sonic Hedgehog was linked to a human brain disease “holoprosencephaly” in 1996. Since then, scientists endeavor to dissect the functions of Sonic Hedgehog in brain development.

Different roles have been described for Sonic Hedgehog during development, acting as a morphogen, mitogen or differentiation factor.

In my studies, I used the mouse model to explore the role of Shh/Gli in pituitary and forebrain development. Firstly, I examined the Gli activity during pituitary development. Shh/Gli signaling is active in the pituitary primordium and pituitary progenitor cells. Secondly, I used fate mapping strategy to trace Hh responding cells in the pituitary. Hh-responsive cells can contribute to multiple pituitary cell types. Thirdly, I examined multiple Gli mutants and found pituitary defects. Both Gli2 mutants and pituitary-specific Gli2 knockouts have proliferation defects in the pituitary. Taken together, Gli2 gene is required to mediate Shh signaling in pituitary cell proliferation.

Chapter 1 Background and Significance

Hedgehog (Hh) signaling is an important signaling pathway which has been studied in many organisms. Hedgehog is indispensible in regulating

patterning, proliferation, survival and growth in both embryos and adult

mammalian systems, especially in the central nervous system (CNS). A decade

ago, it was discovered that signaling via Sonic Hedgehog (Shh), a member of the

Hedgehog family, controls ventral patterning in the spinal cord. Since that time,

Shh has received much attention. It is reported that Shh signaling is required for

pituitary gland development in the mouse although its precise role is not known.

Shh signaling is also involved in the development of other CNS regions such as

the forebrain (telencephalon and diencepahlon). In my thesis work, I focus my

study on the roles of Shh signaling in pituitary primordium specification, cell

proliferation and cell type differentiation as well as telencephalon patterning. In

this chapter, I discuss the background relevant to these studies.

1.1 Introduction to the pituitary gland

The pituitary gland is an endocrine organ regulating growth, reproduction

and endocrine physiology. The pituitary gland is called the endocrine master of

our bodies because of its important and various functions in the endocrine

system. In this section, I introduce the composition and functions of the pituitary

gland and its embryonic development.

1.1.1 The pituitary gland and its function

The pituitary gland is a small, round endocrine organ attached to the hypothalamus and is situated within the sella turcica, a recess in the sphenoid bone at the base of the brain (Figure 1-1).

The pituitary gland is comprised of anterior pituitary, posterior pituitary and the intermediate lobe (Figure 1-1). The anterior pituitary (adenohypophysis) contains six different hormone-producing cell types: melanotropes, corticotropes, gonadotropes, thyrotropes, somatotropes and lactotropes. Melanotropes (M) secrete melanocyte-stimulating hormone (MSH) which stimulates the production and release of melanin in skin and hair. Corticotropes produce adrenocorticotropic hormone (ACTH) which stimulates the secretion of corticosteroids from the adrenal cortex. Gonadotropes secrete follicle-stimulating hormone (FSH) and luteinizing hormone (LH) which regulate gonadal function.

Thyrotropes secrete thyroid-stimulating hormone (TSH) which regulates thyroid gland size and thyroid hormone secretion. Somatotropes secrete growth hormone (GH), which regulates growth and metabolism. Lactotropes synthesize prolactin (PRL) which stimulates milk production. These hormones synergistically control physiological homeostasis both under basal and challenging conditions.

These critical cell types are generated from a common precursor tissue. Thus the anterior pituitary provides an excellent model to investigate cell type specification during organogenesis.

The posterior pituitary (neurohypophysis) contains the terminal axonal projections from the hypothalamus that signal from the hypothalamus to stimulate

Figure 1-1 Location and structure of the pituitary gland in the human brain

(Adapted and modified from Daniel, 1976 [1])

release of pituitary hormones. The posterior pituitary is the connection for hypothalamic-pituitary regulation of the vertebrate endocrine system.

Because the pituitary is the master regulator of the endocrine system, disorders (hypopituitarism or hyperpituitarism) and diseases (pituitary dysfunction and tumors) of the pituitary have a significant impact on human health. In particular, congenital pituitary disorders, in which the pituitary defects are already present at birth, have always attracted tremendous attention.

The congenital pituitary diseases can be classified into two categories according to the function of the genes involved [2]. The first category involves mutations in genes regulating the release or reception of pituitary hormones, like the GHRH receptor gene [3]. The second category involves mutations in genes regulating pituitary development at embryonic stages. Three pituitary disease genes of the latter type are POU1F1/Pit-1, PROP1 and HESX1/RPX, all of which belong to the homeobox class of transcription factors required for pituitary patterning and cell type differentiation. Mutations in the genes encoding the transcription factors Pit-1 and PROP1 have been reported as common causes of combined pituitary hormone deficiency (CPHD), and HESX1 mutations have been identified in children with familial septo-optic dysplasia (SOD), a syndromic form of congenital hypopituitarism involving optic nerve hypoplasia and agenesis of midline brain structures [4, 5] [6, 7]. However, mutation analysis of Pit-1,

PROP1 and HESX1 showed low frequency of mutations in children with sporadic forms of CHPD and SOD, suggesting that these genes can only account for rare cases of human congenital pituitary disorders [8].

1.1.2 Embryonic development of the pituitary

The pituitary gland in all vertebrates arises from two distinct embryonic tissues.

The anterior and the intermediate lobes of the pituitary derive from the oral, non- neural ectoderm while the posterior lobe originates from the neuroectoderm

(Figure 1-2). The pituitary gland arises from the mid-line of the anterior neural ridge (ANR), immediately anterior to the presumptive embryonic hypothalamus.

The anterior neural ridge is the junction between the anterior neural plate and anterior non-neural ectoderm, which can be seen at the rostral margin of the neural plate at E6.5 to E8 in mouse embryos. Fate map analyses in frog, chicken and mouse have shown that anterior neural ridge gives rise to both non-neural structures, including the anterior pituitary, and neural structures, including the posterior pituitary [9]. The neural origin of the adenohypophysis was confirmed in amphibian, Xenopus and rat [10-12]. Moreover, by use of surgical ablation in chick embryos, it has been demonstrated that the anterior neural ridge seems to be committed to form adenohypophyseal tissue as early as the 2- to 4-somitic stages [13]. In mouse, microinjections of dye into early embryos (3-10 somites stage) showed that both pituitary and hypothalamus derive from the anterior neural ridge [14]. These discoveries also indicate that before the first visible appearance of the pituitary primordium, certain cell compartments are already committed to develop into an anterior pituitary.

Figure 1-2 Embryonic development of the pituitary gland in the mouse

The two parts of the pituitary come from neural and nonneural tissues originating in the ANR between E6.5 to E8. At E9, the precursors of the anterior and posterior pituitary have separated into two adjacent domains within the midline of the ANR. When the oral ectoderm and ventral diencephalon come into contact, morphogenetic movements are initiated that will give rise to the mature 3D structure of the gland. The oral ectoderm invaginates to be Rathke’s pouch, which becomes the anterior pituitary at E17. The medial part of the ventral diencephalon evaginates to be infundibulum at E10.5 and grows into the posterior pituitary. (E6.5~E8:dorsal view of neural plate; E9~E17:sagittal view of enlarged pituitary. Adapted and modified from Zhu et al, 2007 [15])

In the mouse, the first sign of pituitary organogenesis occurs at around

E7.5 with the formation of the hypophyseal placode, which is a thickening of

ectoderm in the midline of the anterior neural ridge (reviewed in [16]). As the

anterior neural tube expands during development, the hypophyseal placode

displaces ventrally towards the roof of the future oral cavity. Then, the oral

ectoderm and neural ectoderm, which sit next to each other, come into contact at

E9 (Figure 1-2). The hypophyseal placode gives rise to Rathke’s pouch (RP).

Meanwhile, the neural ectoderm part evaginates to become the infundibulum

(INF), a portion of the ventral medial diencephalon. The infundibulum interacts with the oral ectoderm at about E10 and later becomes the posterior pituitary. By

E12.5, Rathke’s pouch has separated from the oral ectoderm.

Tissue interactions between the neural and oral ectoderm (non-neural part) are critical for the initial stages of pituitary specification. Several lines of evidence suggest that both pituitary induction and cell specification require signals derived from the ventral diencephalon (neural ectoderm). Mutation in T/ebp (Nkx2.1), which is expressed in the ventral diencephalon but not in Rathke’s pouch, causes

apoptosis and degeneration of the nascent pouch [17]. Mutation in Bmp4, which

is expressed in the ventral diencephalon, causes a loss of early RP [17, 18].

Nascent RP isolated from E10.5 mouse embryos will express the presumptive

pituitary marker Isl1 but not the definitive marker Lhx3 when cultured in vitro.

When combined with ventral diencephalic tissues, nascent RP can be induced to

express Lhx3 and other definitive markers [19].

The invagination of Rathke’s pouch brings the pituitary primordium in

close contact with the ventral diencephalon and prechordal plate/notochord.

Because of the close proximity between the nascent pituitary and prechordal

plate, it has been suggested that the formation of Rathke’s pouch and the

infundibulum precedes their commitment to the pituitary fate and may be induced

by morphogenetic signals from prechordal plate/notochord [20]. This raises the

possibility that notochord-derived signals, especially Shh, can affect adjacent

mesenchyme and the composition of extracellular matrix which in turn impacts

the migration of placode-derived cells [21].

Normal pituitary development is dependent upon a complex genetic

cascade of transcription factors either intrinsic or extrinsic to the developing

Rathke’s pouch. These factors dictate organ commitment, cell differentiation, and cell proliferation within the anterior pituitary.

The transcription factor Hesx1/HES-1/Rpx is a member of the paired-like class of homeodomain proteins and functions as a transcriptional repressor. It is

initially expressed in the anterior neural ridge midline and later becomes

restricted to Rathke’s pouch by E9.5~E13.5 [22]. Hesx1/HES-1/Rpx is important

for initial progression and proliferation of the pituitary gland, whereas its

subsequent down-regulation appears to be required for cell determination of

Prop1-dependent lineages [23-25].

Figure 1-3 Transcription factors in pituitary development

Transcription factors Hesx1, Pitx1/2, Pax6 and Isl1 are required for pituitary cell lineage determination. Lim3 is critical for further differentiation of Prop1/Pit1/GATA2 cell lineages

(somatotropes, lactotropes, thyrotropes and gonadotropes). Tbx19 is required for corticotropes and melanotropes. (M-melanotropes; C-corticotropes; S-somatotropes; L- lactotropes; T-thyrotropes; G-gonadotropes; rT-rostral tip thyrotropes. Adapted and modified from Scully and Rosenfeld, 2000 [16])

Two bicoid-related Pitx homeodomain factors, Pitx1 and Pitx2, display distinct but overlapping patterns of expression and exert roles in the development of the pituitary [26]. Targeted disruption of the Pitx1 gene leads to diminished expression of terminal differentiation markers for gonadotrope and thyrotrope

cells, and increased POMC gene expression [27]. In Pitx2-/- mice, the pituitary

gland fails to progress beyond E10.5 and has defects in early proliferation and

patterning [28].

P-LIM/Lim3/Lhx3 is expressed during early anterior pituitary development,

initially with strong uniform expression within Rathke’s pouch at E9.5 [29]. Lim3

encodes a LIM-homeodomain transcription factor that is required for specification

of somatotropes, lactotropes, ganadotropes and thyrotropes [29]. Deletion of the

Lhx3 gene in mice results in failure of the pituitary gland to grow and differentiate,

though Rathke’s pouch forms [29].

Isl1 is the first LIM protein to be expressed during pituitary development.

Between 10.5 and 11.5 dpc, Isl1 expression is restricted to the ventral portion of

the pouch, which becomes rostral tip thyrotropes after E12.5 [19]. Pax6 is

expressed in a dorsal-to-ventral gradient and is essential for establishing ventral-

dorsal cell boundaries in pituitary gland development [30].

As the Rathke’s pouch detaches from the underlying oral ectoderm at

E12/13, cells within the anterior pituitary proliferate and initiate differentiation into

specific lineages. Prop1 (Prophet of Pit1), which encodes a structurally related

paired-like homeodomain activator, is initially detected at E11, with robust

expression continuing until E13.5 to E14.5 [31]. Prop1 is required for initial determination and proliferation of Pit1 cell lineages (somatotrope, lactotrope, thyrotrope) in the anterior pituitary gland [23]. Expression of the ventral marker,

GATA2 (GATA binding protein 2) is induced by Bmp2, a potential morphogen that establishes a ventral to dorsal gradient within the nascent pituitary. GATA2 appears to be an important component of the gonadotrope and thyrotrope developmental programs [32]. GATA2 is normally expressed in pituitary cells located at the anterior tip of Rathke’s pouch, most of which have just exited the cell cycle [33]. At E13.5, Pit1 is expressed in prospective somatotrope, lactotrope and thyrotrope cells [34]. A T-box factor, T-pit, first identified in humans as Tbx19, has been shown to be selectively expressed in POMC-producing cells and to be capable of activating the POMC gene promoter [35]. Lim3-independent cells develop into corticotropes and melanotropes under the regulation of Tbx19 [35].

Under the regulation of a series of temporally and spatially restricted transcription factors, Rathke’s pouch develops into nascent anterior pituitary at

E17.5. The infundibulum of the ventral diencephalon develops into the posterior pituitary which links the anterior pituitary to the hypothalamus. The rest of the ventral diencephalic tissue gives rise to the dorsal thalamus, epithalamus, ventral thalamus, and hypothalamus.

1.2 Shh/Gli signaling in pituitary development

Shh is a member of the Hedgehog family of secreted glycoproteins that has been implicated in the development of many organs and cell groups. Downstream of

Shh signal transduction are Gli family of transcription factors. In this section, I discuss the role of Shh/Gli signaling in pituitary development.

1.2.1 Sonic hedgehog signal transduction pathway

The molecular mechanism, by which extracellular hedgehog signals are received, interpreted and transformed into intracellular cell fate decisions has been studied in both vertebrate and invertebrate species. In vertebrates, the functions of the protein encoded by the single invertebrate gene cubitus interruptus have been divided between the three Gli proteins. Similarly, there are three hedgehog homologues in mammals: Sonic Hedgehog, Indian Hedgehog and Desert Hedgehog.

The initial reception of hedgehog signals is controlled by two transmembrane proteins, the 12-transmembrane protein receptor Patched (Ptc) and the 7-transmembrane G-protein-coupled receptor Smoothened (Smo). In the absence of Shh, Ptc localized to cilia and inhibited Smo by preventing its accumulation within cilia, thereby inhibiting the downstream transduction cascade

[36]. On binding to Shh, Ptc left the cilia, leading to accumulation of Smo and activation of the Shh signaling cascade [36]. Hh signaling is mediated by a complex of cytoskeletal associated proteins including Suppressor of fused

(Su(Fu)), Fused (Fu), PKA, and the Gli proteins. Depending on the status of the pathway, Gli factors can be processed into repressor and activator. These activators and repressors are imported into the nucleus to regulate the expression of target genes (Figure 1-4). Hedgehog interacting protein (Hip) is a

Hedgehog inhibitor. Hip binds to the three mammalian hedgehog proteins and attenuates their bioactivities.

1.2.2 Gli transcription factors

Vertebrates have three distinct Gli genes, Gli1, Gli2 and Gli3, which belong to GLI-Kruppel family. Gli genes share homology with Drosophila gene Ci and different Gli genes have taken on the different roles of Ci. In the absence of

Hh ligand, Ci undergoes proteolytic cleavage to be a repressor and inhibits transcription of Hh target genes. In the presence of Hh, Ci full-length protein enters the nucleus to function as a transactivator [37].

Gli1 (GLI family zinc finger 1) contains a zinc finger domain but does not undergo proteolytic cleavage and has no repressor activity [38]. Gli1 functions as an Hh target gene activator and upregulation of Gli1 serves as readout of active

Hh signaling [39-41]. Transcriptional activation of Gli1 in response to Hh protein does not require protein synthesis but is dependent on the presence of functional

Gli2 and Gli3 protein [40]. Transgenic mice lacking Gli1 are viable and fertile [42].

Figure 1-4 The Shh/Gli signaling pathway

In the absence of Shh, Ptc prevents the accumulation of Smo. The activation of all Gli proteins is inhibited. In the presence of high levels of Shh ligand, Ptc inhibition is relieved;

Smo leads to the inhibition of Sufu and the activation of Gli2- and Gli3-mediated genes.

Gli activators and repressors are translocated into the nuclei to target downstream

genes. (Shh-Sonic Hedgehog; Ptc-Patched; Smo-Smoothened; Su(Fu)-Suppressor of

Fused; PKA-Protein Kinase A; Hip-Hedgehog interacting protein. Adapted and modified

from McMahon, 2000 [43])

Gli2 and Gli3 (GLI family zinc finger 2/3) share a high degree of sequence

similarity, including phosphorylation sites, activator and repressor domains. Gli2

functions as an important transactivator of Hh signaling, as revealed by biochemical analysis and in vivo mouse knock-in genetic analysis [44, 45].

Mouse embryos lacking functional Gli2 show diminished expression of conserved

Hh targets Gli1 and Ptc1 [46, 47]. Gli2 has an important role in mediating the Hh signal during vertebrate development. Gli2 homozygous null mice die prenatally

and exhibit neural tube defects [46, 47]. Shh signaling regulates Gli2

transcriptional activity by suppressing its processing and degradation [45].

However, Gli2 expression is independent of Hh signaling. Indeed, the Hh-related

upstream mechanisms regulating Gli2 activity is still not clear.

Recent studies suggest that Gli3 has retained multiple functions of Ci.

When Hh is absent, Gli3 undergoes PKA-dependent proteolytic cleavage and

functions primarily as a repressor of gene expression [39, 48, 49]. When Hh is

present, Gli3 is not longer processed by PKA-dependent proteasome activity and

full length Gli3 accumulates to act as a transcriptional activator [41, 49]. Loss-of-

function mutations in Gli3 result in phenotypes that largely resemble those

produced by activation of Hh signaling in both mice and humans [50]. Gli3 may

also participate in the activation of Hh target genes [51, 52]. It is still unclear

whether transcriptional activation by Gli3 results from direct transactivation or

derepression.

The Gli code in vertebrate tissue patterning acts through gradients of

repressor and activator functions in time and space [53]. During development, Gli

activator function is dependent on Hh signaling. Without Hh signaling, dominant

Gli repressors inhibit all Hh responsive Gli activitors. Thus Gli activity is reflected

in the ratio of GliA:GliR.

1.2.3 The role of Shh signaling in holoprosencephaly

Holoprosencephaly (HPE) is the most common developmental defects of

the forebrain and midface in humans [54]. HPE is a cephalic disorder in which the

prosencephalon (the embryonic forebrain) fails to develop into bilateral cerebral

hemispheres. Holoprosencephaly consists of a spectrum of facial defects,

malformation of brain structure and impaired brain function. HPE is also a

common cause of significant morbidity and pregnancy loss (1:250 spontaneous

abortions) [55]. In most cases of holoprosencephaly, the malformations are so

severe that babies die before birth. In less severe cases, babies are born but with

facial deformities which usually affect the eyes, nose, and upper lip.

Human phenotype of HPE ranges from cyclopia, proboscis, median or

bilateral cleft lip/palate in severe forms to ocular hypotelorism or solitary median

maxillary central incisor in minor forms. Children with HPE have many medical problems including developmental retardation, feeding difficulties, epilepsy and instable life signs (body temperature, heart rate and respiration). HPE patients

also have endocrine disorders like diabetes insipidus, adrenal hypoplasia,

hypogonadism, thyroid hypoplasia and growth hormone deficiency.

HPE is caused by incomplete cleavage of prosencephaly in early gestation (18d-28d in human). The etiology of holoprosencephaly is heterogeneous and complex, as causes of HPE involve both non-genetic factors

(environmental or metabolic factors) and genetic factors (chromosomal aberrations or genetic anomalies) [56, 57]. The environmental factors includes insulin-dependent diabetes, maternal alcoholism/smoking and prenatal exposure to drugs or to infections [54]. Chromosomal analysis facilitated the identification of 12 candidate regions (HPE1~HPE12) on 11 chromosomes that may contain genes involved in HPE [58, 59].

To date, seven genes have been positively implicated in isolated form of

HPE: SHH, ZIC2 (Zinc Finger Protein of Cerebellum 2), SIX3 (Sine Oculis

Homeobox, Drosophila, Homolog of, 3), TGIF (Transforming Growth Factor-Beta

Induced Factor), PTCH1 (PATCHED1), GLI2 and TDGF1 (Teratocarcinoma

Derived Growth Factor) [57]. SHH was the first gene identified as a cause of HPE in 1996 [60]. SHH was isolated from the human critical region HPE3 and is the major gene implicated in holoprosencephaly. In the human embryo, SHH is expressed in the notochord and the floorplate, suggesting a critical role in early forebrain development [61]. PTCH1 and GLI2 are the key components of SHH signaling pathways and both of them were identified as HPE-associated genes

[62, 63]. ZIC2 is the second identified HPE gene. The zinc finger domain of all mammalian Zic genes is highly homologous with that of the Gli genes. A recent study has linked ZIC2 to SHH in the degree of HPE features, demonstrating an early role for ZIC2 in the axial midline that precedes the expression of SHH [64].

SIX3 mutations are the third most commonly detected gene mutations in HPE patients. SIX3 is expressed in the anterior forebrain and midline structures,

whose development is defective for HPE pathologies. A recent study showed

defective SIX3 function could impair SHH expression in the ventral forebrain,

which consequently caused HPE [65]. Thus, the core of HPE genetic disorders

can be traced to SHH signaling. Other candidate genes involved in SHH

signaling are DISP1 (DISPATCHED1), SMOH (SMOOTHENED), HHIP (Human

Hedgehog Interacting protein) and the lanosterol synthase gene [54].

Shh-/- deficient mice have indistinct cephalic midline with cyclopia and

often die during embryonic development [66]. Cyclopia is a congenital disorder in

which the embryonic prosencephalon fails to properly divide the orbits of the eye

into two cavities. Zic2 knockdown mice show a strong holoprosencephaly

phenotype [67]. Mice with reduced Six3 expression present a failure of forebrain

and eye development [68]. Gli2 homozygous mutants have a variable loss of

midline structures (pituitary gland) and forebrain tissues [42]. However, the

heterozygous mouse always appears normal.

1.2.4 The role of Shh/Gli signaling in pituitary development

The three Gli genes are all expressed in the ventral diencephalon and

Rathke’s pouch [69] while Shh is uniformly expressed in the ventral diencephalon

and oral ectoderm but is excluded from the invaginating Rathke’s pouch during

E9.5 to E12.5 [31, 33]. Meanwhile, Ptc1 is expressed at high levels in all cells of

Rathke’s pouch suggesting that cells of the nascent pituitary gland are receiving

Hedgehog signals.

Deletion of Shh in the mouse results in malformation of midline structures and cyclopia [66]. These Shh deficient mice do not have even a rudimentary

Rathke’s pouch, but this might result from the loss of ventral forebrain structures

[66]. Pituitary-specific blockage of Hh signaling by overexpression of the Shh antagonist Hip under Pitx1 promoter causes pituitary hypoplasia [31]. In E17.5

Pitx1-Hip transgenic embryos, only a cystic rudiment of the pituitary gland was found [31]. In contrast, Shh overexpression in the developing Rathke’s pouch with αGSU promoter, which drives Shh expression in Rathke's pouch as early as

E10.5, results in pituitary hyperplasia [31]. Taken together, these results suggest that Shh signaling is required for proliferation of cells within the mouse pituitary

gland.

In zebrafish, early treatment with cyclopamine, a pharmacological Hh

blocker, causes a transformation from adenohypophysis to lens at pre-

somitogenesis stage, suggesting an early direct role of Hh in pituitary

specification [70]. Over-expression of Shh expands pituitary gene expression and

Zebrafish Hh pathway mutants display a range of pituitary defects [70, 71].

Zebrafish you-too (Gli2 homolog) mutations disrupt anterior pituitary and ventral

forebrain differentiation [72, 73]. These results provide evidence that Shh plays

an essential role during induction, growth, and patterning of the zebrafish

adenohypophysis.

Other studies also indicated the importance of Gli genes during pituitary

development. Gli2-/- mutants have an incompletely penetrant loss of the pituitary

(3/6) while Gli1; Gli2 double homozygous mutants have a completely penetrant

loss of the pituitary [42]. Loss-of-function mutations in the human GLI2 gene are

associated with pituitary anomalies and holoprosencephaly-like features [62].

These observations suggest that Shh signaling is important for normal pituitary development. However, whether and how Shh/Gli signaling affects pituitary development in a spatially and temporally regulated pattern is unclear.

Also, how does Shh signaling interact with other signaling pathways both in time

and space? It is suggested in a previous study that Shh is required in the oral

ectoderm [31]. As Shh is expressed both dorsally (ventral diencephalon) and

ventrally (oral ectoderm), another important question is whether Shh specifies a

“ventral” fate in the pouch. It remains unknown whether Shh functions to specify

the early fate of the placode in mouse as it does in zebrafish. Gli mutant mice

have defects in ventral diencephalon and pituitary formation, although the

generation of different pituitary cell types has not yet been determined [42]. It is

still unknown how Gli genes mediate Shh signaling to control pituitary

specification and proliferation. As formation of Rathke's pouch requires Bmp4

and Fgf8 signals from the diencephalon [17], the pituitary defects in Gli and Shh

deficient mice might result from abnormal ventral diencephalic development.

Further analysis will be needed to determine whether Gli genes are required for

pituitary primordium specification, cell proliferation and/or cell type differentiation.

1.3 Other signals required in pituitary development

From E9.5 to E12.5, multiple signals in addition to Shh are required for proliferation and patterning of Rathke’s pouch [33]. Morphogens including Wnts,

Fgfs and Bmps are involved in cell type determination in the pituitary, as described below.

1.3.1 Bmps

Bone morphogenetic proteins (Bmps) are members of the transforming growth factor-β (TGF-β) superfamily. Bmps transduce signal via Bmp receptor activation and transcription factor phosphorylation. Bmps first bind to serine/threonine kinase receptors type II. Upon activation by type II receptors, type I receptors phosphorylate receptor-regulated Smads (R-Smads), which in turn form complexes with common-partner Smads (Co-Smads). In mouse embryonic development, R-Smads include the transcription factors Smad1,

Smad5 and Smad8. Mice use a single Co-Smad, Smad4. The R-Smad/Co-Smad complexes then translocate and accumulate in the nucleus, thus activating Bmp- responsive genes [74]. The main extracellular inhibiting regulators of BMP ligands are Bmp antagonists Chordin and Noggin. Chordin and Noggin bind different Bmp molecules with different affinities and regulate dorsal-ventral patterning of early embryos in vertebrates.

Bmp signaling is crucial for anterior pituitary development at a very early stage. By over-expressing the Bmp antagonist Noggin in the oral ectoderm and

Rathke’s pouch, Bmp signaling was blocked and caused pituitary arrest at E10

[33]. In another study, the P-OTX/Ptx1 promoter was used to express Noggin in the oral ectoderm and Rathke’s pouch. In the P-OTX/Ptx1-noggin transgenic embryos the development of Rathke’s pouch was arrested, exhibiting a morphological appearance resembling an E10 Rathke’s pouch, even at E17.5

[33]. Removal of Bmpr1a (Bmp receptor 1a) after the specification of pituitary cell

types using an αGSU-Cre mouse line showed that Bmp signaling is required for

isl1 induction and ventral cell type proliferation [75]. Additionally when a dominant

negative form of Bmpr1a was expressed using the αGSU promoter, ventral cell

types, including the Pit1-lineage, were lost [33]. These results suggest Bmp

signaling may be required for formation and cell specification of Rathke’s pouch.

Bmp4 expression was detected at E9 in the ventral diencephalon and

persisted until E14.5 in the infundibulum, while Bmp2 and Chordin are expressed

in the ventral condensing mesenchyme [17, 33, 75]. Bmp4 signaling has shown

to be required for pituitary induction. Mutation in T/ebp/Nkx2.1, which caused

Bmp4 loss in the ventral diencephalon, gave rise to apoptosis of nascent

Rathke’s pouch [17]. In Bmp4-/- mutant mice, the initial ectodermal thickening and

the invagination of Rathke’s pouch fail to occur [17, 18].

Bmp2 has been shown to participate in the development of anterior

pituitary. Bmp2 expressed from the ventral part of Rathke’s pouch at E10.5 may

form a ventral to dorsal signaling gradient but by E12.5, Bmp2 is expressed

throughout Rathke's pouch [19, 33, 75]. Bmp2 signaling is suggested to provide a

ventralizing cue for pituitary development to generate temporally and spatially

Figure 1-5 Multiple signals required for pituitary development

At E9~E10, Shh (green) is expressed in the anterior part of the ventral diencephalon

while Bmp4 and Fgf8 (blue) are expressed in the medial part of the ventral diencephalon.

Shh expression can also be detected in the oral ectoderm adjacent to Rathke’s pouch.

Meanwhile, Bmp2 is expressed in the ventral most part of Rathke’s pouch. At around

E12, Rathke’s pouch is detached from the underlying oral ectoderm. Shh is expressed in

the ventral diencephalon and oral ectoderm while Bmp4 and Fgf8 are expressed in the

infundibulum. Bmp2 is expressed in the ventral pouch. (Adapted and modified from

Treier et al, 1998 [33])

regulated expression of transcription factors. The expression of Isl1, a ventral

marker in Rathke's pouch, is regulated by Bmp signaling. Explant studies

demonstrated that Bmp2 increases Isl1 expression in Rathke's pouch [19].

Therefore, these results suggest Bmp2 signaling is required for ventral cell type

determination in the anterior pituitary.

1.3.2 Fgfs

The fibroblast growth factor (Fgf) gene family is composed of 22 members.

Fgfs induce their biological responses by binding to and activating Fgf receptors

(Fgfrs), a subfamily of cell surface receptor tyrosine kinases (RTKs). Fgf receptors are single spanning transmembrane proteins with an extracellular

ligand-binding region and an intracellular domain harboring tyrosine kinase

activity (Fgfrs). The vertebrate Fgfr gene family consists of four related genes

(Fgfr1–4). Binding of Fgfs and Fgfrs causes receptor dimerization and triggers

tyrosine kinase activation leading to autophosphorylation of the intracellular

domain. Fgfrs then transmit extracellular signals to various cytoplasmic signal

transduction pathways involved in cell growth, cell differentiation, organ

development, tissue maintenance and wound repair, such as the Ras/MAPK

pathway and the PI3 kinase/Akt pathway [76].

Fgf family members, in particular Fgf8 and Fgf10, have been found to play

important roles in pituitary development. Fgf8 expression was detected in the

ventral diencephalon at E9.5 to E14.5 [19, 33]. At E10.5, Rathke’s pouch has

formed and the infundibulum is visible. A current model suggests that Fgf8, expressed in the infundibulum, functions antagonistically to Bmp2. Misexpression

of Fgf8 under the aGSU cis-regulatory elements in early Rathke’s pouch results

in severe dysmorphogenesis and enlargement of pituitary and loss of ventral cell

lineages including gonadotropes, thyrotropes, somatotropes, and lactotropes [33].

This result suggests Fgf8 prevents ventral-cell-type determination within the pituitary. As Bmp2 signaling is required for ventral cell type determination, it is possible that Fgf8 expressed in the ventral diencephalon/infundibulum counteracts the ventral BMP signal. This proposed dorso-ventral gradient is suggested to induce several temporally and spatially restricted transcription factors, including Pax6, Six3, Nkx3.1 in the dorsal pouch and Isl1, Brn-4, Msx1 in

the ventral pouch [16].

Transgenic studies and in vitro organ culture have shown that Fgf8

stimulates proliferation and Lhx3 expression in developing Rathke’s pouch,

mimicking the roles of the infundibulum [19, 33]. Mouse embryos homozygous

null for Titf1 (also known as T/ebp and Nkx2.1) lose Fgf8 expression specifically

in the ventral diencephalon, which results in a rudimentary Rathke's pouch [17].

In addition, targeted deletion of Fgfr2, a candidate receptor for Fgf8 in the

pituitary, gives rise to a poorly developed Rathke's pouch with severe apoptosis

[77, 78]. Moreover, the Fgf10 knockout mouse also loses its pituitary through an

apoptotic mechanism [77]. All these results suggest a role of Fgf signaling in

initial pituitary organogenesis and survival of pituitary cells.

1.3.3 Wnts

The canonical Wnt pathway is a conserved cell-cell signaling mechanism

in many animals that regulates gene expression via Tcf/Lef DNA-binding factors.

In the absence of Wnt ligand, cytoplasmic β-catenin interacts with APC and Axin scaffold proteins. The APC/GSK-3/Axin complex promotes the proteolytic degradation of the β-catenin, leaving Tcf/Lef to form transcription-repressing complexes with transcriptional co-repressors [79]. In the presence of extracellular

Wnt signaling, Wnt protein binds to cell-surface receptors of the Frizzled family and a coreceptor of the LRP-5/6/arrow family, thus activating Dishevelled family proteins. The APC/GSK-3/Axin destruction complex is inhibited and as a consequence, β-catenin is stabilized and tranlocated into the nucleus where it forms transcription activation complexes with Tcf/Lefs. The Wnt signaling network regulates diverse processes during development such as cell fate determination, structural remodeling, cell polarity and morphology, cell adhesion, and growth.

Members of the Wnt family, Wnt4 and Wnt5a, are expressed in Rathke’s pouch and ventral diencephalon respectively [33]. They have also been shown to be required for anterior pituitary development. Analysis of Wnt4−/− mice at E17.5

revealed a clear reduction in cell number in the anterior pituitary compared to

their wild-type littermates [33]. Wnt5a-/- mice have distorted morphology in the

dorsal part of the pituitary [80]. These results indicate that Wnt signaling is

involved in differentiation or expansion of ventral cell types in the anterior

pituitary.

1.3.4 Signaling interactions in pituitary development

Multiple signals in the developing pouch seem to be crucial for pituitary

cell type differentiation. However, it is not clear how the expression and activity of

the various growth factors are controlled. Considering the complicated nature of

these signaling pathways, pituitary development is likely to involve an intricate

balance between activators and inhibitors of these pathways as well as careful coordination of cross-talk between them. Simple gradients may not be sufficient

for proper morphogenesis of Rathke’s pouch. Signaling interactions,

concentration and exposure time of individual signals should be taken into

consideration. Determining how the downstream signal transduction pathways

integrate these signals in temporal sequence is the biggest challenge in this field.

Dorsal to the developing pituitary, Shh is expressed in the anterior ventral

diencephalon while Bmp4 and Fgf8 are expressed in the medial ventral

diencephalon (Figure 1-5). In the ventral part, Shh is expressed in the oral

ectoderm adjacent to Rathke’s pouch while Bmp2 is expressed in the ventral

Rathke’s pouch. The expression patterns of Shh, Bmp4, Fgf8 and Bmp2 suggest

possible interactive roles of these signaling molecules during pituitary

organogenesis, patterning, proliferation and cell differentiation.

During pituitary organogenesis and patterning, a series of transcription

factors are expressed in the developing pouch (Figure 1-5). Some of these

transcription factors are known downstream targets of specific growth factors or

have interactions with signaling molecules. For instance, GATA2, an important determinant of the gonadotropes and thyrotropes, is a Bmp target [32]. Direct interactions between β-catenin and the tissue-specific homeodomain factor

Prop1 activate Pit1 gene transcription, which is required for pituitary cell-fate determination [81]. Transient dorsal expression of Pax6 in Rathke’s pouch is essential for establishing a sharp boundary between dorsal and ventral cell types, based on the inhibition of Shh ventral signals [30]. Searching for mechanisms which maintain the balance of signaling pathways during pituitary growth would be beneficial to the treatment of pituitary diseases, especially pituitary tumors.

1.4 Shh/Gli signaling to ventral telencephalon

Shh signaling pathway is also involved in the development of the ventral forebrain, especially in dorsoventral patterning. In this section, I introduce the embryonic development of the telencephalon and the role of Shh/Gli signaling in telencephalon development.

1.4.1 Introduction to the ventral telencephalon

The telencephalon is the embryonic structure from which the mature cerebrum develops. The cerebrum comprises what most people think of as the

"brain". It lies in front or on top of the brainstem and in humans is the largest and most superior of the five major divisions of the brain. It determines intelligence

and personality, senses smells and touches, interprets sensory impulses, controls motor function, learns, plans and organizes.

During embryonic development, the telencephalon is the anterior portion of the brain, anterior to the prospective diencephalon (Figure 1-6). The telencephalon or cerebrum, together with the diencephalon, constitutes the forebrain. The telencephalon gives rise to the cerebral cortex, basal ganglia, corpus striatum and olfactory bulb. The dorsal telencephalon (pallium) develops into the cerebral cortex while the ventral telencephalon (subpallium) becomes the basal ganglia.

1.4.2 The embryonic development of the ventral telencephalon

In vertebrates, the anterior neural epithelium undergoes a series of morphological subdivisions to generate vesicle-like structures known as the prosencephalon (forebrain), mesencephalon (midbrain), and rhombencephalon

(hindbrain). During vertebrate embryonic development, the prosencephalon, the most anterior of three vesicles that form from the embryonic neural tube, is further subdivided into the telencephalon and diencephalon. The telencephalon then forms two lateral telencephalic vesicles which develop into the left and right cerebral hemispheres.

Figure 1-6 Telencephalon development

Upper diagram shows a frontal view of brain vesicles during early embryonic

development. Lower diagrams show semi-coronal sections of E12.5 mouse brains from

rostral (left) to caudal (right). Blue lines illustrate the VZ areas and dotted blue lines

show the SVZ. Red dotted lines separate the region of LGE/MGE/CGE from cortex.

Green area indicates Shh expression in the MGE. (VZ-ventricle zone; SVZ-subventricle zone; LGE-lateral ganglionic eminences; MGE-medial ganglionic eminences; CGE- caudal ganglionic eminences. Adapted and modified from Campbell, 2003 [82])

The telencephalon can be further divided along dorsoventral axis as

dorsal telencephalon and ventral telencephalon. The dorsal telencephalon cells

contribute to cortical projection neurons and migrate radially into the cortical mantle. The majority of cortical interneurons derive from the ventral

telencephalon and migrate tangentially into the developing cortex.

The ventral telencephalon is comprised of three proliferative zones: a

medial domain that is known as the medial ganglionic eminence (MGE), and two

posterior and lateral regions that are designated the lateral ganglionic eminence

(LGE) and the caudal ganglionic eminence (CGE) (Figure 1-6). Each of these

regions contributes neurons to different populations in the basal ganglia, which

gives rises to neuronal diversity of the telencephalic structures. Several lines of

evidence suggest that proliferating cells and postmitotic cells in these ganglionic

eminences migrate into different layers of the cerebral cortex.

In utero fate mapping and embryonic transplantation studies suggest LGE

cells migrate ventrally and anteriorly and give rise to the projecting neurons in the

striatum and interneurons in the olfactory bulb while MGE cells migrate dorsally

via the neocortical subventricular zone into the cerebral cortex and give rise to

the major population of projection neurons and interneurons of the ventral

telencephalon and cortical plate [83-87]. CGE cells contribute to distinct cortical

and subcortical cell populations via a unique migration pattern which is distinct

from the MGE and LGE [88].

1.4.3 Shh signaling in telencephalon development

In the developing spinal cord, distinct neuronal subtypes are generated from discrete progenitor domains located along the dorso-ventral axis [89].

However, how the regional production of specific neuronal subtypes occurs within the ventral telencephalon is still unclear. Recent studies have shown that the migration pattern and positioning of transplanted cells is largely determined by donor tissues, suggesting that local signals within the ventral telencephalon determine cell fate and behavior of telencephalic neurons [83, 88, 90-92].

Shh is expressed in early telencephalic tissues, initially in the prechordal plate [93]. At E8.5~E9, Shh and Nkx2.1, which determines fate of MGE-derived progenitors, are expressed in the neural epithelium of the ventral telencephalon

[94]. At E9.5, Shh and Nkx2.1 are expressed in the MGE [95]. By E12.5 in the telencephalon, Shh is still expressed in the mantle zone of the MGE while Nkx2.1 is strongly expressed throughout the proliferative regions of the MGE [95]. The fact that Shh expression consistently overlaps with expression of the MGE marker Nkx2.1 indicates Shh signaling might specify ventral identity and MGE fate within the telencephalon. Consistent with this idea, recent studies have shown that Shh is required to induce and maintain Nkx2.1 in the developing ventral telencephalon [96, 97].

Other genetic studies also suggest Shh signaling is required in telencephalon development. Shh-/- mutants display defects in the presumptive forebrain [66]. However, Shh and Nkx2.1 expression appeared normal in the

subventricular zone of the MGE in Gli1 and Gli2 mutants [42]. Gli3 mutant mice show a strong dorsal telencephalic phenotype, but Shh;Gli3 double mutants show Nkx2.1 expression and a morphologically distinct MGE, suggesting that the primary function of Shh signaling is to prevent the production of excessive Gli3 repressor [98-102]. Conditional targeting using FoxG1-Cre to remove Smo in the rostral neural plate and anterior neural ridge of mouse embryos at E8.5 results in a loss of all ventral telencephalic tissues, suggesting an early requirement for hedgehog signaling in ventral telencephalic patterning [103]. However, conditional mutants using Nestin-Cre to remove Smo in the neural progenitor populations of mouse forebrain at E10~E12 have minor defects, suggesting the main role of Shh signaling at this stage is to maintain progenitor cells and cell migration [104-106].

Chapter 2

Shh/Gli signaling is active in the pituitary primordium

and pituitary cell progenitors

2.1 Abstract

Previous results provide evidence that Shh plays an essential role during induction, growth, and patterning of the zebrafish adenohypoyhysis [70-73].

Moreover, Hh signaling has been shown to play an early direct role in pituitary induction in zebrafish [70]. However, it remains unknown whether Shh exerts its function on early induction of pituitary in the mouse as well. Fate mapping studies were performed to determine whether Hh responding cells can contribute to multiple pituitary cell types. Active Hh signaling was found in the anterior neural ridge and early E7~E8 Hh-responding cells contributed to Rathke’s pouch at E12.

These results suggest Shh signaling is involved in the early development of the pituitary. Hh responsive cells labeled at E10 contributed to all five ventral cell types in the anterior pituitary, indicating a potential role of Shh signaling in cell specification of the anterior pituitary.

2.2 Materials and methods

2.2.1 Mouse breeding and genotyping

Mouse breeding was carried out following approved CWRU IACUC

protocols. For detection of Gli expression, Gli1lacZ/+ and Gli2lacZ/+ heterozygous

male mice were mated with CD1 WT females. For fate mapping experiments,

Gli1-CreER male was crossed with R26R females to generate Gli1-CreER;R26R

embryos. To generate homozygous Gli mutants, both male and female Gli1lacZ/+,

Gli2zfd/+ and Gli3xt/+ mice were bred and then mated. The genotyping of Cre,

Rosa, Gli1lacZ, Gli2lacZ, Gli2zfd and Gli3xt mice was performed as described in

previous papers [44, 107-109].

2.2.2 Fate mapping

To carry out fate mapping studies, Gli1-CreER mouse line was used to express tamoxifen-inducible Cre under the transcriptional regulation of Gli1

(Figure 2-1). R26R has the lacZ gene inserted into the ubiquitously expressed

Rosa locus and preceded with a stop cassette flanked with LoxP sites [107].

Gli1-CreER mice were crossed with R26R and pregnant females were gavaged

with tamoxifen (4mg per 40g body weight). For detection of early Hh-responding

cells, tamoxifen injection was given at E6 to label cells at E7 to E8 and embryos

were collected at E12.5. For detection of the Hh contribution to the anterior

pituitary, tamoxifen injection was given at E9 to label cells at ~E10 and embryos

were obtained at E17.5. X-gal and β-galactosidase (β-gal) stainings were then used detect lacZ expression in labeled Hh-responding cells.

2.2.3 X-gal staining and Immunohistochemistry

For X-gal staining, embryos were dissected in cold PBS and fixed in 0.5% glutaraldehyde for 15-30 minutes at 4°C. For immunohistochemistry (IHC), embryos were fixed in 4% paraformaldehyde (PFA) for 1-3 hours. Then these embryos were washed with PBS, sunk in 30% sucrose/PBS and cryo-embedded in OCT (Tissue-Tek). Tissue sections were collected in sagittal orientation at 10-

12 μm on a Leica Cryostat. At least three embryos were analyzed for each embryonic stage. Methods of X-gal staining and IHC were described in previous papers [44, 110]. Anti-β-galactosidase antibodies (1:500, Biogenesis) were used to detect Cre activity. Hoechst 33258 (Molecular Probes) was used for nuclear counter-staining. The following antibodies were used to examine the five ventral cell types in the anterior pituitary: ACTH (1:1000), GH (1:1000), LH (1:500), PRL

(1:2500) and TSH (1:500) (all from National Hormone & Peptide Program). All sections were then photographed with a Leica DMLB epi-fluorescence microscope fitted with a SPOT camera in the Genetics Imaging Facility

(supported by NIH-NCRR, RR-021228).

Figure 2-1 Fate mapping strategy

A. Gli1-CreER and R26R mouse lines are crossed to generate Gli1-CreER; R26R embryos. Tamoxifen injection can induce Cre-LoxP recombination in the Hh-responding cells (Gli1 as Hh target). Hh-responding cells are labeled permanently at tamoxifen injection time point and can be stained by X-gal or β-gal. (Adapted from Ahn and Joyner, 2004 [107])

B. In Hh-responding cells, Cre recombinase on Gli1 genomic locus is transcribed and translated. Cre-ER is a fusion gene between Cre and a mutant form of the ligand-binding domain of the estrogen receptor (ER). Upon administration of Tamoxifen, Cre-ER is activated by 4-hydroxy (OH)-tamoxifen (TM). Active CreER-TM fusion protein translocates from the cytoplasm to the nucleus, allowing for temporally controlled activation of lacZ gene by removing LoxP-flanked STOP cassette (Adapted and modified from Zhang et al, 2005 [111]).

2.3 Results

2.3.1 Shh signaling is active in the pituitary primordium

2.3.1 a Active Hh signaling in the anterior neural ridge

At around E8 in mouse embryos, Shh is expressed in the prechordal plate and notochord [103]. At that stage, the anterior neural ridge (ANR), which carries both anterior and posterior precursors, can be seen at the rostral margin of neural plate, just anterior to the prechordal plate (Figure 2-2 f). Therefore, pituitary precursors in the midline structures of anterior neural ridge may receive early Shh signals to specify pituitary primordium.

To examine if Shh signaling is active in the pituitary primordium, I utilized the Gli1lacZ mouse line to detect Hh activity in the ANR. Gli1lacZ has a lacZ gene inserted in the Gli1 locus. As Gli1 is a direct target for Hh signaling, lacZ expression can serve as a read-out of Hh signaling [110]. Gli1lacZ/+ embryos were collected at E8 8AM (~E8.25) and whole-mount X-gal staining was performed. In the ventral view and the side view of early E8 embryo (6 somites), I found X-gal staining at the rostral margin of the neural plate, where the ANR resides (Figure

2-2 a,b). Stained embryos were then embedded and sectioned in sagittal orientation. lacZ expression was detected in the ANR in medial sections but not in lateral ones (Figure 2-2 c).

Figure 2-2 X-Gal staining of E8 Gli1lacZ embryos showed active Hh signaling in the anterior neural ridge.

a Side view of an early E8 (6 somites) Gli1lacZ embryo showed X-gal staining in the anterior neural ridge (ANR), indicated by red dash.

b Front view of the same embryo in panel a showed X-gal staining in the ANR (red dash).

c High magnification picture of the midline ANR in sagittal section of E8 (6 somites) Gli1lacZ embryo. Inset: sagittal section of 6 somites Gli1lacZ embryo with red dash boxed area showing the ANR.

d Side view of 12 somites Gli1lacZ embryo showed X-gal staining in the ANR (red arrow).

e Sagittal view of bisected Gli1lacZ embryo in panel d showed X-gal staining in the ANR (red arrow), where is the junction of prospective diencephalon and oral ectoderm.

f Dorsal view of mouse neural plate at E8. Shh is expressed in the prechordal plate and the notochord (green). The most anterior part of the neural plate filled with red is the ANR. Red dashed area filled with blue is the midline ANR carrying pituitary precursors.

The ANR carries the pituitary origins of both the anterior pituitary and the posterior pituitary, as the oral ectoderm and the neural ectoderm, respectively.

My result showed X-gal staining in the ANR in medial sagittal sections of early E8 embryos. It suggests active Hh signaling in the midline structures of the ANR, which carries the precursor of pituitary gland. Detection of early Hh activity in the anterior neural ridge, indicated by Gli1-lacZ expression in early E8 embryos, suggests that Shh signaling is active in pituitary precursors. A current model suggests Shh signaling is required for pituitary proliferation at E9.5 to E12.5 [31].

My result suggests that Shh signaling may function as early as E7~E8, before the induction of the initial pouch.

2.3.1 b Hh-responding cells in the ANR contribute to Rathke’s pouch

Active Hh signaling detected in the ANR suggests that Hh may play an early role in specifying the pituitary primordium. To ascertain Hh-responding cells in the ANR can contribute to the pituitary primordium, I used the fate mapping method to label Hh-responding cells in the ANR (Figure 2-1).

Gli1-CreER mouse line has a tamoxifen inducible Cre under the transcriptional regulation of Gli1 [107]. R26R mouse line has a lacZ gene inserted into the ubiquitously expressed Rosa locus that is preceded by LoxP- flanked STOP cassette [107]. Gli1-CreER mouse line was crossed with R26R to generate Gli1-CreER; R26R embryos with the administration of tamoxifen. Hh- responding cells can be detected by X-gal staining. Pituitary precursors in the

anterior neural ridge develop into Rathke’s pouch and infunbibulum. Thus I expected Hh-responding cells in the ANR contribute to further-developed pituitary tissues.

In mouse embryos, Shh is expressed in the neural plate at E6.5 and Gli1 expression is detected at late E7 [44]. Tamoxifen induced Cre-LoxP recombination is usually active at about 6 hours post injection and last till 36 hours post injection [112]. To trace the early Hh-responding cells before the formation of Rathke’s pouch, tamoxifen injection should be performed earlier than E9. I crossed the Gli1-CreER and R26R mouse line and injected the pregnant female mice with 4mg per 40g body weight tamoxifen at E6 12PM

(Figure 2-3 a). Gli1-CreER; R26R embryos were collected at E12.5.

Both whole mount and section X-gal staining was performed to detect Hh responsive cells. E12.5 embryos were bisected and then stained with X-gal. Blue staining was found in Rathke’s pouch, the infundibulum and the hind brain

(Figure 2-3 c). In sagittal sections of embryos, a small group of X-gal-labeled cells was detected in the pouch (Figure 2-3 d). Since the tamoxifen injection at

E6 12PM labels the Hh-responding cells during E7.75 to E8 (Figure 2-3 a), these blue cells are considered as the initial population of Hh responding cells coming from the ANR and contributing to both the anterior pituitary and posterior pituitary.

Figure 2-3 Hh-responding cells in the ANR contribute to Rathke’s pouch. a Fate mapping of Hh responsive cells at E7~E8 (red bar) by tamoxifen induction at E6 12PM (purple arrow). b Diagram of Rathke’s pouch at E12.5 in sagittal sections. c Whole mount X-Gal staining of bisected Gli1-CreER;R26R embryo at E12.5 showed Hh-responding cells in Rathke’s pouch labeled at E7 to E8. Tamoxifen gavage was given at E6 12PM. Rathke’s pouch and infundibulum was illustrated with red dashes. (RP-Rathke’s pouch; INF-infundibulum; HB-hindbrain) d X-Gal staining of E12.5 Gli1-CreER;R26R embryo sections showed Hh-responding cells in Rathke’s pouch labeled at E7 to E8. Tamoxifen gavage was given at E6 12PM.

2.3.2 Gli expression gradient in developing Rathke’s pouch

Shh is expressed in the ventral diencephalon and the oral ectoderm during

pituitary patterning, cell proliferation and cell type determination [33]. Inhibition of

Hh signaling in Rathke’s pouch by Pitx1-Hip transgenic strategy caused pituitary

hypoplasia and a loss of induction of ventral transcription factors expressed at

the Shh boundary [31]. These results suggest Shh signaling plays a critical role

in pituitary development. However, when and how Hh signaling contributes to

pituitary cell type specification has not been determined. To determine the

temporal and spatial requirements of Shh/Gli signaling in pituitary cell type

differentiation, I performed Gli expression analysis.

Gli1 is a direct Hh target and its expression pattern serves as good

indicator of active Shh signaling. Gli1 expression was found in both the ventral

diencephalon and Rathke’s pouch [69]. Here I used Gli1lacZ embryos at different

stages to analyze Gli expression pattern.

Gli1lacZ/+ embryos were collected at E8 (~13 somites) and whole mount X- gal staining was performed. Strong expression was detected in the diencephalon

(Figure 2-4 a). Stained embryos were sectioned in the anterior coronal orientation as shown in Figure 2-4 (a’,b’, red lines). Coronal sections of E8 embryos showed Gli activity in the ventral diencephalon and underlying oral ectoderm (Figure 2-4 a’). Enlarged view of coronal sections showed Gli1 expression in the ventral diencephalon at late E8 (Figure 2-3 a’,c’).

Figure 2-4 Gli activity can be detected in the ventral diencephalon and Rathke’s pouch. a Side view of E8 (13 somites) Gli1lacZ embryo showed Gli activity in the diencephalon. a’ and b’ are the coronal sections obtained from this embryo shown in cutting direction of red lines in panel A. c’ is the zoomed in view of the red box in panel a’, showing X-gal stained cells in the midline of ventral diencephalon. Scale bars in the pictures stand for 100 µm.

b-e X-gal staining of Gli1lacZ sagittal sections at E9 (B), E10 (C), E11 (D) and E12 (E) showed Gli activity in the Rathke’s pouch. Anterior is to the left.

(RP-Rathke’s pouch; VD-ventral diencephalon; Di-diencephalon; INF-infundibulum)

At E9-E12, Gli1lacZ/+ embryos were sectioned in sagittal orientation for X- gal staining. Rathke’s pouch arises at ~E9 and is not separated from the oral ectoderm until E11. Before RP is detached from the oral ectoderm, Gli1 expression is higher at the anterior side of the pouch than the posterior (Figure 2-

4 b,c). At E11, the anterior to posterior Gli expression gradient in Rathke’s pouch is not so obvious. Gli expression is quite even in the pouch at this time (Figure 2-

4 d). At E12, Gli expression is much stronger in the ventral part (Figure 2-4 e).

2.3.3 Hh-responding cells contribute to ventral cell types in the anterior pituitary

There are six cell types in the anterior pituitary: melanotropes (M) in the dorsal part and somatotropes (S), lactotropes (L), thyrotropes (T), gonadotropes

(G), corticotropes (C) at the ventral part. At E17.5, all the cell types become mature and are able to secrete different hormones as MSH, GH, Prl, TSH, LH and ACTH, respectively. To investigate whether Hh responsive cells in RP contribute to different pituitary cell types, I used fate mapping to label the Hh- responding cells at E10 (described above).

Tamoxifen injection was given at E9 6PM so that Hh responsive cells could be labeled at around E10. Embryos were collected at E17.5 and sectioned in sagittal orientation. Different anti-hormone antibodies were used as cell type specific markers. Double immunostaining of β-gal and cell type specific markers were performed. As shown in Figure 2-5, double labeling cells (yellow) of β-gal

Figure 2-5 Hh-responding cells contribute to all five ventral cell types in the anterior pituitary. a-e Double immuno-staining of beta-Gal and pituitary hormone markers showed Hh- responding cells and five ventral cell types of the anterior pituitary. Different pituitary cell types were indicated by ACTH (C); GH (S); LH (G); Prl (L) and TSH (T). a’-e’ Enlarged view of a-e showed double labeled cells for each pituitary cell type (white arrow).

(blue-nuclei; green-β-Gal; red-pituitary hormone specific antibody; yellow-double labeling)

(green) and cell-type-specific marker (red) in the anterior pituitary were shown.

Cell-type-specific hormone labels the different cell types. In each cell type, β-gal- positive cells are fate mapped Hh-responding cells. Double labeling cells are Hh- responding cells which contribute to certain cell type of the anterior pituitary.

Double labeling cells were detected in every cell type, suggesting that Hh- responding cells contribute to all five ventral cell types. In each individual cell type, Hh-responding cells only occupied a small portion of the total cells. It might reflect the low efficiency of tamoxifen induced Cre-LoxP system. Further analysis is planned to determine whether there is contribution preference to certain cell type of Hh-responding cells labeled at different time period.

2.4 Discussion

Active Hh signaling was detected in the midline of the anterior neural ridge, which is the precursor of both the anterior and posterior pituitary (Figure 2-2).

Shh is expressed in the prechordal plate adjacent to the midline structures of anterior neural ridge [94]. Thus the pituitary primordia are receiving Shh signals at E8, before the initial formation of Rathke’s pouch. Hh responding cells labeled at E7~E8 were found in Rathke’s pouch (Figure 2-3). This suggests Shh/Gli signaling may play a role in specifying pituitary precursor cells in the ANR as in driving their proliferation. To further test this hypothesis, ANR specific Gli knockout analysis will be performed.

Hh-responsive cells in RP contribute to five ventral cell types of the anterior pituitary (Figure 2-5). This suggests Hh responding cells can develop into multiple cell types. To further determine the role of Shh/Gli signaling in cell type specification, affected cell types in Gli mutants or pituitary-specific Gli mutants will be investigated. Loss of certain pituitary cell type(s) is expected if

Shh/Gli signaling plays a role in pituitary cell differentiation.

At E9 to E10, Gli expression has an anterior to posterior gradient in

Rathke’s pouch, reflecting higher Hh response at the anterior side of the pouch

(Figure 2-4 b,c). However, after Rathke’s pouch is separated from the underlying oral ectoderm, this gradient is no longer present (Figure 2-4 d). Later at E12, Gli gradient shifted to a dorsoventral model, with higher Hh response in the ventral part of the pouch (Figure 2-4 e).

As proposed in a previous paper, Rathke’s pouch receives Shh signals from the oral ectoderm at E9 to E12 [31]. Blocking Hh signaling specifically in RP caused pituitary hypoplasia [31]. However, this result does not indicate oral ectoderm as the only Shh source. According to my result, expression of Gli1 and

Gli2 suggests higher Hh response in the anterior side of Rathke’s pouch at E9 and E10 (Figure 2-4, appendix I). Anterior to posterior pattern of Hh response in

Rathke’s pouch did not reflect ventral shh signals (oral ectoderm) as a major source. At E9 and E10, Shh is expressed at high level in the ventral diencephalon besides in the oral ectoderm [33]. This suggests that Rathke’s pouch may receive Shh signals from the ventral diencephalon as well.

2.5 Acknowledgements

I thank Kristen McDonnell, a former lab member, for her kind help with my work. I thank Drs. S. Camper for providing antibodies. I also thank Dr. Patricia

Conrad for assistance with microscopy and imaging.

Chapter 3

Direct and indirect requirements for Shh/Gli signaling

in early pituitary development

This Chapter was published as Wang, Y., et al. (2010). "Direct and indirect requirements for Shh/Gli signaling in early pituitary development." Dev Biol

(in press). I did all the experiments and quantification analysis in this chapter and contributed to the writing.

3.1 Abstract

Induction of early pituitary progenitors is achieved through combined activities of signals from adjacent embryonic tissues. Previous studies have identified a requirement for oral ectoderm derived Sonic Hedgehog (Shh) in specification and/or proliferation of early pituitary progenitors, however how different Gli genes mediate Shh signaling to control pituitary progenitor development has not yet been determined. Here we show that Gli2, which encodes a major Gli activator, is required for proliferation of specific groups of pituitary progenitors but not for initial dorsoventral patterning. We further show that the action of Gli2 occurs prior to the closure of Rathke’ pouch. Lastly, we show that Shh/Gli2 signaling controls the diencephalic expression of Bone morphogenetic protein 4 (Bmp4) and Fibroblast growth factor 8 (Fgf8), two genes that are known to play critical roles in patterning and growth of Rathke’s pouch.

Our results therefore suggest both cell-autonomous and non-cell autonomous requirements for Gli2 in regulation of pituitary progenitor specification, proliferation and differentiation.

Key words: Gli1, Gli2, Gli3, Shh, mouse, pituitary, patterning, proliferation

3.2 Introduction

The pituitary gland is a master endocrine organ that produces a number of hormones regulating many essential physiological functions in the body. It is comprised of adenohypophysis (anterior and intermediate lobes) and neurohypophysis (posterior lobe). The anterior lobe contains five different hormone-producing cell types that secrete adrenocorticotropic hormone (ACTH), growth hormone (GH), prolactin (PRL), thyroid stimulating hormone (TSH), luteinizing hormone (LH) and follicle-stimulating hormone (FSH) while the posterior lobe contains nerve endings that secrete oxytoxin and vasopressin [15,

Reviewed in 113]. Both hypopituitarism and hyperpituitarism have been identified in a number of human congenital pituitary disorders [4-7, 62].

Development of the pituitary gland requires extensive interactions between different embryonic tissues. Between ~E8.5 and E9.0, a portion of the oral ectoderm thickens and invaginates to form a rudimentary structure called

Rathke’s pouch (RP). Several lines of evidence suggest that induction of RP requires signals derived from the diencephalon. First, mutation in T/ebp (Nkx2.1), which is expressed in the ventral diencephalon but not in RP, causes apoptosis and degeneration of nascent RP [17]. Second, mutation in Bmp4, which is expressed in the ventral diencephalon, causes a loss of early RP [17, 18]. Similar phenotypes were also observed in embryos over expressing Noggin, a BMP antagonist, in the oral ectoderm [33]. On the other hand, mutation in Noggin results in expansion of pituitary tissues [75]. Lastly, analysis of embryos mutant for Fgfr2, which encodes a receptor for FGF1/3/7/10 and is expressed in the

diencephalon, revealed a requirement for FGF signaling in the diencephalon [78].

In both Fgfr2 and Fgf10 mutants, increased apoptosis causes severe defects in infundibulum and RP development [77, 78]. By ~E11, RP closes off from the oral ectoderm and by ~E12, RP detaches from the oral ectoderm. Interestingly, the critical period for pituitary progenitor specification appears to be between ~E10 and ~E12/13. Nascent RP isolated from E10.5 mouse embryos will express early markers of pituitary pouch Isl1 but not Lhx3 (Lim1) when cultured in vitro. When combined with ventral diencephalic tissues, nascent RP can be induced to express Lhx3 and other pituitary markers [19]. The action of diencephalic tissues can be recapitulated by beads soaked with FGF8, suggesting at least that FGF8 secreted from the diencephalon is critical in specifying different pituitary cell types. However, by E11.5, the pituitary explants become refractory to FGF8 induction [19]. By E12/13 (E13/15 in rat embryos), isolated RP can differentiate into different pituitary cell types even when cultured alone, suggesting that pituitary progenitors have already been specified when RP detaches from the underlying oral ectoderm [114].

In addition to dorsal diencephalic signals, another signaling molecule Shh, which has been shown to function as a morphogen and induce different cell types at a concentration-dependent manner in several embryonic tissues

[Reviewed in 115], is expressed in the oral ectoderm immediately adjacent to RP

[33]. The requirement of Shh signaling in pituitary development is supported by the following experiments. Over-expression of Shh in the developing RP using an

αGSU promoter, which drives ectopic expression of Shh in RP starting from

~E12.5, results in an expansion of ventral pituitary cell markers TSH and LH and causes pituitary hyperplasia [31]. On the other hand, blocking Shh signaling in

RP through expression of a Shh antagonist, Hedgehog-interacting protein (Hhip), using the Pitx1 promoter causes pituitary hypoplasia [31]. However, interpretation of some of these results is complicated by the fact that manipulations of secreted signals can affect multiple tissues. In addition to the oral ectoderm, Shh is also abundantly expressed in the ventral diencephalon, immediately above the early RP. As a result, RP may be exposed to Shh derived from both oral ectoderm and ventral diencephalon, and the above experiments might have affected oral ectoderm and diencephalon development in addition to development of RP. Preliminary analysis of the pituitary phenotype resulting from a loss of Gli2, which encodes the major Gli activator, provides some initial insight into the requirement of Gli genes. Mutations in Gli2 result in a variable loss of pituitary, demonstrating a requirement of Shh/Gli2 signaling in pituitary development [42]. However, whether different cell types are generated in the anterior pituitary or whether the development of posterior pituitary is affected has not yet been determined. To address how Gli genes mediate Shh signaling to control pituitary specification and proliferation, we analyzed pituitary phenotypes of embryos mutated for individual Gli genes, which are required for the transcription output of Shh signaling. We then conditionally disrupted Gli genes in

RP and found a critical role for Shh signaling before RP detached from the oral ectoderm. Lastly, we found Shh signaling is also required in the diencephalon to control pituitary development through the regulation of two critical growth factors.

3.3 Materials and methods

Mouse breeding

Mouse breeding was carried out according to protocols approved by Case

Western Reserve University (CWRU) Institutional Animal Care and Use

Committee. Rosa26-lacZ Reporter (R) mouse [116] was used to monitor Cre-

mediated recombination. To generate FoxG1Cre;Gli conditional mutant embryos,

FoxG1Cre;Gli2zfd/+ male mice were bred to Gli2flox/flox;R/R and

FoxG1Cre;Gli2zfd/+;Gli3xt/+ were bred to Gli2flox/flox;Gli3xt/+;R/R female mice. To

generate embryos with activated Hh signaling in RP, Pitx2Cre male mice were

bred to R26SmoM2 female mice. To generate Gli2 mosaic embryos, Gli2lacZki/+

mice were bred with Gli2zfd/+; Rosa26 (R26)-EGFP/R26-EGFP mice. The resulting morula stage embryos were co-cultured with CD1 embryos. Embryos

that had aggregated after overnight culture were then transferred into pseudo- pregnant female for further development. The genotyping of Cre, Gli2flox, Gli2lacZki,

Gli2zfd and Gli3xt mice has been described [109, 117-121].

Histology, immunostaining and RNA in situ hybridization

Embryos were dissected in cold PBS and fixed in 4% paraformaldehyde (PFA)

for 1-3 hrs at 4°C, washed with PBS, sunk in 30% sucrose and embedded in

sagittal orientation in OCT (Tissue-Tek). Tissue sections were collected at 12 μm on a Leica Cryostat. About 3-6 embryos were analyzed for each embryonic stage.

Methods of X-gal staining, antibody staining and RNA in situ hybridization have

been described (Bai et al., 2002; Bai and Joyner, 2001). The following antibodies

were used: Pax6 (1:500, Covance), Isl1 (1:100, Abcam), Lhx3 (1:100,

Developmental Studies Hybridoma Bank), BrdU (1:20, Becton-Dickinson), Ki67

(1:500, Abcam), phospho-Histone H3 (1:100, Upstate), cleaved Caspase-3

(1:200, Cell Signaling), ACTH (1:1000), GH (1:1000), LH (1:500), PRL (1:2500) and TSH (1:500, all from National Hormone & Peptide Program) and β- galactosidase (1:500, Biogenesis). Hoechst 33258 (Molecular Probes) was used to visualize nuclear staining. All sections were examined with a Leica DMLB epi-

fluorescence microscope fitted with a SPOT camera in the Genetics Imaging

Facility (supported by NIH-NCRR, RR-021228).

BrdU labeling, cell cycle, cell death analysis and quantification

Pregnant mice were injected intraperitoneally with BrdU (Sigma) at 100μg/g body

weight 1hr prior to dissection. The percentage of BrdU+, Ki67+, pHH3+,

Caspase3+ cells was calculated by dividing the number of the positive cells by

the total number of nuclei (Hoechst) inside RP or specific regions. To calculate

the fraction of S-phase cells within cell cycle, we divided the number of BrdU+

cells by the total number of Ki67+ cells and converted the number to a

percentage. For each time point, we used 2 adjacent mid-sagittal sections from

each embryo and 4-6 embryos from at least two different litters for analysis. As

the number of Caspase3+ cells is low in both WT and Gli2 mutants, we counted

3-4 sections from each embryo (total 1-5 Caspase3+ cells per embryo, with n=3

for Gli2 mutants and n=6 for WT). The number in charts was displayed as mean

± S.E.M. Student's t-test was used to calculate the P value and to determine

whether the results were significantly different from each other (** indicates

P<0.01; * indicates P<0.05).

Quantification of pituitary cell types was applied at E17.5. To ensure consistent sampling of cell types among different embryos, we performed immunostaining analysis for each marker on 2 adjacent, mid-sagittal sections from each embryo. A total of 6 WT embryos, 6 Gli2-/- embryos and 4

FoxG1Cre;Gli2-/- embryos from at least three litters were used in the analysis.

We used the following method to normalize the contribution of EGFP-labeled cells to RP of mosaic embryos. We first selected an area in the dorsal mid/hindbrain region, which does not have active Shh/Gli signaling, and calculated the percentage of EGFP+ cells over the total number of cells. This number was used as a baseline for the level of mosaicism in that embryo. We then calculated the percentage of EGFP+ cells within RP or in different regions of

RP, and normalized the percentage against the baseline in that embryo. Results from three embryos were then used to determine whether there was a significant difference between the contribution of WT and Gli2 mutant cells.

3.4 Results

3.4.1 Loss of Gli2 leads to a selective loss of specific pituitary cell types

Previous studies reported a variable loss of pituitary (3/6) in Gli2 mutants when examined at E12.5 [42]. To determine how Gli2 mediates Shh signaling to influence pituitary development, we examined patterning of the pituitary gland and the production of different pituitary cell types in wild type (WT) and mutant embryos.

We first examined the expression of two markers that are normally expressed throughout E12.5 pituitary pouch, Lhx3 and Prop1. Lhx3 encodes a

LIM-homeodomain transcription factor that is required for specification of pituitary cell lineages [29] and Prop1 encodes a paired-like homeodomain transcription factor required for initial determination of the Pit1 (Poulf1)-expressing lineage of the pituitary [23]. In Gli2 mutants, both Lhx3 and Prop1 were found throughout the pituitary gland, suggesting that early pituitary cells were specified normally

(Figure 3-1A-D). We then examined the expression of markers for different regions of the pituitary pouch. Gata2 encodes for a ventrally restricted transcription factor that is normally expressed in pituitary cells located at the anterior tip of RP, most of which have just exited cell cycle [32, 33] (Figure 3-1E).

In Gli2 mutants, Gata2+ cells were found in the anterior tip of the pouch, as was the case in WT embryos (Figure 3-1F). We also examined the expression of two other pituitary markers, Pax6 and Isl1. Pax6 is normally expressed in a dorsal-to- ventral gradient and Isl1 is restricted to the pituitary tip at E12.5 (Figure 3-1G,I).

The expression pattern of Pax6 and Isl1 in Gli2 mutants is similar to the pattern

seen in WT embryos (Figure 3-1H,J). This demonstrates that loss of Gli2 does not affect the overall patterning of the pituitary.

Although patterning of the Gli2 mutant pituitary is largely normal, most Gli2 mutant pituitaries were found to be smaller than WT pituitaries (Figure 3-1), suggesting defects in cell proliferation and/or excessive cell death in mutant pituitary tissues. To distinguish these two possibilities, we examined expression of Ki67, a marker for all proliferating cells, BrdU incorporation, a marker for cells in S-phase, and phospho- Histone H3 (pHH3), a marker for cells in M-phase. At

E12.5, we detected a significant reduction in the percentage of cells expression

Ki67 in Gli2 mutant pituitaries (Figure 3-1K,L,Q, P=0.009). Additionally, we found a significant reduction in the percentage of BrdU+ cells in Gli2 mutants (Figure 3-

1K,L,Q, P=0.0006). Finally, we also found reduction in the number of cells in M- phase based on pHH3 levels (Figure 3-1M,N, P=0.0123). To exclude the possibility that the smaller pituitary pouches found in Gli2 mutants were caused by excessive cell death, we examined the expression of cleaved-Caspase3 and found no significant difference between mutant and WT pouches (Figure 3-1O,P).

Together, these data suggest that loss of Gli2 affects proliferation of pituitary progenitor cells at E12.5.

Figure 3-1 Gli2 mutant pituitaries have normal patterning but show defects in proliferation. (A-F) RNA in situ hybridization of wild type (WT) and Gli2 mutants using probes of Lhx3, Prop1 and Gata2. (G-J) Immunostaining of Pax6 and Isl1. (K-P) Analysis of proliferation and cell death by immunostaining of Ki67, BrdU, pHH3 and Caspase3 in Rathke’s pouch. Note: The strong background staining in (L) is caused by elevated levels of mouse endogenous IgG in Gli2 mutants. (Q) Quantification of BrdU incorporation, the expression of Ki67, pHH3 and Caspase3. All embryos (n=6) were at E12.5. * indicates P<0.05, ** indicates P<0.01. Scale bars are 100 µm.

To determine whether loss of Gli2 affects the production of different

pituitary cell types, we analyzed the expression of cell-type specific markers at

E17.5, a stage when all mature cell types should be present (Figure 3-2A-J). Five hormone-secreting cell types we examined include corticotropes (C), which secrete ACTH; thyrotropes (T), which secrete TSH; gonadotropes (G), which secrete LH; somatotropes (S), which secrete GH; and lactotropes (L), which secrete PRL. We found that all five markers were detected in Gli2 mutant pituitaries (Figure 3-2A-J). However, when the number of different cell types was counted in Gli2 mutants, we found a drastic reduction in the number of three cell types, C, S and L (Figure 3-2K), suggesting a specific requirement of Gli2 in

specification of these three cell types.

3.4.2 Progenitor proliferation in the nascent pituitary requires cell-

autonomous expression of Gli2

Several possibilities could account for the proliferation defects in Gli2

mutant pituitaries. First, Gli2 is required cell-autonomously in RP to respond to

Shh signal derived from the oral ectoderm, as suggested by previous studies [33].

Second, Gli2 could be required for cells in RP to respond to diencephalic derived

Shh. Third, Gli2 may be required in the diencephalon for the expression of other

growth signals, as diencephalic defects were found in Gli2 mutants in our

previous study [42]. To distinguish these possibilities, we generated Gli2 mosaic embryos and analyzed mutant cells in an otherwise WT background.

Figure 3-2 Production of specific cell types is affected in Gli2 mutants at E17.5. (A-J) Immunostaining of different pituitary cell type markers ACTH, GH, LH, PRL and TSH in WT and Gli2 mutant pituitaries. (K) Quantification of each pituitary cell type in WT and Gli2 mutants (* indicates P<0.05, ** indicates P<0.01, n=6 embryos). Two mid-sagittal sections from each embryo were used to quantify the number of each cell type.

Two different Gli2 alleles, Gli2zfd and Gli2lacZki [109, 119], were used to generate Gli2 mutant embryos. EGFP+ embryos were derived by crossing

Gli2lacZki/+ and Gli2zfd/+; EGFP/EGFP mice. The EGFP-labeled embryos were then aggregated with morula stage, un-labeled CD1 embryos in vitro. After overnight incubation, those embryos that had successfully aggregated were transferred into pseudo-pregnant females and allowed to develop. Embryos were then dissected and genotyped with primers for the two Gli2 alleles. Those that contained Gli2zfd, Gli2laczki and WT Gli2 alleles were mosaic embryos containing

WT and Gli2 mutant cells while those that contained only Gli2lacZki and WT Gli2 alleles were used as controls. As expected, both control and mosaic embryos appeared to have morphologically normal RP (Figure 3-3A-F). To determine whether Gli2 mutant cells can compete with WT cells and adopt a pituitary cell fate, we stained mosaic pituitary with Lhx3 antibody (Figure 3-3A,D). We found that all EGFP+ Gli2 mutant cells expressed Lhx3 in mosaic pituitary examined

(Figure 3-3B,E, n=3), suggesting that Gli2 mutant cells can contribute to pituitary and adopt pituitary cell fate in mosaic embryos (Figure 3-3C,C’,F,F’). However, there were much fewer EGFP positive cells in mosaic Gli2 pouches (Figure 3-

3E,K,N) than in control (Figure 3-3B,H). To determine whether EGFP+ mutant cells are less likely to be integrated into mosaic pituitary, we divided RP into two regions, anterior wall (A) and posterior wall (P), along the pouch lumen (Figure 3-

3M). When we calculated the percentage of EGFP+ cells in each division and normalized this number to the level of mosaicism in each embryo (see Materials and Methods), we found mutant cells contributed significantly less to the anterior

wall of the pituitary (Figure 3-3N, PA= 0.002). Interestingly, mutant cells contributed less to the posterior wall as well, although the reduction was not as dramatic (PP=0.031). Lastly, mutant cells were also underrepresented in the infundibulum (PI=0.011), suggesting a requirement of Shh/Gli2 signaling in the development of posterior pituitary.

One potential reason why there are fewer mutant cells in the anterior wall of the pouch is that mutant cells do not proliferate as well as WT cells. To determine whether this was the case, we counted the number of EGFP+ BrdU+ mutant cells and the total number of unlabeled, BrdU+ S-phase cells (Figure 3-

3G-L), and calculated the percentage of mutant cells over the total number of S- phase cells in each region (Figure 3-3O). Indeed, we found much fewer proliferating EGFP+ cells in anterior wall of the pouch (Figure 3-3O). There were also fewer proliferating EGFP+ cells in the posterior wall, although the reduction was not as drastic as in the anterior wall of the pouch, suggesting that the reason there were fewer Gli2 mutant cells inside RP is that there is a cell-autonomous requirement of Gli2 for cells to proliferate.

Figure 3-3 Gli2 mutant cells do not proliferate as well as WT cells especially in the anterior edge of the pituitary, as revealed by mosaic analysis. (A-F) Immunostaining of Lhx3 in WT and Gli2 mosaic embryos. (C’,F’,I’,L’) are higher magnification images of corresponding boxes indicated. Note: EGFP is expressed in cytoplasm while Lhx3 or BrdU is expressed in nuclei. (G-L) Immunostaining of BrdU and EGFP in WT and Gli2 mosaic embryos. EGFP+ cells in WT embryos (A-C, G-I) were Gli2+/+ while EGFP+ cells in Gli2 mosaic (D-F, J-L) were Gli2zfd/lacZki. (M) Schematic representation of the subdivisions of the pituitary. The anterior pituitary was outlined by dotted lines. White arrowheads indicate cells double-labeled by both green and red. (N) Distribution of EGFP+ cells in anterior wall (A) and posterior wall (P) of the anterior pituitary, and in infundibulum. (O) Distribution of BrdU+;EGFP+/BrdU+ cells in different regions of the anterior pituitary. See Material and Methods for normalization of the number of EGFP+ cells in each embryo. Scale bars: 100 µm. * represents P<0.05, ** represents P<0.01, n=3 embryos.

3.4.3 Removal of Gli2 function before the closure of RP causes proliferation defects in some pituitary progenitors

Previous studies have shown that pituitary development is controlled by

Shh expressed from the oral ectoderm [31, 33]. To determine the critical period of Shh/Gli signaling that controls pituitary proliferation, we conditionally removed

Gli2 function in the pituitary by using a conditional Gli2 allele and two Cre lines,

FoxG1Cre [122] and Pitx2Cre [123].

FoxG1 (Forkhead box G1) is initially expressed in the anterior neural ridge and in the early telencephalon [124]. To determine whether FoxG1Cre can be used to remove Gli2 function before RP closes, we used FoxG1Cre to recombine a Rosa26-lacZ reporter (R) and monitored the activity of β-galactosidase expressed from the Rosa26-lacZ reporter. We found that at E9.5, all cells encompassing the invaginating RP were positive for β-galactosidase (Figure 3-

4A). The ubiquitous staining of β-galactosidase+ cells within RP was confirmed at E10.5 and E12.5 (Figure 3-4B,C), suggesting that FoxG1Cre can be used to disrupt Gli2 function before the closure of RP. As FoxG1 is not expressed in the diencephalon, the manipulation should not affect the normal role of the diencephalon in regulating the growth and patterning of the pituitary.

At E12.5, the conditional Gli2 mutants (FoxG1Cre; Gli2flox/zfd) appeared morphologically normal. Analysis of early pituitary markers, Lhx3, Porp1, Gata2,

Pax6 and Isl1, also revealed a pattern of expression resembling that of the WT embryos (Appendix II Figure S1A-K), consistent with our studies that Gli2 does not control pituitary patterning (Figure 3-1). Nevertheless, we reasoned that if

Gli2 is required to mediate Shh signaling to control progenitor proliferation before the closure of RP, then we would expect proliferation defects in the FoxG1Cre;

Gli2flox/zfd conditional mutants. To this end, we examined the proliferation of

conditional mutant cells at E12.5 by analyzing BrdU incorporation and expression

of Ki67 and pHH3. Compared with WT embryos, we found no significant differences in the percentage of cells incorporating of BrdU, or cells expressing

Ki67 and pHH3 in conditional Gli2 mutants (Figure 3-4D,G). Furthermore,

removal of Gli2 did not change the pattern of cell death, as there was no

significant change in the number of cells expressing Caspase3 (Figure 3-4F,I).

To further determine whether there are regional differences in cell proliferation,

we divided the pouch into four regions: anterior (dorsal and ventral, Ad and Av respectively), rostral tip (rT) and posterior (P) (Figure 3-4J). At this stage, most of

the rT cells had already exited the cell cycle while the rest of the cells in RP were

still proliferating. Interestingly we found a decrease in the percentage of BrdU

incorporation in the Av region of the FoxG1Cre;Gli2 mutants (Figure 3-4K).

However, the reduction in BrdU incorporation appeared to reflect a reduction in

proliferation, rather than changes in cell cycle length, as the number of

proliferating cells (Ki67+) was also decreased and the ratio of BrdU/Ki67

remained largely unchanged (Figure 3-4L).

Figure 3-4 Early loss of Gli2 function results in proliferation defects in pituitary progenitors. (A-C) FoxG1Cre activity can be detected before the closure of RP, as detected by X-gal staining at E9.5 (A) and E10.5 (B), and by immunostaining at E12.5 (C). (D-I) Analysis of proliferation and cell death in E12.5 WT and FoxG1Cre;Gli2 conditional mutants. (J) Schematics of sub-divisions of the pituitary. (K,L) Quantification of BrdU incorporation and the ratio of BrdU/Ki67 in different regions of the pituitary. Scale bars: 100 µm. * represents P<0.05, n=3 embryos.

If proliferation of progenitors in the Av region was affected, then we would

expect to see a reduction in the number of mature pituitary cells generated from this region. We therefore quantified the number of each mature pituitary cell type in FoxG1Cre;Gli2 mutants at E17.5 (Figure 3-5A-J). We found a significantly smaller number of corticotropes (C) and lactotropes (L) (Figure 3-5K), suggesting a requirement of Gli2 in the proliferation and differentiation of their precursors.

The reduction in the number of pituitary cells could impede normal growth during the early postnatal period. To test whether this is the case, we monitored the growth of FoxG1Cre;Gli2 mutant pups. Compared with WT pups, mutant pups had a lower body weight (Figure 3-5L), and even though mutant pups were born alive, most died within 30 days after birth. The growth defects in

FoxG1Cre;Gli2 mutants likely reflect disruption of pituitary function, although it might also result from loss of Gli2 function in endodermal tissues.

We also used Pitx2Cre, which is active in RP [123] but slightly later than

FoxG1Cre, to disrupt Gli2 function. Pitx2Cre mediated recombination of Rosa26-

lacZ can be detected in a few cells at E9.5 and more cells at E10.5 and E12.5

(Figure 3-6A-C). However, when Pitx2Cre was used to remove Gli2 function, we

found no obvious defects in patterning or proliferation of mutant pituitaries at

E12.5 (data not shown), suggesting that Gli2 is only required before Pitx2Cre is

active in RP.

3.4.4 Overlapping roles of Gli genes in pituitary development

Previous studies have shown that the three Gli genes share overlapping and

redundant functions [51]. To address whether Gli2 is the primarily Gli

transcription factor to mediate Shh signaling during pituitary development, we

analyzed the pituitary phenotypes of Gli1 and Gli3 mutants at E12.5. Both

mutants had normal pituitary patterning and proliferation at E12.5 (data not

shown), consistent with the notion that Gli2 is the primary Gli transcription factor

in the pituitary. We next examined the pituitary development in Gli2/Gli3 double

mutants, which do not express Gli1 and therefore lose all Gli function [51], and

found no pituitary in all double mutants examined (n=4). One potential reason for

the complete loss of pituitary in Gli2/Gli3 double mutants is that Gli mediated Shh

signaling is required in the diencephalon for the initial formation of RP. To

address this issue, we generated conditional mutant embryos that lost all Gli

function specifically in the pituitary using FoxG1Cre (FoxG1Cre; Gli2flox/zfd;Gli3xt/xt).

At E12.5, the pituitary gland in these conditional mutant embryos was smaller.

However, when pituitary specific cell markers were used to examine the initial patterning, we found that Lhx3 is expressed normally throughout the pituitary gland (data not shown). In addition, Isl1 showed typical staining in the rostral tip and Pax6 appeared to express in a dorsal-to-ventral pattern, similar to WT embryos at this stage (Appendix II, Figure S1K,L). These results confirm that early pituitary patterning is not affected by loss of Gli function, and that unregulated expression of Gli repressor in pitx1-Hhip transgenic embryos may be responsible for the severe pituitary phenotypes.

Figure 3-5 Early loss of Gli2 function results in the reduction of two groups of pituitary cell types at E17.5. (A-J) Immunostaining of different pituitary cell type markers. (K) Quantification of different pituitary cell types in WT and FoxG1Cre;Gli2 mutants (* denotes P<0.05, ** denotes P<0.01, n=6 embryos). (L) FoxG1Cre;Gli2 mutants did not grow as well as WT pups after birth. Seven pups from four litters were tracked for each genotype. The number on each dot indicates surviving pups at that time. Scale bars: 100 µm.

Figure 3-6 Activation of Hh signaling after the closure of RP resulted in over proliferation of the pituitary. (A-C) X-gal staining revealed the activity of Pitx2Cre from E9.5 to E12.5. (D-K) Analysis of patterning markers in E12.5 WT embryos (D-G) and in embryos with activated Hh signaling (Pitx2Cre;SmoM2) (H-K) using RNA in situ hybridization (D-F,H-J) and immunostaining (G, K). (L, M) Immunostaining of BrdU incorporation and the expression of Ki67. (N) Quantification of BrdU incorporation and Ki67 expression. (* indicates P<0.05, n=6 embryos). (O-V) Activation of Hh signaling did not affect pituitary patterning, as revealed by RNA in situ hybridization of markers at E14.5. Scale bars: 100 µm.

3.4.5 Activation of Hh signaling in RP increases pituitary proliferation

without affecting patterning

If disruption of Gli function affects pituitary cell proliferation, then we

expect that activation of Shh signaling increase pituitary proliferation. To test this,

we activated Hedgehog (Hh) signaling using Pitx2Cre and a conditional allele

encoding an active form of Smo inserted in the Rosa26 locus (R26SmoM2) [118].

At E12.5, Pitx2Cre; R26SmoM2 mutant pouch appeared similar to wild type pouches, as Lhx3, Prop1, Gata2 and Isl1 were expressed normally (Figure 3-6D-

K). The expression pattern of these markers remained unchanged even when

examined at E14.5 (Figure 3-6O-V), suggesting that activation of Hh signaling using Pitx2Cre does not alter the general patterning of the pituitary.

We next examined markers for cell proliferation in mutant pituitary pouches. Although there were no significant differences in the number of Ki67+ and pHH3+ cells in the whole pituitary pouch (data not shown), the number of proliferating cells (BrdU+ and BrdU/Ki67) was significantly increased in the Ad

region of the pouch (Figure 3-6L,M,N). The observation that cells in the dorsal anterior region over proliferate in response to Shh signaling suggests that these cells may normally respond to Shh derived from the diencephalon. The

phenotype became more prominent at E14.5, as Pitx2Cre;SmoM2 mutant

pituitaries were much bigger in size than WT controls (Figure 3-6O-V). Together, these data support our observation that the primary role of Shh/Gli2 signaling is to control the proliferation, rather than patterning, of pituitary progenitors.

3.4.6 Patterning of pituitary requires diencephalic function of Gli2

In addition to the partial loss of anterior pituitary as reported previously

[42], we found that all Gli2 mutants had no posterior pituitary (n=43). The

complete loss of the posterior pituitary suggests a requirement of Gli2 in ventral

diencephalon. As diencephalon-derived signals such as BMP4 and FGF8 are

required for induction and proliferation of the anterior pituitary [17, 19, 33], we were curious whether the expression of these two signals was perturbed in Gli2 mutants. We therefore examined the expression of Shh, Bmp4 and Fgf8 in Gli2

mutants using in situ hybridization on E9.5 embryos and on embryo sections

(Figure 3-7). We found a normal expression of Shh in the ventral diencephalon of

Gli2 mutants (Figure 3-7A,B). However, while Bmp4 was detected strongly in the

medial ventral diencephalon in WT embryos (Figure 3-7E), no Bmp4 expression

was detected in the ventral diencephalon of Gli2 mutants (Figure 3-7F, n=3

embryos). Similarly, while Fgf8 is strongly expressed in the diencephalon of WT

embryos (Figure 3-7I), the expression in Gli2 mutants was greatly reduced (Fig.

7J, n=3 embryos). Similar results were obtained when in situ hybridization was

performed on E9.5 embryo sections (Figure 3-7C,D,G,H,K,L, n=7 embryos).

Together, these results suggest that Gli2 is required for diencephalic expression

of Bmp4 and Fgf8 and that the loss of expression of these two genes in

diencephalon may be responsible for the variable pituitary phenotypes in Gli2

mutants.

Figure 3-7 Gli2 is required for the expression of Bmp4 and Fgf8 in the ventral diencephalon. (A-L) RNA in situ hybridization of Shh, Bmp4 and Fgf8 in E9.5 embryos (A,B,E,F,I,J) and embryo sections (C,D,G,H,K,L). Red arrows indicate diencephalic expression. Scale bars: 100 µm.

3.5 Discussion

Shh signaling is primarily required for proliferation, rather than

specification, of early pituitary progenitors

Shh signaling has been shown to be involved in controlling cell fate specification

and proliferation in different tissues. In the spinal cord, Shh can induce different

types of spinal cord neurons based on the concentration of Shh or the length of

action [115]. In the cerebellum, however, the major function of Shh appears to control proliferation of granular precursors [125]. Analysis of Gli mutants and gain-of-function embryos provided further support that Gli transcription factors, in particular Gli2, mediate different functions of the Shh signaling in different tissues.

For example, Gli2 mutants do not generate floor plate cells and have only residual V3 interneuron in the spinal cord [46, 47]. Similarly, Gli2 mutants have defects in the generation of ventral telencephalic neurons [112]. On the other hand, Gli2 mutants show reduced proliferation of granular progenitors but no loss of cell types in the cerebellum [120].

Because Shh mutants have severe patterning defects during early embryonic development, the role of Shh signaling in pituitary development has previously only been examined in gain- or loss-of-function transgenic embryos

[31]. Blocking Shh signaling by expressing Shh antagonist Hhip in the oral ectoderm caused loss of anterior pituitary markers Gata2 and brn4 (Pou3f4), and a smaller pituitary. On the other hand, ectopic expression of Shh in the anterior pituitary resulted in the expansion of anterior marker Gata2, increased production of pituitary cells and a bigger pituitary. In both situations, changes in the

expression of anterior markers were accompanied by a corresponding change in cell proliferation. Our current study provides several lines of evidence that

Shh/Gli signaling functions primarily to promote pituitary cell proliferation. First, the patterning of Gli2 mutant pituitaries is largely normal but the pituitaries have a clear proliferation defect. Second, Gli2 mutant cells are less likely to contribute to the pituitary, in particular to anterior pituitary, because these mutant cells do not proliferate as well as WT cells in mosaic embryos. Lastly, patterning of the early pituitary is normal in conditional Gli2 mutants when Gli2 function is disrupted by using FoxG1Cre or Pitx2-Cre. Even when both Gli2 and Gli3 are disrupted in the pituitary, cells in RP still express appropriate pituitary markers at E12.5.

Our analysis also revealed a critical window of Shh/Gli signaling in controlling proliferation and differentiation of pituitary progenitors. Removal of

Gli2 using FoxG1Cre affects proliferation and differentiation of several pituitary progenitors. These effects are direct results of loss of Gli2 function in RP, as diencephalic expression of Bmp4 and Fgf8 is not altered in these embryos

(Appendix II, Figure S2). Additional removal of all Gli function in the pituitary have a more drastic effect, as FoxG1Cre;Gli2;Gli3 mutant pituitary glands are much smaller than FoxG1Cre;Gli2 pituitary glands. However, removal of Gli2 function using Pitx2Cre, which becomes active slightly later that FoxG1Cre, does not appear to have obvious effect on patterning or proliferation of the pituitary progenitors. Nevertheless, ectopic activation of Shh signaling using Pitx2Cre causes massive over-proliferation of all pituitary progenitors of the pituitary, suggesting that although Shh/Gli signaling is not required at this stage, pituitary

progenitors are still responsive to ectopic Shh signaling. It further suggests that

Shh signaling must be tightly regulated even at this later phase.

Although Hedgehog signaling is largely conserved across species, clear differences exist as to how Hh/Gli signaling regulates pituitary development in mouse and fish. In mouse, Gli2 functions as the major activator and is required for pituitary development [42], while Gli1 and Gli3 are dispensable, although Gli1 has redundant function with Gli2. Furthermore, conditional removal of Gli2 or all

Gli function in pituitary reveals that Hh/Gli signaling controls proliferation, rather than patterning, or early pituitary progenitors. Our analysis also reveals that

Hh/Gli signaling controls diencephalic expression of Bmp4 and Fgf8, which have been shown to control early pituitary patterning and proliferation. On the other hand, zebrafish dtr/gli1 is required for normal development as dtr/gli1 null mutants have a pituitary phenotype including a reduced expression of pituitary markers nk2.2 and Lim3, and reduction in the number of somatotropes, lactotropes and anterior corticotropes [126, 127]. Blocking the function of yot/gli2 results in minor pituitary phenotypes as the number of corticotropes is reduced.

However, blocking the function of both dtr/gli1 and yot/gli2ab function results in a significant reduction in almost all anterior pituitary cell types [127], a phenotype that is more severe than any single or compound mouse Gli mutants. It is possible that the differences observed are species specific, as the morphogenetic process of pituitary formation in zebrafish is distinct from that of the mouse or chick in at least two aspects. First, the zebrafish pituitary primordium is formed as a solid structure at the anterior edge of the head, which

then moves caudally into the head without a pituitary-specific morphogenetic movement similar to that in chick or mouse. Second, the formation of zebrafish pituitary gland does not involve a structure equivalent to Rathke’s pouch and the invagination of oral/placodal ectoderm [71, 128]. In addition, there are differences in the transcriptional regulation of Gli genes in mouse and fish, as mouse Gli2 is not regulated by Hh signaling while the zebrafish dtr/gli1 appears to be regulated by Hh signaling. Alternatively, the more severe pituitary phenotype in zebrafish could also result from morpholino knockdown of gene function in both pituitary and diencephalon.

Shh/Gli2 mediated diencephalic signal is required for pituitary development

Our analysis of pituitary-specific Gli mutants revealed that blocking of Shh signaling within pituitary progenitors reduces proliferation and differentiation of pituitary progenitors but does not significantly affect patterning of the pituitary gland. But if that is the case, why would half of the Gli2 mutants lack a pituitary?

The answer appears to be in the ventral diencephalon. Starting from E8.0, Shh is strongly expressed in the ventral diencephalon, immediately above the forming

RP. Our previous analysis identified loss of ventral diencephalic tissues and reduced expression of Nkx2.1 in some Gli2 mutants. As ventral diencephalon is an important tissue in the induction of RP, we examined the expression of diencephalic marker genes between E8.5 and E9.5. We found that Gli2 mutants had reduced expression of Bmp4 and Fgf8, which are growth factors shown to be critical for the development of a definitive pouch [17, 33]. Our study therefore

identified two requirements for Shh/Gli signaling that are critical for pituitary progenitor development: one within RP to promote proliferation of pituitary progenitors and a second role to control expression of Bmp4/Fgf8 in the ventral diencephalon to control early patterning of RP. Future work will be needed to determine whether the Gli2 transcription factor can bind directly to Bmp4/Fgf8 regulatory elements to control their expression in the ventral diencephalon.

3.6 Acknowledgements

We thank Drs. M. Rosenfeld and S. Camper for providing in situ probes,

W. Jiang of CWRU transgenic facility for performing the embryo aggregation experiments, Dr. P. Conrad for microscope assistance and Drs. R. Atit, K.

Molyneaux and R. Conlon for comments and suggestions. This study was supported in part by a CWRU fund (CBB) and an NIH R01 HL093484 (JFM).

Chapter 4

Patterning of the ventral telencephalon requires

positive function of Gli transcription factors

This Chapter was published as Yu, W., Wang,Y., et al. (2009). "Patterning of

ventral telencephalon requires positive function of Gli transcription

factors." Dev Biol 334(1): 264-275 (see Appendix III for copyright license of

reusing this article in current thesis). I contributed to Figure 5-6 in this

chapter and supplementary Figure 1 in appendix IV.

4.1 Abstract

Three Gli transcription factors in neural precursors. However, whether Gli- mediated Shh signaling is also required to induce different cell types in the ventral telencephalon has been controversial. In particular, loss of Shh has little effect on dorsoventral patterning of the telencephalon when Gli3 is also removed.

Furthermore, no ventral telencephalic phenotypes have been found in individual

Gli mutants. To address this issue, we ﬁrst characterized Shh-responding ventral telencephalic progenitors between E9.5 and E12.5 and found that they produce neurons migrating to different layers of the cortex. We also discovered a loss of

Nkx2.1 and Nkx6.2 expression in two subgroups of progenitors in embryos lacking major Gli activators. Finally, we analyzed the telencephalic phenotypes of embryos lacking all Gli genes and found that the ventral telencephalon was

highly disorganized with intermingling of distinct neuronal cell types. Together,

these studies unravel a role for Gli transcription factors in mediating Shh

signaling to control speciﬁcation, differentiation and positioning of ventral

telencephalic neurons.

4.2 Introduction

The ventral telencephalon is the source of several types of neurons,

including striatal, olfactory and cortical interneurons and is comprised of three

proliferative zones: the lateral, medial and caudal ganglionic eminences (LGE,

MGE and CGE). Following their generation, many MGE-and LGE-derived neurons undergo tangential migration to reach the cortex or olfactory bulb [129-

131]. Recent studies have further shown that the migration pattern and positioning of the transplanted cells are largely determined by donor tissues, suggesting that local signals within the ventral telencephalon determine cell fate and behavior of telencephalic neurons [83, 88, 90-92, 132].

Shh, a member of the Hedgehog (Hh) family of secreted proteins that specify many central nervous system (CNS) cell types[115], is expressed in early telencephalic tissues. The initial Shh expression appears in the prechordal plate

[93]. By E9, a number of morphogenetic events transform the telencephalic anlage into a set of paired vesicles and Shh is expressed in the neural epithelium of the ventral telencephalon [94]. By E12, Shh expression is shifted into the mantle area, and can no longer be detected in the neuroepithelium of the MGE

[95]. Both genetic and gain-of-function studies conﬁrm the importance of Shh signaling in telencephalon development. Speci ﬁcally, mutations in Shh or

conditional loss of components of the Shh pathway at an early stage (~E8.5)

using FoxG1-Cre lead to the loss of all ventral telencephalic tissues [66, 103,

133], suggesting an early requirement for Shh in the dorsoventral patterning of

the telencephalon. On the other hand, disruption of Shh signaling pathway

between E10 and E12 using Nestin-Cre has a more limited effect, as it only affects the dorsal-most expression of Nkx2.1 [104-106]. Conversely, ectopic dorsal expression of Shh leads to the induction of ventral telencephalic markers and repression of dorsal markers [97, 101, 124, 134, 135]. Shh signaling, therefore, could be a local signal linking progenitor cell speci ﬁcation to the fate of the migrating post-mitotic ventral neurons.

Although Shh is clearly required for telencephalic development, it is less clear whether and how Shh is involved in specifying different cell types in the ventral telencephalon. Shh signaling is mediated by Gli transcription factors, which include Gli1, Gli2 and Gli3 [37, 136, 137]. Both Gli1 and Gli2 function primarily as transcriptional activator, as revealed by biochemical analysis and in vivo mouse knock-in genetic analysis [38, 44, 45], and Gli3 functions primarily as a transcriptional repressor [48, 49]. However, examination of the MGE marker

Nkx2.1 expression in mice mutated for individual Gli genes or of Gli1/2 double mutants has not revealed obvious defects in patterning of ventral telencephalon

[42]. On the other hand, Gli3 mutant mice show a strong dorsal telencephalic phenotype [98-102], but ventral patterning appears to be largely normal. These analyses suggest that Gli3 repressor is required in the dorsal telencephalon.

Interestingly, removal of Gli3 from Shh mutant embryos largely rescues dorsoventral patterning defects in Shh mutants [101], suggesting that the primary function of Shh signaling is to prevent the production of excessive Gli3 repressor.

However, because Gli3 has also been shown to function as a weak activator in vivo [51, 138], it remains to be determined whether there is any Gli activator

function in the ventral telencephalon, and whether Gli genes have a

compensatory function in mediating Shh signaling to in ﬂuence speciﬁcation and

behavior of neurons derived from ventral telencephalon.

To address whether Gli activators are required for patterning of ventral

telencephalon, we analyzed an array of domain-speciﬁc ventral telencephalic

markers in Gli2 and Gli1/2 double mutants, and identiﬁed the loss of Nkx2.1 and

Nkx6.2 expression in two subgroups of progenitors in the interganglionic sulcus.

We also analyzed the ventral telencephalic phenotypes of pan-Gli mutant

embryos and showed that Gli genes play multiple roles in telencephalon

development, including speciﬁcation, differentiation and positioning of ventral

telencephalic neurons.

4.3 Materials and methods

Mouse breeding

For fate-mapping experiments, Gli1-CreER/+;R26R/R26R [107] male mice

were bredwith6–8-week-old CD-1 females (Charles River). Noon of the day of

detection of a vaginal plug is considered E0.5. Tamoxifen (Sigma, T-5648) was dissolved in corn oil at a concentration of 20 mg/ml and was fed to pregnant female using a gavage-feeding needle (FST) at a dose of 4 mg/40 g body weight, at 5 PM on different gestational days. The genotyping of Gli1-lacZ, Gli2-lacZ,

zfd xt Gli2 and Gli3 mice were as described [44, 109, 121]. Double heterozygous

mutant mice were maintained on an outbred background.

Histology, immunohistochemistry and RNA in situ hybridization

Mouse embryos were dissected in cold PBS and ﬁxed in 4%

paraformaldehyde (PFA) for 1 h at 4°C. Adult mouse brains (8-10 weeks old)

were dissected out after transcardiac perfusion and post-ﬁxed with 4% PFA for 3

h. Embryos and brains were washed with PBS, cryo-protected in 30% sucrose and embedded in OCT (Tissue-Tek). Tissues were sectioned at 12 μm (embryos) or 20 μm (adult brain) on a Leica Cryostat. At least three brains were analyzed for each time point. Immunohistochemistry, X-gal staining, H&E staining and

RNA in situ hybridization were performed essentially as described [44, 110].

Subtype identity of labeled cells was determined by double immunoﬂuorescence staining using antibodies against β-galactosidase and parvalbumin, somatostatin or calretinin. Antibodies used were: GABA (1:2000, Sigma), calretinin (1:2000,

Chemicon), parvalbumin (1:2000, Sigma), somatostatin (1:250, Chemicon), β- galactosidase (1:500, Biogenesis; or 1:500, 5 prime-N3 prime), Pax6 (1:20,

Developmental Studies Hybridoma Bank), Gsh1/2 (1:200, Kenneth Campbell,

Cincinnati Children's Hospital Medical Center, Cincinnati, OH, USA), BrdU (1:20,

Becton-Dickinson), Ki67 (1:500, Abcam), phospho-Histone H3 (1:100, Upstate), cleaved Caspase-3 (1:200, Cell Signaling), and Nkx2.1 (TTF1, 1:500, Epitomics).

Nuclear counter-staining was performed with Hoechst 33258 (Molecular Probes) or with 0.005% nuclear fast red (Polyscientific, Inc.). Probes for in situ hybridization have been described previously [101]. Sections were examined with a Leica DMLB epifluorescence microscope fitted with a SPOT camera in the

Genetics Imaging Facility (supported by NIH-NCRR, RR-021228).

Fate-mapping analysis

Coronal sections of the brain were stained by nuclear fast red or Hoechst nuclear dye to reveal layer structures. To better quantify the position of labeled cells in the cortex, we divided the cortex into ten bins along the radial axis, with bin 1 closest to the white matter (WM) and bin 10 next to the pial surface. Only those cells located within primary motor cortex and somatosensory cortexes were used for quantiﬁcation (Paxinos and Franklin, The Mouse Brain). In order to map the location of the labeled cells, X-gal stained cells were assigned to one of the ten bins according to their location in the cortex. Three sections from one animal, located about 0.12 mm apart, were used for quantiﬁcation. Three mouse brains were used for each time point. The numbers on the histograms represent the mean percentage of labeled cells in one bin over the total number of β- galactosidase labeled cells in the section ± S.E.M.

BrdU labeling, cell cycle and cell death analyses

E12.5 pregnant mice were injected intraperitoneally with BrdU (Sigma) at

100μg/g body weight 1h prior to dissection. The fraction of BrdU+ cells was calculated by counting the number of those positive cells in 75 μm wide areas spanning radially from the ventricular surface to the edge of VZ/SVZ (as judged by dense nuclear staining) and dividing the total number of nuclei in that area

(each segment contains ~250 nuclei). To calculate the fraction of S-phase cells, the number of BrdU+ cells was divided by the total number of Ki67+ cells. To

calculate the fraction of pH3+ cells, the number of those positive cells bound by a box (120 μm× 94 μm) in the VZ/SVZ was divided by the total number of nuclei.

For cell death analysis, the total number of Caspase3+ cells in the MGE or LGE was divided by the size of the area and converted to a percentage. Data from 3–

5 embryos from at least two different litters were pooled together to determine average and S.E.M for each genotype. Student's t-test was used to calculate the p value and to determine whether the results were signi ﬁcantly different from each other.

4.4 Results

4.4.1 A subset of ventral telencephalic progenitors receives Hh signaling and expresses Gli1-lacZ

In order to visualize those cells that respond to Hh signaling, we examined the expression pattern of Gli1-lacZ, in which a lacZ gene is expressed from the

Gli1 genomic locus and thus provides a sensitive readout of Hh activator signaling [110]. At E12.5, Gli1-lacZ was found in the interganglionic sulcus, with a higher level of expression on the MGE side (Figure 4-1A, B), similar to previous reports [102, 139]. The expression pattern suggests that cells in the interganglionic sulcus, in particular those on the side of the MGE, receive a high level of Hh signaling. We next examined the expression pattern of Gli2-lacZ, where a lacZ gene is expressed from the Gli2 genomic locus [44], and found that the ventral limit of Gli2-lacZ expression coincided with that of Gli1-lacZ on the

MGE side of the interganglionic sulcus (Figure 4-1C, D). The expression of these

Hh-dependent transcriptional activators in the ventral telencephalon raises the

intriguing possibility that different ventral telencephalic progenitors are generated

in response to Hh signaling, as they are in the spinal cord.

Figure 4-1 X-gal staining of Gli1-lacZ and Gli2-lacZ on telencephalic sections of E12.5 mouse embryos. (B) and (D) are higher magnification images of the boxed regions in (A) and (C) respectively. Arrows indicate the ventral expression limits of Gli1 and Gli2. Note the overlapping ventral expression limit of Gli1 and Gli2 in the ventral telencephalon. LGE, lateral ganglionic eminence; MGE, medial ganglionic eminence. Scale bar: 0.15mm (A, C).

4.4.2 Hh-responding progenitors produce progressively superficial cortical

interneurons

To determine whether different cell types are generated from Hh- responding progenitors between E9.5 and E12.5, we fate mapped neurons produced from these Gli1-expressing progenitors using an inducible genetic

approach [140], and examined cell fate 8 weeks after birth when telencephalic neurons have completed their migration and express mature cell type markers.

This strategy involves using Gli1-CreER, in which an inducible Cre is inserted

into the endogenous Gli1 locus [107], and a Rosa26 reporter (R26R) [116]. A

single application of Tamoxifen (TM) allows for activation of Cre, recombination

of the reporter allele and the labeling of distinct populations of Gli1+ cells during

a ~24h time window beginning about 6h after TM administration. Since

dorsoventral patterning of the telencephalon has already been established by

E12.5 [104-106], we sought to determine the inﬂuence of Hh signaling on

telencephalic progenitors expressing Gli1 prior to E12.5.

We ﬁrst examined the expression of β-galactosidase ~36h after TM

delivery to determine the initial descendants of cells expressing Gli1 (Figure 4-2A,

B). Very few β-galactosidase+ cells were found in the telencephalon when TM

was given at 5 PM on E7.5 (data not shown). However, when TM was given on

E8.5 or later, many β-galactosidase+ cells were detected in the ventricular layer

of the dorsal MGE 36 h later (Figure 3-2C). In particular, β-galactosidase+ cells

labeled prior to E11.5 occupy a broader domain in the MGE than cells labeled on

E11.5, as assessed by co-staining of Nkx2.1 (Appendix IV Figure S1),

suggesting a mechanism that reﬁnes Hh-responding cells to the dorsal MGE

during this time period. When analyzed at 2 months after birth, many β-

galactosidase+ cells were found in the striatum, ventral striatum, ventral septum

and the cortex (Figure 4-2D). Because most of the cortical interneurons are

known to derive from the ventral telencephalon, in particular from the MGE,

Figure 4-2 Different cortical interneurons were generated from progenitors responding to Hh signaling from ~E9.5 to ~E12.5. (A) Schematic of Tamoxifen (TM) injection to analyze the initial populations of Hh-responding progenitors in the ventral telencephalon. TM was given at 5 PM on E8.5 to E11.5 to label Hh-responding cells from ~E9.5 to ~E12.5. (B) Coronal section of an E11.5 telencephalon showing the MGE and LGE. Red dotted box indicates the area shown in (C). (C) Immuno ﬂuorescence staining of β- galactosidase to reveal the initial population of cells responding to Hh signaling. Embryos were analyzed ~36 h after TM injection. White dotted lines indicate the edge of the ventral ganglionic eminence. (D) Coronal section of a 2-month-old brain section stained with X-gal to reveal the labeled cells. Cells expressing Gli1 at ~E9.5 were permanently labeled with β-galactosidase from the Rosa26 reporter. Labeled cells were found in somatosensory cortex (a, b), striatum (c), piriform cortex (Pir) (d) and septum. (E) Layer positioning of cortical interneurons correlates with the timing of exposure to Hh signaling. TM was given on E8.5, E9.5 and E11.5 to label progenitors responding to Hh signaling at ~E9.5, ~E10.5 and ~E12.5. The position of labeled cells in 2-month-old cortex was determined by X-gal staining, followed by nuclear fast red staining to reveal the different layer structure of the cortex (I to VI). Only labeled cells in the somatosensory and motor cortexes were counted. To facilitate calculation, the cortex was divided into 10 bins along the radial axis, with bin 1 closest to the white matter (WM) and bin 10 closest to the pial surface. Arrows indicate X-gal stained cells. (F) Layer distribution of labeled cells derived from progenitors responding to Shh signaling at different developmental stages. X-gal stained cells were placed into each bin and plotted in the graph. The approximate layer numbers are also indicated. The numbers are expressed as mean percentage±S.E.M.Three coronalsections from eachbrainwere examined,and threebrains were usedforcalculation.Total numbers oflabel cellanalyzed are: n=106 (TM E8.5), n =215 (TM E9.5) and n =324 (TM E11.5). Scale bar: 0.1 mm (D).

we focused our analysis on these β-galactosidase+ cells in the somatosensory

and motor cortexes of 8-week-old adult mouse brains.

To quantify the contribution of labeled cells to different cortical layers, we stained coronal sections of the brain with nuclear fast red to reveal the laminar structure, divided the radial axis of the cortex into ten bins along the pial surface and the white matter (WM), and assigned each labeled cell to one of these bins

(Figure 4-2E), as previously described [92, 141, 142]. We found that the majority of labeled cells derived from ~E9.5 Hh-responding progenitors (TM injection at 5

PM of E8.5) were located close to the white matter (Figure 4-2E), with about 88% of the progeny located within the inner half (bins 1–5, approximately layers V and

VI) of the cortex (total labeled cells n =106, 3 adult brains) (Figure 4-2F). In

contrast, when progeny derived from ~E10.5 Hh-responding progenitors were

analyzed, ~68% of the cells (n = 215) occupied intermediate locations (bins 3–6,

approximately layers V and IV) in the cortex. Lastly, we found ~72% of labeled

cells (n = 324) derived from ~E12.5 Hh-responding cells were located near the

pial surface of the cortex (bins 6–9, or approximately layers IV, III and II). Our

results thus show an ‘inside-out’ pattern of ventrally derived cortical interneurons

derived from Gli1 expressing progenitors.

To determine whether different subpopulations of cortical inter-neurons

are generated at different stages, we examined the expression of three cortical

interneuron markers, parvalbumin (PV), somatostatin (SST) and calretinin (CR)

[83, 87, 90, 92, 143] within the labeled cells (Appendix IV Figure S2A). We found

similar numbers of PV+ neurons were generated from progenitors responding to

Hh signaling from E9.5 to E11.5. In contrast, there was an increase in the

number of SST+ and CR+ neurons generated from progenitors responding to Hh

signaling from E9.5 to E11.5 (Appendix IV Figure S2B). Collectively, these analyses suggest that the temporal response of telencephalic progenitors to Shh, mediated primarily by Gli2, correlates with the generation of different waves of telencephalic neurons.

4.4.3 Gli activators are required for the specification of two progenitor groups in the telencephalic sulcus

Previous studies have shown that Gli activator is required for speci ﬁcation of different spinal cord progenitors but not for generation of different digits in the limb [44, 46, 47, 107]. To determine whether Gli activator is required for speciﬁcation of telencephalic progenitors, we analyzed the expression of critical transcription factors in E12.5 embryos lacking different Gli activators.

We ﬁrst examined the expression of Nkx2.1, which normally marks the entire extent of the MGE, up to the point of the interganglionic sulcus (Figure 4-

3A, A’). In Gli1 mutant embryos, the expression of Nkx2.1 remained throughout

the MGE, similar to wild-type (WT) embryos (Figure 4-3B, B’). In contrast, in Gli2

mutant embryos, although Nkx2.1 was still expressed strongly in the MGE, the

dorsal-most expression of Nkx2.1 in the interganglionic sulcus was lost (Figure 4-

3C, C’, arrow). Strikingly, the region where Nkx2.1 expression was lost overlaps

with the ventral expression of Gli1 on the MGE side of the sulcus (Figure 4-1B),

suggesting that cells normally receiving the highest level of Shh signaling are

Figure 4-3 Gli activator is required for the development of subgroups of ventral telencephalic progenitors. (A–D’) The expression of Nkx2.1 was affected in Gli2 and Gli1/2 double mutants. In WT and Gli1 mutant, Nkx2.1 was expressed in the interganglionic region (A, A’,B, B’). However, in Gli2 or Gli1/2 mutants, the dorsal expression of Nkx2.1 was lost (C, C’, D, D’, indicated by white arrows). (E–H’) Loss of Nkx6.2 expression in Gli2 and Gli1/2 mutants. Nkx6.2 was expressed throughout the interganglionic region in WT and Gli1 mutants (E, E’,F,F’). However, the expression of Nkx6.2 was lost in Gli2 and Gli1/2 mutants (G, G’,H,H’). The overall production of Lhx6+ MGE neurons (I–L) and GAD67+ GABAergic neurons (M–P) did not appear to be dramatically altered. By E14.5, the reduction in the dorsal-most Nkx2.1 domain (Q, Q’, yellow arrowheads) persisted in Gli2 mutants. Furthermore, the leading edge of migrating Lhx6+ neurons was reduced (R, R’, red arrowheads). The expression of Ebf1, which labels LGE-derived neurons, did not appear to be altered in Gli2 mutants (S, S’). By E18.5, all Gli2 mutants (4/4) showed enlarged telencephalic ventricles, reduced numbers of Lhx6+ cortical interneurons (T, T’) and tyrosine hydroxylase (TH) ﬁbers (V, V’, white arrowheads). The Ebf1+ striatum (Str) was largely normal (U, U’). NCX, neocortex; GP, globus pallidus. Scale bar: 75 μm(A–P).

either lost or transformed into a more dorsal identity. To further study the shift of the dorsal Nkx2.1 border, we examined the expression of Nkx6.2, which encodes a homeobox transcription factor speciﬁcally expressed in the inter-ganglionic sulcus between the MGE and LGE throughout prenatal stages and is dependent on Shh signaling [106, 144]. In Gli1 mutants, like in WT embryos, Nkx6.2 appeared to be expressed at the highest level in the sulcus, with a decreasing gradient towards the MGE and LGE (Figure 4-3E, E’,F,F’). However, in Gli2 and

Gli1/2 double mutant embryos, the expression of Nkx6.2 was lost in the sulcus

(Figure 4-3G, G’,H, H’). The loss of Nkx6.2 expression further conﬁrms that when

Shh signaling is disrupted in Gli2 mutants, or in Gli1/2 double mutants, speciﬁcation of Nkx6.2+ telencephalic progenitors is affected. The analyses of these Gli mutant embryos raise the possibility that Gli activator is required to

specify two subgroups of telencephalic progenitors: Nkx2.1+;Nkx6.2+;Gli1high progenitors located on the MGE side of the interganglionic sulcus and

Nkx2.1−;Nkx6.2+;Gli1low progenitors located on the LGE side of the sulcus (see summary in Figure 4-8).

To determine whether the overall production of post-mitotic neurons is affected in the ventral telencephalon of different Gli mutants, we examined two additional neuronal markers at E12.5. Lhx6 encodes a LIM homeodomain transcription factor that is expressed in all cells derived from the MGE [145]. In

Gli1, Gli2 or Gli1/2 double mutants, the expression of Lhx6 appeared unchanged

(Figure 4-3I–L). We next examined the expression of GAD67, which encodes a rate-limiting enzyme in the synthesis of GABA, and is expressed in all ventral

telencephalic-derived inhibitory GABAergic neurons. In Gli1, Gli2 or Gli1/2 mutants, GAD67 was detected throughout the mantle area surrounding the MGE and the LGE, similar to that in WT embryos (Figure 4-3M-P). The similar expression of these markers in Gli mutant and WT embryos shows that the loss of two subgroups of telencephalic progenitors does not dramatically alter the overall production of interneurons at E12.5.

By E14.5, we found that one out of four Gli2 mutant brains had enlarged ventricles (Appendix IV Figure S3). Even in those Gli2 mutant embryos with normal morphology, the reduction in the dorsal Nkx2.1 domain was apparent

(Figure 4-3Q, Q’). Furthermore, the leading population of Lhx6+ neurons migrating towards the cortex was reduced (Figure 4-3R, R’), consistent with a requirement of Nkx2.1 for the expression of Lhx6 [146]. On the other hand, Ebf1+ population did not appear to be affected (Figure 4-3S, S’). By E18.5, however, all four Gli2 mutant brains were found to have enlarged ventricles, as previously noted [147], and reduced ventral telencephalon (Appendix IV Figure S3). Other defects were also noted, including a reduction in Lhx6+ interneurons in the cortex

(Figure 4-3T, T’) and reduced innervation of tyrosine hydroxylase ﬁbers in the mutant ventral telencephalon (Figure 4-3V, V’). The Ebf1+ striatum, however, does not appear to be signiﬁcantly reduced (Figure 4-3U, U’).

4.4.4 Removal of all Gli genes disrupts production and proliferation of ventral telencephalic neurons

The three Gli genes are expressed in an overlapping manner in many

embryonic tissues. Recent studies have shown that in addition to Gli1 and Gli2,

Gli3 can also function as an activator in vivo to mediate Hh signaling [51, 138].

To determine whether Gli3 may play a role in ventral telencephalon development, we examined the expression of Gli3 by RNA in situ hybridization. As previously reported [102], Gli3 is strongly expressed in the dorsal telencephalon. In addition,

Gli3 was also detected in the interganglionic sulcus, with a ventral expression limit similar to that of Gli2 (Figure 4-4A, B), suggesting that Gli2 and Gli3 may promote Gli1 expression in the ventral telencephalon. Indeed, we found a weakened Gli1-lacZ expression in the interganglionic sulcus of Gli2 mutants

(Figure 4-4C, D). Furthermore, we found a slight reduction in Gli1-lacZ

expression in the absence of Gli3 (Figure 4-4E), and a complete absence of Gli1-

lacZ expression in Gli2/3 double mutants (see Figure 3-2 in [51]).

To determine whether ventral telencephalon patterning is normal in the

absence of all Gli genes, we examined the expression of progenitor and neuronal

markers in E12.5 Gli2/3 mutant embryos, which lack the expression of all three

Gli genes (hereafter referred to as pan-Gli mutants). We found that Nkx2.1

expression was largely restricted in the MGE, although the dorsal expression

was reduced and somewhat diffuse in the interganglionic sulcus (Figure 4-5A, F,

insert). The restricted expression of Nkx2.1 suggests that speciﬁcation of MGE

progenitors is not greatly altered in Gli2/3 mutants. Next we examined the

expression of Lhx6, which is expressed in MGE-derived post-mitotic neurons,

and found a severe reduction in Gli2/3 mutants (Figure 4-5B, G). To determine

whether the production of LGE-derived post-mitotic neurons is affected, we

examined the expression of Ebf1 and also found a reduction in the number of

Ebf1+ neurons (Figure 4-5C, H). A similar reduction in GAD67, which is expressed in both MGE and LGE post-mitotic neurons, was also found in Gli2/3 mutant embryos (Figure 4-5D, I).

Figure 4-4 Gli2 and Gli3 have overlapping function in regulating Gli1 expression in the ventral telencephalon. (A, B) RNA in situ hybridization showing that Gli3 is expressed in the interganglionic sulcus of E12.5 telencephalon. White arrowhead indicates the approximate area of Gli1 expression. (C–E) Immunoﬂuorescence staining of Gli1-lacZ in the telencephalon of E12.5 WT, Gli2 and Gli3 embryos. Gli1 expression was dramatically reduced in Gli2 mutants (D). In Gli3 mutants, Gli1 expression was reduced but remained restricted in the interganglionic sulcus (E). White lines indicate ganglionic eminences. White bracket lines indicate the expression of Gli1-lacZ. Scale bar: 0.1 mm.

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Figure 4-5 Generation of ventral telencephalic neurons is affected in Gli2/3 double mutants. (A, F) Nkx2.1 expression, which is normal detected in MGE, did not appear to be signiﬁcantly altered in Gli2/3 mutant embryos. Note: the dorsal limit of Nkx2.1 expression was reduced in Gli2/3 double mutants (insert). (B, G) Expression of Lhx6, which marks MGE-derived neurons, was reduced in Gli2/3 mutants. (C, H) Expression of Ebf1, which marks LGE-derived neurons, was also reduced in Gli2/3 mutants. (D, I) GAD67, which is expressed in both the MGE and LGE-derived neurons, was found to expand into the dorsal telencephalon, as indicated by red arrowheads. (E, J) Oligodendrocyte precursors were generated in Gli2/3 double mutants, although in a smaller number in the mutant telencephalon. Red asterisks indicate the PDGFRa+ cells. Note: PDGFRa is also expressed strongly outside of the neural tube. (F, G, I, J) were adjacent sections of the same embryo. Black arrows indicate the dorsoventral limit, as determined by Pax6 expression on an adjacent section. Scale bar: 90 μm.

In addition to the reduced expression of markers for ventral telencephalic

neurons, we also found a reduction in the expression of neuronal marker TuJ1

(Figure 4-6A, E). To address whether disrupted neurogenesis is caused by defects in proliferation of MGE and/or LGE progenitors, we examined BrdU incorporation and Ki67 expression in telencephalic progenitors. BrdU was given to pregnant mice 1 h prior to dissection at E12.5 to label S-phase cells and Ki67 staining was used to reveal all proliferating cells (Figure 4-6B, C, F, G). We found that the fractions of S-phase progenitors in MGE and LGE of Gli2/3 double mutants were signiﬁcantly lower (p b 0.01, Student's t-test) (Figure 4-6I). 101

Moreover, the percentages of proliferating Ki67+ MGE and LGE progenitors were

also signiﬁcantly reduced in Gli2/3 double mutants (p b 0.01) (Figure 4-6K).

However, within the cycling population, there was a signi ﬁcant increase in the

fraction of S-phase cells (BrdU+/KI67+) in both MGE and LGE progenitors

(Figure 4-6J), suggesting a faster cell cycle time. Indeed, the fraction of M phase

cells, indicated by phospho-Histone H3 staining, was also increased (LGE: WT

4.01± 0.13, mutant 5.66±0.41; MGE: WT 4.3±0.3, mutant 5.11±0.36). In addition

to changes in cell proliferation, we also found a signi ﬁcant increase in the number

of Caspase-3+ cells in both MGE and LGE progenitors in Gli2/3 mutants (p b

0.01) (Figure 4-6D, H, L). Together, these results suggest that loss of Gli genes signiﬁcantly affects the proliferation and survival of MGE and LGE progenitors.

The reduced number of post-mitotic neurons in the ventral telencephalon

of Gli2/3 mutants could also be caused either by defects in the differentiation of

neuronal progenitors or by a delay in the generation of neurons. Because most

Gli2/3 double mutants die by E13, we cannot assess whether the number of

post-mitotic neurons is restored at later developmental stages. However, if there

is a developmental delay in Gli2/3 double mutant embryos, then we would expect

an absence of PDGFRα+ oligodendrocyte precursors from the ventral

telencephalon, because these cells are produced in WT embryos around E12.5

[95, 148, 149]. In Gli2/3 double mutants, as in WT embryos (Figure 4-5E),

PDGFRα+ cells were detected in the telencephalic region and outside of the

neural tube (Figure 4-5J), although the number of positive cells was reduced

compared with WT embryos. This result argues against a developmental delay in

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embryos that cannot respond to Shh, but instead suggests that Gli genes control

the fate and differentiation of post-mitotic neurons in the ventral telencephalon.

Figure 4-6 The production of post-mitotic neurons is affected in Gli2/3 double mutant telencephalon. (A, E) Tuj1 staining of coronal sections of WT and Gli2/3 telencephalon. (B, C, F, G) Triple immunofluorescence staining of BrdU (green), Ki67 (red) and nuclei (blue) in the MGE and LGE region of WT and Gli2/3 mutant embryos. (D, H) Immunostaining of cleaved Caspase-3 in WT and mutant embryos. There was a significant reduction in the numbers of BrdU+ cells (I) and Ki67+ cell (K) in Gli2/3 mutants. However, the fraction of BrdU+ cells within proliferating Ki67+ cells was significantly increased in Gli2/3 mutants (J). There was also a significant increase in the number of Caspase-3+ cells in Gli2/3 mutants. **p<0.01; *p<0.05. Scale bar: 0.1 mm (A, E); 40 μm (B, C, F, G); 20 μm (D, H). 103

4.4.5 Intermingling of different neuronal groups in Gli2/3 mutants

In addition to a reduced number of post-mitotic neurons, we noticed that some Lhx6+ neurons occupied a more dorsal position in Gli2/3 mutants (Figure

4-5G). Similarly, some GAD67+ ventral neurons were also found to occupy

ectopic positions in the mutant dorsal telencephalon (Figure 4-5I, red

arrowheads). The ectopic expression of these markers suggests a disruption in

the arrangement of distinct neuronal domains in the mutant telencephalon.

Alternatively, the phenotype could be caused by a diencephalic displacement of

dorsal telencephalic tissues, resulting in a joining of diencephalic and dorsal

telencephalic tissues as has been demonstrated in Gli3 mutants [99]. To

distinguish these two possibilities, we ﬁrst performed RNA in situ hybridization of

FoxG1, a forkhead box containing transcription factor that is normally expressed

speciﬁcally in telencephalic, but not diencephalic tissues [150]. In WT, Gli1, Gli2,

Gli1/2 mutants, FoxG1 is expressed throughout the telencephalon except in the

dorsal cortical hem/choroid plexus anlage region (Figure 4-7A–C, indicated by

red arrowheads). As expected, FoxG1 was not detected in most of the dorsal

telencephalic region of Gli3 mutants (Figure 4-7D), since this region has been

transformed into diencephalic fate [99]. On the other hand, we found that in

Gli2/3 double mutants, FoxG1 was expressed throughout the telencephalon

except in the cortical hem/choroid plexus anlage region (Figure 4-7E). Thus,

unlike Gli3, the dorsal telencephalic region of Gli2/3 mutants retains

telencephalic characteristics.

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Figure 4-7 Dorsoventral patterning in different Gli mutants. (A–E) RNA in situ hybridization of telencephalic marker FoxG1. FoxG1 is normally expressed in telencephalic but not in diencephalic tissues (A). Red arrowheads indicate the expression limit of FoxG1.In Gli2 (B), Gli1/2 (C) and Gli2/3 double mutants (E), FoxG1 was detected throughout the telencephalon, suggesting that the dorsal tissues in these mutants are of telencephalic origin. On the other hand, in Gli3 mutants, FoxG1 was not detected in the dorsal region (D). (F–J) The expanded expression of Dlx2 suggests a dorsal expansion of ventral markers in Gli2/3 double mutants. Arrowheads indicate the dorsal expression limit of Dlx2. (K–O) Immunofluorescence staining of Pax6 and Gsh2 in different Gli mutant telencephalons, with higher magni fication images shown to the right. Normally Pax6 is expressed in the dorsal telencephalon (K, green channel) and Gsh2 is expressed in the ventral telencephalon (K, red channel). White arrows indicate the ventral limit of Pax6 expression. In Gli2 and Gli1/2 double mutants, the expression of Pax6 and Gsh2 was similar to that in WT embryos. In Gli3 mutants, Gsh2 was found to expand dorsally (N), reflecting a partial diencephalic transformation. In Gli2/3 mutants,

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Pax6 remained expressed in the dorsal telencephalon (O, green channel). However, Gsh2 expression expanded into the dorsal telencephalon (O, red channel). (P, R) Expansion of sFRP2 in WT and Gli2/3 mutants. sFRP2 is normally expressed in a small stripe in the corticostriatal area, as indicated by an arrowhead in (P). However, the domain of sFRP2 was expanded in the Gli2/3 mutants (indicated by a bracket in R). (Q, S) Expansion of Isl1+ neurons into the dorsal telencephalon. Isl1+ neurons are normally located ventrally to the sFRP2 domain (white arrowhead in Q). In Gli2/3 mutants, Isl1+ neurons were found to expand into the dorsal telencephalon to overlap with the sFRP2 domain (S, white bracket indicates the sFRP2 domain. R and S are adjacent sections). Scale bar: 0.1 mm.

The detection of ventral neurons in the Gli2/3 dorsal telencephalon could

be due to defective migratory pattern of the post-mitotic neurons or expansion of

ventral progenitors to the dorsal domain. We therefore examined the expression

of Dlx2, which labels all ventral telencephalic progenitors [151]. We found a

normal expression of Dlx2 in Gli1, Gli2 and Gli1/2 double mutants (Figure 4-7F–

H). In Gli3 mutants, Dlx2 expression was restricted in the tissues of telencephalic

origin (Figure 4-7I). Interestingly, in Gli2/3 double mutants, the expression of Dlx2

was detected in both ventral and dorsal regions of the telencephalon, suggesting

that ventral progenitors expanded into the dorsal telencephalon (Figure 4-7J).

To further determine whether different dorsal and ventral progenitors

intermingle as a result of expansion of ventral progenitors, we performed double immunoﬂuorescence staining of Pax6 and Gsh1/2. Pax6 and Gsh2 are normally expressed in a complementary pattern from E10.5 to E12.5, with Pax6 expressed in the dorsal telencephalic progenitors and Gsh2 expressed in the ventral telencephalic progenitors [152]. In WT embryos at E12.5, Pax6 and Gsh2 are co- expressed only in a few rows of cells in the corticostriatal region (Figure 4-7K).

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The mutually exclusive expression pattern was maintained in Gli1, Gli2 and

Gli1/2 double mutants (Figure 4-7K, L, M). In Gli3 mutants, Gsh2 was detected in the dorsal region, likely because of the diencephalic transformation (Figure 4-7N).

Interestingly, in Gli2/3 double mutant embryos, the expression pattern of Pax6 was largely unperturbed (Figure 4-7O, green channel). However, Gsh2 expression was expanded into the dorsal telencephalon and thus overlapped broadly with Pax6 expression (Figure 4-7O, red channel).

To conﬁrm that the expression domains of different markers intermingle as a result of loss of Gli function, we examined the expression of two additional makers, sFRP2 and Isl1. sFRP2 encodes an inhibitor of Wnt signaling and is expressed in the ventral-most region of the dorsal (pallial) telencephalon (Figure

4-7P) [153] and Isl1 is expressed in neurons ventral to the sFRP2 domain in WT embryos (Figure 4-7Q). In Gli2/3 double mutants, the expression domain of sFRP2 was greatly expanded (Figure 4-7R, indicated by a bracket). Moreover,

Isl1+ neurons clearly expanded into the dorsal region, resulting in a partial overlap with sFRP2 in Gli2/3 double mutants (Figure 4-7S). Together, these results suggest that both neural progenitors and neurons have lost their distinct spatial organization in the absence of Gli function.

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4.5 Discussion

Previous studies have shown that Gli1 and Gli2 are expressed in the

interganglionic sulcus, suggesting that this region receives a high level of Hh

signaling [102, 139]. We analyzed telencephalic patterning in embryos lacking

major Gli activators in Gli2 and Gli1/2 double mutants, and found that the

expression of three critical transcription factors, Nkx2.1, Nkx6.2 and Gli1, was

signiﬁcantly reduced in progenitors receiving high levels of Shh signaling.

Furthermore, we found that from E9.5 to E12.5, Shh-responding progenitors give

rise to neurons occupying successively more superﬁcial layers in the cortex.

Finally, when all Gli transcription factors are removed, speci ﬁcation and

production of ventral interneurons are severely perturbed. Together, our results

uncover multiple roles for Gli proteins in speci ﬁcation, differentiation and survival

of telencephalic progenitors.

Transcriptional activation mediated by Gli proteins is required to induce

distinct progenitor populations in the ventral telencephalon

Shh signaling has been shown to specify distinct cell fates in a

concentration-dependent manner in the developing spinal cord, with a higher

concentration of Shh inducing ﬂoor plate cells and a progressively lower

concentration of Shh inducing V3 interneurons, motor neurons and V2–V0

interneurons (reviewed in [89]). The ability of Shh signaling to induce different

cell types is dependent on the Gli family of transcription factors. In particular, it

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has been shown that a gradient of Gli activator is able to mimic Shh signaling to induce different cell types in the developing chick spinal cord [154].

It is known that different subtypes of interneuron are generated from different regions of the ventral telencephalon at different developmental stages in a predictable spatial–temporal sequence [83, 90, 143]. The LGE contains at least two progenitor populations, with one population expressing Dlx2 and Er81 that generates olfactory bulb interneurons, and a second population expressing Dlx2 and Isl1 that contributes to striatal neurons [144]. Furthermore, the dorsal-most

LGE cells, located next to the Pax6 expressing pallium, have the distinct ability to contribute to the lateral cortical stream and migrate to the developing ventral telencephalon [155]. A recent study has proposed to further divide the LGE into four progenitor groups (pLGE1–4) and the MGE into ﬁve progenitor groups

(pMGE1–5), based on a combinatorial expression pattern of transcription factors in the ventral telencephalon [156].

Shh signaling has been postulated to inhibit the repressor function of Gli3 in the telencephalon [101]. Our analysis further suggests that Gli activator mediate Shh signaling to establish distinct progenitor populations in the ventral telencephalon. Based on the unique responses to Shh signaling, progenitors in the interganglionic sulcus can be divided into two subpopulations (Figure 4-8), one located on the MGE side and express Gli1high, Nkx6.2 and Nkx2.1 (group 1

in Figure 4-8), and the second group located on the LGE side and express Gli1low and Nkx6.2, but not Nkx2.1 (group 2). The two Shh responsive progenitor groups identiﬁed in our study appear to correspond to pLGE4 and pMGE1. In Gli2

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mutants, speciﬁcation of both progenitor groups is disrupted. More dorsally, speciﬁcation of two other progenitors (groups 3 and 4) does not appear to be

affected in the absence of Gli2 or Gli1/2. It is likely that a low level of Shh

signaling, which is sufﬁcient to attenuate the level of Gli3 repressor, is adequate

for the speciﬁcation of these two progenitor groups. Therefore, Shh signaling in

the ventral telencephalon may have two different functions: to repress the

function of Gli3 and at the same time, activate downstream transcriptional targets

through Gli activators (Figure 4-8), by a mechanism similar to that in the spinal

cord [51-53, 154].

Consistent with the previous observation that dorsal MGE progenitors

(pMGE1) have a bias for SST+ neurons [139, 156, 157], we found that ~20% of

progeny generated from E8.5 Gli1-expressing progenitors were SST+ neurons,

and the number increased to ~25% at E11.5, suggesting that Hh signaling may

have a role in the speciﬁcation of this group of neurons. In addition, examination

of β-galactosidase+ cells 36 h following TM injection revealed that cells labeled

at E9.5 and E10.5 spread over a broader region in the MGE, compared with cells

labeled at E11.5 that are limited to the interganglionic sulcus. Together, these

results raise the possibility that Hh signaling may involve in speci ﬁcation and

consolidation of SST+ progenitors in the dorsal MGE.

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Figure 4-8 Schematic of Shh/Gli signaling in the ventral telencephalon. Based on the expression of different transcription factors, ventral telencephalic progenitors responding to Shh signaling can be divided into four groups (1–4). The ventral-most progenitors (groups 1) express Gli1, Nkx6.2 and Nkx2.1, and are located in the MGE side of the interganglionic sulcus. Group 2 progenitors express Gli1, Nkx6.2 but not Nkx2.1, and are located on the LGE side of the interganglionic sulcus. These two groups of progenitors are not speciﬁed in the absence of Gli activator. More dorsally, another two groups of progenitors have been de ﬁned previously. One group produces Er81+ neurons and the other group generates isl1+ neurons. These four groups of progenitors are absent when Hh signaling is conditionally disrupted at E8.5, but are not affected in the absence of Gli activator. It is likely that groups 3 and 4 of progenitors require only a low level of Shh signaling to inhibit the formation of excess Gli3 repressor. Shh is also likely required to attenuate Gli3 repressor in most of the MGE cells, since conditional loss of Hh responsiveness eliminates all ventral telencephalic cells. See text for details. LGE, lateral ganglionic eminence. MGE, medial ganglionic eminence.

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Proliferation and organization of ventral telencephalon require Gli- mediated Hh signaling

Several defects in ventral telencephalon development were observed in

Gli2/3 double mutant embryos, including dorsal expansion of ventral telencephalic markers, the intermingling of different progenitor domains and the reduced differentiation of ventral neurons. These phenotypes have never been observed in Gli2 mutants (data not shown) and are distinct from the phenotypes observed in Gli3 mutants.

In Gli3 mutant, Gli1, Ptc1 and Shh were detected in the mutant ventral telencephalon, suggesting a normal ventral patterning [102, 158]. On the other hand, in the dorsal forebrain, there is a diencephalic displacement of telencephalic tissues, resulting in a joining of diencephalic and telencephalic tissues and the expression of several markers normally expressed in both ventral telencephalon and diencephalon [99-102]. As the loss of Gli3 repressor has been shown to increase the range of Shh signaling in the limb and to cause expansion of intermediate cell types in the ventral spinal cord [159], possibly through derepression of Gli activator, an increase in Shh signaling from the zona limitans intrathalamica (Zli) is a likely reason for the diencephalic displacement of dorsal telencephalic tissues in Gli3 mutants. Consistent with this, there appears to be an enlargement of the diencephalon based on a caudal shift of Wnt1 expression in

Gli3 mutants [158].

The phenotypes of pan-Gli mutants are different from the phenotypes of

Gli3 in at least three aspects. First, FoxG1 is expressed throughout pan-Gli

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mutant telencephalon, suggesting that the telencephalic characteristic is

maintained in these embryos. Second, the expression of Gli1 and Nkx6.2 was

not detected in the ventral telencephalon of pan-Gli mutants ([51] and data not

shown). The loss of these two markers thus suggests an additional loss of ventral

telencephalic progenitors in pan-Gli mutant embryos. Third, different ventral

telencephalic cell types intermingle in Gli2/3 mutants, suggesting that positional information is lost. A similar phenotype has also been observed in the pan-Gli

mutant spinal cord [51, 52] or in the Smo;Gli3 double mutant embryos [160]. Our

analysis therefore provides further support that Gli genes are required for the

establishment of separate neuronal domains throughout the ventral neural tube.

Finally we found that the production of post-mitotic neurons in the ventral telencephalon is severely reduced when all Gli function is removed. This phenotype is more severe than the phenotype of any single Gli mutant or Gli1/2

double mutants. Most likely this phenotype is caused by an overlapping as well

as unique functions between different Gli proteins. Indeed, it appears that

different tissues have different requirements for the activator or repressor

function. For example, only Gli repressor is needed for digit pattering in the limb

[107, 161, 162] while Gli activator is required for cell fate speci ﬁcation in the

spinal cord [51, 154, 159]. Based on the analysis of the telencephalic phenotypes

of different Gli mutants, we propose that only Gli3 repressor is required for the

suppression of ventral telencephalic and/or diencephalic markers in the dorsal

telencephalon, while the three Gli proteins are involved in controlling the

speciﬁcation of distinct progenitors, the separation of different progenitor

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domains and the differentiation of telencephalic neurons.

The fact that some aspects of dorsoventral patterning remain intact in embryos devoid of all Gli function also raises the question of what other signaling pathways, in addition to Shh, specify the remaining cell types. Our analysis of spinal cord development suggests that both Wnt and Shh signaling pathways are involved in controlling cell type generation [163], although one recent report excludes Wnt signaling in cell type speci ﬁcation in the ventral telencephalon

[164]. Another likely signal is FGF signaling. It is known that several of the FGF ligands and FGF receptors are expressed in the developing telencephalon

(reviewed in [165]). In particular, in Fgfr1;Fgfr2 double mutant embryos, many ventral progenitor cells are not speci ﬁed [166]. The loss of ventral progenitor phenotype is similar to that of conditional loss of Smo in the telencephalon [103], raising the possibility that FGF signaling functions in a linear pathway with Shh signaling [166]. However, the phenotype of Fgfr1;Fgfr2 double mutants is clearly more severe than that of the pan-Gli mutant embryos, because dorsal markers are not expanded into the ventral domains in pan-Gli mutants. The differences in phenotypes would therefore suggest that FGF signaling functions parallels that of

Shh signaling, for example in lineage commitment, as has been recently shown in the ES cells [167], although it is also possible that the Gli genes may have function independent of Shh signaling. Future challenges will be to delineate the relationship between Shh and FGF signaling in patterning and generation of telencephalic neurons.

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4.6 Acknowledgements

We thank Dr. Alexandra Joyner for her support and encouragement during

the initial phase of the work, and for stimulating discussions on the project. We

thank Drs. Sohyun Ahn, Sandra Blaess and Mark Zervas for their effort in the initial collaborative Gli1-CreER fate-mapping project, and for comments on the

manuscript. We thank Drs. Gord Fishell, Joshua Corbin and Kenneth Campbell

for providing probes, antibodies and suggestions, and Man-Sun Sy and Ron

Conlon for critical reading of the manuscript. Pax6 monoclonal antibody from the

Developmental Studies Hybridoma Bank was developed under the auspices of

the NICHD and maintained by the Department of Biological Sciences at The

University of Iowa. This work was supported by a startup fund from CWRU and in

part by a New Scholar Award from the Ellison Medical Foundation.

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Chapter 5 Conclusions and Future Directions

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5.1 Summary

Studies of signaling molecules and transcription factors during embryonic

development of the pituitary gland have demonstrated that multiple signals are

required in pituitary organogenesis, proliferation and differentiation [4, 6, 19, 23-

30, 32-34, 75, 77, 78, 80, 81, 123, 168, 169]. It was already known that Shh signaling is required for embryonic development of the pituitary [31]. My thesis

work aimed at identifying how Shh/Gli signaling regulates pituitary development

in the mouse embryo.

My main discovery is that Gli2 is required to control cell proliferation within

Rathke’s pouch. First, loss of Gli2 had no impact on pituitary patterning. Gli2-/-

mutant pituitaries and pituitary-specific Gli knockouts (FoxG1-Gli2 and Pitx2-Gli2)

expressed normal patterning markers in Rathke’s pouch. Second, Gli2-/- mutant

pituitaries have all five ventral cell types of the anterior pituitary although the

pituitary size is much smaller. Third, pituitary-specific Gli2 knockouts (FoxG1-Gli2)

have proliferation defects in Rathke’s pouch and reduced numbers of

corticotropes and lactotropes. These results indicate the major role of Gli2 in

pituitary is regulating proliferation rather than determination of pituitary cell

lineages or cell types.

Previous research has shown variable defects within the pituitary of Gli

mutants [42]. In my study, I examined Gli1lacZ/lacZ, Gli2zfd/zfd (same as Gli2-/- in this

thesis), Gli3xt/xt and Gli2zfd/zfd;Gli3xt/xt mutants. The Gli1 and Gli3 homozygous

mutant embryos all have pituitaries expressing normal cell type markers. Gli2-/-

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mutant embryos have a high ratio of anterior pituitary loss (around 75%) both at

E12.5 (24/32) and E17.5 (15/21). None of the Gli2 mutants had infundibulum or the posterior pituitary. As the infundibulum and the posterior pituitary come from the ventral diencephalic tissues, my result is consistent with a previous paper suggesting Gli2 mutants have tissue loss within the diencephalon [42].

5.2 Early Shh signaling in the ANR may specify the pituitary primordium

Active Hh signaling was detected in the anterior neural ridge (ANR), which is the precursor of both anterior and posterior pituitary. In zebrafish, Hh signaling has an early direct role in pituitary induction [70]. This early Hh activity in the anterior neural ridge may also have a role in specifying the mouse pituitary primordium. Further fate mapping analyses suggest Hh responsive cells labeled at E8 contribute to the Rathke’s pouch at E12.5. At E6.5~E8 in the mouse embryo, pituitary origins reside in the midline structure of ANR [9]. These results indicate that Shh/Gli signaling may play a role in specifying ANR cells during early pituitary development in mice.

To further prove this hypothesis, conditional knockouts studies were performed. However, FoxG1-Gli2 mutants still had proper developed Rathke’s pouch. The most obvious defect of these FoxG1-Gli2 mutants was reduced proliferating rates in the anterior RP. FoxG1-Cre mediated recombination is present in the rostral part of the neural plate at E7.5, but recombination at this stage appeared mosaic throughout the anterior neural plate [103]. At this stage,

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Gli2 activity is already in the neural plate [110]. Thus FoxG1-Gli2 mutants did not knockout Gli2 activity thoroughly in the anterior neural ridge. Thus, the early role of Shh signaling remains to be tested.

5.3 Two different phases of Shh/Gli signaling in pituitary proliferation

Shh is expressed in the ventral diencephalon (E9-E14) and the adjacent oral ectoderm (E10-E12) of Rathke’s pouch while Ptc1 is highly expressed inside

the developing pouch [31, 33]. Gli1, Gli2, and Gli3 are also expressed in the

ventral diencephalon and within Rathke’s pouch [69]. Therefore, the developing

pituitary gland is competent to receive and respond to Shh signals. However,

which source of Shh is required for pituitary patterning and proliferation remains

unclear.

As proposed in a published paper, Rathke’s pouch responds to Hh

signaling from the oral ectoderm underneath [31]. Shh over-expression in the

developing Rathke’s pouch with αGSU promoter resulted in expansion of pituitary

ventral cell types. In contrast, pituitary specific blockage of Hh signaling by over-

expressing the Shh antagonist Hip gave rise to pituitary hypoplasia. However,

Pitx-Hip transgenic would block all Shh signals in Rathke’s pouch, no matter

where Shh emaninates. Shh can be detected in the ventral diencephalon besides

the oral ectoderm [33]. Rathke’s pouch may receive Shh signals from the

adjacent ventral diencephalon as well.

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During pituitary patterning and proliferation (E10-E12), Shh is expressed

in the ventral diencephalon and oral ectoderm (Figure 5-1). At E10, Rathke’s

pouch, which is not yet separated from the adjacent oral ectoderm, receives Shh

signals both from the ventral diencephalon and the oral ectoderm. As Rathke’s

pouch grows and expands, it is detached from the oral ectoderm at E11. Shh/Gli

signaling, from the ventral diencephalon and the oral ectoderm, promotes

pituitary cell proliferation in anterior wall and tip region, respectively. At E12.5,

the infundibulum is about to be completely detached from the ventral

diencephalon. Rathke’s pouch receives Shh signals from the underlying oral

ectoderm. Shh/Gli signaling from the oral ectoderm promotes pituitary cell

proliferation in the ventral anterior region of Rathke’s pouch.

At E10, Gli expression was found in the whole Rathke’s pouch. However, I

observed an anterior to posterior Gli gradient in Rathke’s pouch, reflecting higher

Hh response in the anterior wall of the pouch. Meanwhile, Shh is expressed in

the anterior ventral diencephalon, which is adjacent to the anterior RP (Figure 5-

1). This indicates Rathke’s pouch may receive Shh signals from the ventral

diencephalon besides the oral ectoderm. Later at E12, Gli expression gradient

shifts to ventralhigh to dorsallow, when Rathke’s pouch is completely separated from the underlying oral ectoderm and is distant from the diencephalon. Rathke’s pouch may only receive Shh signals from the oral ectoderm at this time.

In the present thesis, I discovered a critical role of Shh/Gli signaling in

pituitary cell proliferation, especially in the anterior side of Rathke’s pouch. This

part of RP later becomes the anterior lobe of the mature pituitary gland. The

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Figure 5-1 Shh/Gli signaling is required for pituitary cell proliferation.

At E10, Shh is expressed in the anterior ventral diencephalon and the oral ectoderm adjacent to Rathke’s pouch (green). Gli expression gradient is from anterior to posterior (left to right in the diagram). At E11, Shh is expressed in the anterior ventral diencephalon and the underlying oral ectoderm (green). Gli expression gradient is not obvious at this time. Rathke’s pouch is separated into three regions by red dashed lines: the anterior wall (A), the posterior wall (P) and tip region (Tip). Cell proliferations in the anterior wall and the tip region are promoted by Shh/Gli signaling. At E12, Shh is expressed in the ventral diencephalon and the underlying oral ectoderm (green). Gli expression gradient is from ventral to dorsal (bottom to top in the diagram). Rathke’s pouch is separated into four regions by red dashed lines: the dorsal anterior region (Ad), the ventral anterior region (Av), the posterior region (P) and the rostral tip region (rT). Cell proliferations in the ventral anterior region are promoted by Shh/Gli signaling. Precursors of the five ventral cell types, corticotropes (C), somatotropes (S), lactotropes (L), thyrotropes (T) and gonadotropes (G) are located in the anterior region of Rathke’s pouch. Precursors of dorsal cell type melanotropes (M) are located in the posterior region. (RP-Rathke’s pouch; OE-oral ectoderm; VD-ventral diencephalon; INF- infundibulum; ↑P-promote proliferation; Adapted and modified from Scully and Rosenfeld, 2002 [16])

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posterior side behind the lumen (pituitary cleft), which develops into the intermediate lobe of the pituitary gland that contains melanotropes (M), was less affected.

Mosaic analysis of Gli2 showed that Gli2 mutant cells did not proliferate well in E11 Gli2 mosaic mutants. In the anterior wall and tip region of Rathke’s pouch, proliferating cells contributed by the Gli2 mutant lineage were significantly reduced. This suggests Shh/Gli signaling is required for cell proliferation in the anterior wall and tip region of Rathke’s pouch. At E11, Shh expression is in the anterior ventral diencephalon, which is adjacent to the anterior wall of Rathke’s pouch. Rathke’s pouch may receive Shh signals from the ventral diencephalon to promote cell proliferation in the anterior RP. In the ventral side, Shh expression in the oral ectoderm may promote cell proliferation in the tip region.

In mouse embryos, precursors of different pituitary cell types are assumed to reside in different parts of Rathke’s pouch (Figure 5-1). The GATA2+ cell lineage including gonadotropes (G) and thyrotropes (T) is located at the most ventral part of RP [32, 33]. The Pit1+ cell lineage including somatotropes (S), lactotropes (L) and thyrotropes is located at the anterior side of the pouch lumen and dorsal to GATA2+ lineage [23, 169]. The Tbx19 lineage including corticotropes (C) is located at the anterior side of the pouch lumen as well [35].

The posterior side of the pouch lumen develops into the intermediate lobe of the pituitary which contains dorsal cell type melanotropes (M) [170].

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I used FoxG1-Gli2 mutants to remove Gli2 function specifically in the

Rathke’s pouch before RP detachment (before E10). In my proliferation analyses, the Rathke’s pouch at E12.5 was divided into rostral tip (rT), posterior (P), ventral anterior (Av) and dorsal anterior (Ad) regions (Figure 5-1). Rostral tip region contains matured pituitary cells while the other parts are still under proliferation and differentiation [19]. In FoxG1-Gli2 mutant pituitaries, we found fewer proliferating cells in the anterior RP and significantly reduced proliferation rate in the Av region. As FoxG1-Gli2 mutants had fewer proliferating cells in the anterior

RP at E12, fewer progenitors of pituitary cells were expected. As a result, fewer cells of certain pituitary cell types (corticotropes and lactotropes) were found in the anterior pituitary at E17.5. The precursors of these cell types once resided in the anterior wall of Rathke’s pouch (Figure 5-1). These results indicate Shh/Gli function is important for proliferation in anterior Rathke’s pouch.

I then used Pitx2-SmoM2 mutants to over-express active form of Smo in the Rathke’s pouch. At E12, the number of proliferating cells was significantly increased in the anterior RP of Pitx2-SmoM2 mutants, especially in the Ad region.

At E14.5, Pitx2-SmoM2 mutant pituitaries were much bigger in size than WT controls, with the dorsal anterior region of RP greatly expanded. Direct activation of Hh signaling in RP enhanced the overall Hh response in Rathke’s pouch.

Assuming Rathke’s pouch only receives Shh signals from the oral ectoderm at

E12, ventral part of the pouch should be highly responsive to Hh signaling. Over- expression of Smo by Pitx2 would greatly increase the Hh signaling level in the dorsal RP, where Hh response (Gli1 activity) is not as strong as in the ventral

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part. To prove my hypothesis, future work is required to determine the temporal and spatial requirement of Shh signals.

5.4 Gli2 may be required for diencephalon-derived signals Bmp4 and Fgf8

The Bmp, Shh, Wnt and Fgf signaling pathways are involved in the

development of the pituitary gland. The expression pattern of Bmp4, Fgf8 and

Shh in the ventral diencephalon indicates possible roles of signaling interactions

during pituitary development. Previous studies have shown that Bmp4 and Fgf8

activity from the ventral diencephalon are required for the development of a

definitive pouch [17, 33]. Thus, Bmp4 and Fgf8 are required for both the anterior

pituitary (coming from oral ectoderm) and the posterior pituitary (coming from

ventral diencephalon).

Shh is known to be required at the early stage of diencephalon

development and Gli2 expression is also found in the ventral diencephalon

besides within the Rathke’s pouch [66, 69]. A previous paper reported loss of

diencephalic tissues in the Gli2 mutants [42]. In our study, Gli2 mutants had a

high ratio of anterior pituitary loss while no posterior pituitary was detected in

these mutants. Further examination showed that Bmp4 and Fgf8 expression was

greatly reduced in the ventral diencephalon of Gli2 mutants. Reduction of

diencephalon-derived signals in Gli2 mutants indicates Gli2 may mediate Bmp4

and/or Fgf8 signaling in the ventral diencephalon to regulate pituitary induction.

Yet, it remains to be determined whether Bmp4 and/or Fgf8 expression is up-

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regulated by activated Hh signaling (Pitx2-SmoM2). Future work will be needed to determine whether Gli2 protein directly bind to Bmp4 and/or Fgf8 gene regulatory elements.

5.5 Mouse Gli2 mutant phenotypes correlate with human HPE

Genes discovered in the mouse often lead to the discovery of human genetic diseases and thus reveal the mechanism at a genetic level. In the field of pituitary studies, this is also the case. For instance, the discovery of the mouse gene Pou1f1 and Prop1 facilitate the identification of the mutated human genes in hypopituitarism and pituitary related dwarfism [4, 169]. However, some genes which are reported necessary for normal pituitary growth in mice have not shown their significance in human patients. The ability of mouse mutants to predict human genetic deficiency in pituitary diseases is remarkable, although the correspondence is imperfect as genome variations always exist between the two species [171].

In humans, alterations in SHH signaling are associated with a number of pathological states, such as basal cell carcinoma (BCC) and holoprosencephaly

(HPE) [172]. Gain-of-function of Shh in the epidermis induces basal cell carcinomas of the skin [173, 174]. Patients carrying heterozygous mutations in

SHH present holoprosencephaly phenotypes ranging from severe to minor [55,

66]. Several other genes with direct or indirect links to hedgehog have been implicated genetically in human HPE, including ZIC2, SIX3, PTCH1, GLI2, DISP1,

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SMOH and HHIP [54, 62-65]. Thus SHH signaling plays a significant role in HPE

etiology.

Human GLI2 gene loss-of-function mutations are associated with pituitary

anomalies and HPE-like features [62]. However, only seven out of total 390

patients with HPE were identified with GLI2 mutations. Clinical studies suggested

the pituitary and facial structures were the most sensitive to a reduction in GLI2 activity among all HPE phenotypes [62]. Novel nonsense GLI2 mutations were

also identified in patients without HPE but with hypopituitarism and ectopic

posterior pituitary lobe [175]. Gli2 deficient mice either lost their anterior pituitary

or had pituitary hypoplasia (Chapter 3). Gli2-/- mice also had severe defects in the

ventral diencephalon and lost their posterior pituitary (Chapter 3). All these

discoveries suggest an important role for the GLI2 gene in pituitary development.

However, genome variations and brain structures difference between mice

and human beings may affect conserved GLI2 function in pituitary development.

SHH and GLI2 mutations found in patients with pituitary defects are all

halpoinsufficient. In the mouse embryos, heterozygous Shh or Gli2 loss-of-

function mutations caused no pituitary phenotype. Homozygous Gli2 loss-of-

function mutations gave rise to pituitary defects and could not survive afterbirth.

This discrepancy could be caused by the functional divergence of Gli2 proteins in

various species. Human beings may be more sensitive to GLI2 level or one copy

of Gli2 might be sufficient to maintain homozygous Gli2 level in mice. Difference

in signaling molecules of Shh pathway should be considered as well.

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5.6 Future directions

As my thesis work shows the importance of Shh/Gli signaling in cell proliferation of the embryonic pituitary, much remains unknown in this field.

Future studies will need to determine whether and how Shh/Gli is required in early specification of the pituitary primodium and cell type specification of the anterior pituitary. Search for interactions between Shh and other signaling pathways during pituitary development is also a challenging task. As pituitary continues to grow after birth, Shh defects may affect adult pituitary secretion as well.

5.6.1 Shh/Gli signaling is required for early specification of the pituitary primordium

In mouse embryos, Shh/Gli signaling is active in the pituitary primordium and pituitary cell progenitors (Chapter 2). Mosaic analysis showed Gli2 is cell autonomously required in Rathke’s pouch (Chapter 3). In zebrafish, Hh singaling plays an early direct role in pituitary induction [70]. However, morphogenetic process of pituitary formation in zebrafish is different from that of the mouse. To determine whether Shh/Gli signaling plays a role in specifying mouse pituitary primordium, I propose the following experiments.

In Chapter 3, I used FoxG1-Cre mouse line to remove Gli activity in the anterior neural ridge and found defects in pituitary cell proliferation but not in the formation of pituitary primordium. At E7~E8, FoxG1 regulated Cre activity was

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mosaic in the anterior neural ridge, thus FoxG1-Gli2 mutants might still preserve

partial Gli functions. Here I propose to use another Cre line to fulfill the ANR

removal of Gli functions. Hesx1Cre mouse line has Hesx1 regulated Cre activity

detected in the anterior neural plate (E7.5), anterior forebrain region (E9) and

Rathke’s pouch/ventral diencephalon (E10.5) [176]. Hesx1Cre mouse line will be

crossed with Gli2flox/zfd to remove Gli2 activity in the anterior neural plate. If

Shh/Gli signaling is required for early specification of the pituitary primordium,

Hesx1-Gli2 mutants will have defects in the formation of initial Rathke’s pouch.

Considering the redundant function of Gli2 and Gli3, Hesx1-pan-Gli mutants will

be examined as well.

Another experiment to determine the role of Shh/Gli signaling will be in

vitro culture of the anterior neural ridge. Micro-dissection will be performed on mouse embryos to collect the ANR tissue. At E7~E8, the anterior neural ridge, which is the origin of the pituitary, will be isolated from the neural plate and cultured. Hesx1Cre;Rosa26-EGFP mouse embryos will be used to visualize GFP-

labeled anterior neural plate [176, 177]. The anterior neural plate explants will be

dissected from 6-somite (late E7 and early E8) mouse embryos with glass

needles in PBS, then transferred to 500 ml of 2.5% pancreatin (Sigma), 0.5%

trypsin (Sigma) and incubated 15 min at 4°C [178]. After the above treatment

with enzymes to separate tissue layers, ANR explants will then be transferred to

culture medium (Dulbecco’s modified Eagle medium with streptomycin and

penicillin, plus 15% fetal calf serum). All explants were cultured in individual

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drops of medium under mineral oil on plastic petri plates (Fisher) in a 37°C humidified incubator with 5% CO2 [178].

In the control group, explants will be exposed to Shh-N (biologically active amino-terminal fragment of mouse Shh protein) for 48 hr. After 24~48 hrs culture, multiple markers including Lhx3 and Pitx1/2 will be used to determine the early specification of the pituitary primordium. In the control group, the cultured ANR explants are expected to express early pituitary markers. The concentration of

Shh-N treatment will be adjusted to obtain the optimum result (0.4nM~4nM). In the second group, ANR explants will be cultured without addition of Shh-N treatment. If Shh is required for induction of pituitary progenitor cells, this group of ANR explants should be unable to express Lhx3 and Pitx1/2. The third group of the ANR explants will be exposed to Shh-N (optimum concentration) while treated with Shh blocker. Anti-Shh treatment will be added in the explants culture medium, including antibodies (anti-Shh-N) or cyclopamine [179]. After 48 hr culture, the third group of explants should fail to express pituitary markers as well as the second group. However, increasing the concentration of Shh-N in the presence of anti-Shh-N antibody should restore pituitary induction. In the fourth group, the ANR explants will be cultured with high concentration of Shh-N

(~13nM) in the presence of anti-Shh-N treatment. This group of explants is expected to express Lhx3 and Pitx1/2.

To further examine the role of Shh/Gli signaling in the specification of pituitary progenitors, the ANR explants will be collected at 12-somite (late E8 and early E9) stage and cultured for 72~96 hr with or without the Shh-N/anti-Shh-N

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treatment (Figure 5-2). Pituitary-specific markers including ACTH, TSH, isl1 and

αGSU will be examined. Whether Shh signaling is required for specification of the pituitary primordium and pituitary progenitor cells will be revealed.

Figure 5-2 In vitro culture of the ANR explants

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5.6.2 Shh/Gli singaling is required in the ventral diencephalon for pituitary

cell proliferation

As suggested in my thesis, Shh signals may be required both from the

ventral diencephalon and oral ectoderm. To determine whether Shh signals are

required in ventral diencephalon for pituitary proliferation, I will perform tissue specific loss-of-function studies of Shh. Ventral diencephalon specific Shh

knockout mutants will be designed to investigate the requirement of Shh source.

Nkx2.1Cre and Shhflox mouse lines will be use to remove Shh expression in the

anterior diencephalon as early as E10 [180-182]. If the ventral diencephalon

derived Shh signal is required for pituitary cell proliferation, these conditional

mutants will display pituitary hypoplasia.

5.6.3 Shh/Gli singaling is required for cell type specification of the anterior

pituitary

In the current thesis, Shh/Gli signaling is required to regulate pituitary cell

proliferation. However, Gli2 mutants and RP-specific Gli2 mutants displayed a

preference of pituitary cell type loss. Though FoxG1-Gli2 mutants had

proliferation defects only in the ventral anterior region of Rathke’s pouch at E12,

it caused dramatic loss of pituitary cell types (corticotropes and lactotropes) at

E17. To investigate the requirement of Shh signals in cell type specification of the

anterior pituitary, I will use a series of fate mapping to trace Hh-responding cells.

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How these Hh responsive cells labeled at different time periods contribute to different pituitary cell types will be analyzed and compared.

In my preliminary data, Hh-responding cells at E10 in Rathke’s pouch were found to contribute to all five ventral cell types in the anterior pituitary. Gli1 expression gradient in RP shifted during E10 to E12, indicate different Hh response in Rathke’s pouch at different time points. Moreover, each pituitary cell type differentiates and proliferates at different time period under the regulation of transcription factors (Figure 1-3).

To determine whether Shh signals at different stages contribute to the cell proliferation of different pituitary cell types, I will label Hh responsive cells at E10,

E11 and E12. Embryos will be collected at E17.5 and double immunostaining will be applied. β-gal staining reveals labeling of the Hh-responding cells while cell type specific hormonal antibodies will be used to mark different cell types in the

anterior pituitary. The double labeling cells of β-gal and cell type specific

hormone in each section are Hh-responding cells which contribute to certain cell

type. Analysis of multiple embryos for different labeling time points will show us

how Shh/Gli signaling specifies different pituitary cell types.

5.6.4 Interactions between Shh and other signaling pathways

The Wnt, Bmp, Fgf, and Shh pathways were found to have profound

effects on pituitary development. This puts signaling molecules regulating

pituitary development in a complicated model rather than a simple one. Signaling

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interactions may exist during pituitary development and it would be worthwhile to identify precisely how these pathways interact.

5.6.4 a Shh interacts with Bmp2 to regulate pituitary differentiation

It has been shown that Gli2 mediates Bmp2 expression in osteoblasts in response to Hedgehog signaling [183]. Bmp2 has been shown to play a major role in osteogenesis, including chondrogenic and osteogenic differentiation [184].

Shh signaling has been shown to be necessary for skeletogenesis, as Shh deficient mutants had severe skeletal abnormalities [66]. Shh stimulates Bmp2 promoter activity and osteoblast differentiation. Over-expression of Gli2 enhanced Bmp2 expression in osteoblast precursor cells. Genetic deletion of the

Gli2 gene inhibited Bmp2 expression in osteoblasts. Moreover, Gli2 binding sites have been identified in the Bmp2 promoter by promoter deletion analysis, ChIP assays and EMSAs [183].

Several lines of evidence suggest Shh might regulate Bmp2 to control cell proliferation of anterior pituitary. First of all, Bmp2 expression was detected in the pituitary, at the boundary of the oral ectoderm and Rathke’s pouch during E10.5 to E14.5. Shh is expressed in the adjacent oral ectoderm during E9.5 to E12 [33].

Secondly, loss-of-function studies of both Shh and Bmp2 showed similar pituitary phenotypes. Blocking Shh signals by Pitx1-Hip reduces the size of Rathke’s pouch and results in a smaller pituitary. The same phenotype appears when

Bmp2 is ablated specifically in Rathke’s pouch [31, 33, 75]. Moreover, Pitx1-Hip

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transgenic embryos have no Bmp2 or Gata2 (Bmp2 target) expression in the

Rathke’s pouch. This suggests blocking Shh in the Rathke’s pouch may disrupt

Bmp2 signaling in pituitary. In contrast, over-expression of Shh using aGSU-Shh results in an induction of Bmp2 [31]. Together, these results suggest that Shh signaling may regulate Bmp2 expression during pituitary proliferation. As Gli2 function is required in pituitary cell proliferation, I hypothesize that Gli2 activates

BMP2 in response to Shh to regulate cell proliferation in the anterior pituitary.

To determine whether Gli2 mediates Bmp2 signaling during pituitary proliferation, the expression of Bmp2 and a possible readout pSmad1 will be examined in Rathke’s pouch of Gli2-/- or pituitary-specific Gli2 mutant embryos. pSmad1 is a phosphorylated form of Smad1, an intracellular signaling component of the Bmp pathway [185]. Smad1 is phosphorylated in the forming infundibulum and ventral Rathke’s pouch of wild type embryos where Bmps

(Bmp2 or Bmp4) are expressed [75]. pSmad1 can serve as an effective readout of Bmp signaling during pituitary proliferation. Down-regulation of Bmp signaling is expected in the Gli2-/- or conditional Gli2 mutants. However, it is possible that pSmad1 activity can still be detected in the anterior pituitary as Bmp4 signaling also exerts roles in development of anterior pituitary [17, 33, 97].

To test whether activation of Shh signaling upregulates Bmp2 signaling, I will activate Hh signaling in Rathke’s pouch and examine Bmp2 expression levels

(Pitx2-SmoM2, as described in Chapter 3). High levels of Bmp2, Bmpr1a and pSmad1 are expected in the Rathke’s pouch due to upregulated Bmp signaling in

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response to Hh. However, Bmpr1a and pSmad1 activity alone cannot indicate

the status of Bmp2, since these factors respond to all Bmps.

To ascertain that loss of Bmp2 signaling is responsible for pituitary

proliferation defects in Gli2 mutants, gain-of-function analysis of Bmp2 will be

performed using an αGSU-Bmp2 transgene [32]. In the Gli2 mutants I examined,

αGSU expression was detected in the Rathke’s pouch. This makes it realistic to

use αGSU-Bmp2 transgenic mouse to express ectopic Bmp2 in the Gli2-/- mutants. If Gli2 mediates Bmp2 signaling in response to Shh to regulate pituitary proliferation, over-expression of Bmp2 in the Gli2 pituitary should rescue the

pituitary proliferation defects. Since Gli2 could be required in both pituitary and

ventral diencephalon, it is possible that αGSU-Bmp2 cannot rescue the pituitary

phenotype of Gli2-/- mutants. If this is the case, then I will attempt to use αGSU-

Bmp2 to rescue the pituitary defects in pituitary specific Gli2 mutants. Pituitary

cell proliferation is expected to be restored by the rescue of the αGSU-Bmp2

transgene.

5.6.4 b Gli2 regulate Bmp2 transcription by direct binding sites

In a recent paper, Gli2 regulates Bmp2 gene transcription and thus

mediating osteoblast differentiation in response to Hh signaling [183]. They found

overexpression of Gli2 enhances Bmp2 promoter activity and mRNA expression

in osteoblast precursor cells. In contrast, genetic ablation of the Gli2 gene results

in significant inhibition of Bmp2 gene expression in osteoblasts. To test whether

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Gli2 protein binds to Bmp2 regulatory elements to regulate Bmp2 transcription in the pituitary, I propose the following experiments.

To determine whether Gli2 is sufficient to transactivate Bmp2 transcription,

Rathke’s pouch will be isolated to get primary cells at E10. Pituitary primary cells

will be cotransfected with luciferase Bmp2 promoter reporter constructs and Gli2

expression plasmid [183]. Bmp2 promoter activity will be determined by

luciferase level. Control experiment will be pituitary primary cells transfected with

only luciferase Bmp2 promoter reporter constructs but without Gli2 expression

plasmid. Another control assay will be applied on cells transfected with luciferase

Bmp4 promoter reporter or promoter reporters of other presumably unaffected

genes. Gli2 expression should stimulate Bmp2 promoter activity.

To determine whether Gli2 is required for Bmp2 transcription, zinc finger

DNA binding domain of Gli2 protein will be mutated. Pituitary primary cells will be

cotransfected with luciferase Bmp2 promoter reporter constructs and mutated

Gli2 expression plasmid. Bmp2 transcription level will be measure and compared

to WT Gli2 expression plasmid. Gli2 mutation should impede Bmp2 transcription

level.

To determine whether Gli2 bind to specific sites in the Bmp2 promoter, I

will use ChIP (chromatin immunoprecipitation) assay. Previous investigations

have demonstrated that Gli proteins transactivate target genes through a DNA

consensus sequence, GACCACCCA [186]. Moreover, promoter deletion assay,

chromatin immunoprecipitation and electrophoretic mobility shift assays, provided

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direct evidence that Gli2 physically interacts with the Bmp2 promoter [183]. I have constructed the IREShyg2-Gli2Fc plasmid which can express Gli2-fused Fc fragment and can be selected by hygromycin. Chromatin from pituitary primary cells transfected with Gli2 expression vector will be cross-linked and sonicated into small fragments. The protein-DNA complexes will be immunoprecipitated by incubation with IgG antibody and protein A agarose beads. De-cross-linked and purified DNA will be used as a template for PCR, using the primer sets to amplify two fragments of the mouse Bmp2 5’-flanking promoter. Total DNA before immunoprecipitation will be used as a positive control for PCR and the negative controls will be untransfected primary cells. It then can be determined whether

Gli2 protein physically binds with the Bmp2 promoter at the specific regions which contain putative Gli-responsive elements.

5.6.5 Further study of Shh signaling in the pituitary gland

My thesis work demonstrates an important role of Shh/Gli signaling in cell proliferation of the embryonic pituitary, but it remains to be determined whether

Gli defects can cause pituitary dysfunction after birth. Recent studies suggest the adult pituitary contains a cell population retaining early embryonic characteristics and expressing high mRNA level of Shh/Ptc1 [187]. Moreover, expression of

Shh/Ptc/Gli in adult human pituitary cells suggests Shh signaling may have a direct role in hormone secretion and cell proliferation in the adult brain [188, 189].

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In all FoxG1-Gli2 conditional knockouts I examined, the pituitary gland was still present with proliferation defects in the Rathke’s pouch. However, these

FoxG1-Gli2 mutant pups only survived one month after birth. Compared with controls, mutant mice had lower body weights. Clinical studies indicate increased mortality in preterm infants may be associated with hypoprolactinemia, which is a human disease caused by a deficiency in lactotrope-secreted hormones [190].

Reduced cell number of lactotropes was found in the FoxG1-Gli2 mutant embryos, but it is not clear whether hypoprolactinemia was the cause of death in these mutant pups.

To examine the circulating hormonal level of lactotrope-produced prolactin

(PRL), I will collect blood samples from WT and FoxG1-Gli2 mutant pups at P0 and P14 (early morning). Plasma (or serum) samples will be separated by centrifugation and assayed for PRL by means of traditional RIA

(radioimmunoasssy) methods [190]. By comparison of WT and FoxG1-Gli2 pups,

PRL level can predict whether embryonic removal of Gli2 in RP causes insufficient lactotropes. Other pituitary secreted hormones as ACTH, GH, LH,

TSH, will be assayed as well to determine whether loss of Gli2 function gives rise to postnatal hypopituitarism.

It was recently reported Shh and Gli1 can increase proopiomelanocortin

(POMC) transcription and ACTH production through cross-talk with the CRH

(corticotropin-releasing hormone) pathway suggest a continuous role of Shh/Gli in the adult pituitary [188]. High numbers of Shh immunopositive cells were found in the corticotroph population of the normal anterior pituitary [189]. Shh

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stimulates ACTH secretion and POMC (ACTH precusor) transcription in the mouse corticotroph cell line AtT-20 [188]. Moreover, Shh signaling has anti-

proliferative effects in AtT-20 cells [189]. However, Shh signaling mainly

promotes cell proliferation in the embryonic pituitary. As I found significant

reduction of corticotropes in the FoxG1-Gli2 mutant embryos, removal of Shh

signaling components would be expected to reduce ACTH secretion. Considering

the nature of AtT-20 line as pituitary tumor cells, Shh may function in a different

way in pituitary growth and hormone secretion within the mature or neoplastic

pituitary. Whether Shh/Gli signaling has a balancing role on pituitary cell

proliferation by antagonizing over-proliferation of pituitary neoplasia and

promoting the cell proliferation of pituitary hypoplasia in adults would be another

interesting topic.

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Appendix I Supplemental Materials for Chapter 2

Figure S1 Gli expression in Rathke’s pouch displayed an anteriorhigh to posteriorlow gradient. a-b Gli1 expression in the anterior Rathke’s pouch (left side) is higher than the posterior

Rathke’s pouch (right side). c-d Gli2 expression in the anterior Rathke’s pouch (left side) is higher than the posterior

Rathke’s pouch (right side).

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Appendix II Supplemental Materials for Chapter 3

Figure S1 Pituitary patterning is largely normal in conditional Gli mutants.

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Figure S2 No signicant changes in the expression of Bmp4 and Fgf8 were observed in FoxG1Cre;Gli2 mutant embryos at E9.5.

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Appendix III Copyright License for Journal Articles in Chapter 4

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Appendix IV Supplemental Materials for Chapter 4

Figure S1 Early Hh-responding cells occupy a broader region in the MGE than late Hh- responding cells. Cells were stained for ß-galactosidase 36 hours after TM injection.

Nkx2.1 was used as a marker for MGE progenitors. Cells labeled on E9.5 and E10.5

were found to nest inside the Nkx2.1+ domain while cells labeled on E11.5 were found

only near the edge of dorsal Nkx2.1 domain. White arrowheads indicate ß-

galactosidase+ cells. White dotted lines outline the edges of the MGE and LGE.

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Figure S2 Ventral telencephalic progenitors responding to Hh-signaling between E9.5

andE12.5 generate different subsets of cortical interneurons. (A) Double

immunofluorescence staining to show co-localization of ß-galactosidase with GABA, parvalbumin (PV) or somatostatin (SST) in 8-week-old adult cortex. (B) Progenitors responding to Hh-signaling between ~E9.5 and ~E12.5 generated different combinations of interneuron subtypes. Alhough the percentages of PV+ cells remained constant

(29.7%±3.8%, TM E8.5; 31.5%±4.8%, TM E9.5; 30.2%±9.3%, TM E11.5), the percentages of SST+ cells increased from 18.9%±9.6% (TM E8.5) to 26.5% and

25%±13.3% (TM E9.5 and TM11.5 respectively). Similarly, the percentages of calretinin

(CR+) cells increased from almost negligible (0% and 1.7%±1.4%, TM E8.5 and E9.5 respectively) to 9.1%±0.8% (TM E11.5). The number is expressed as mean percentage

± SEM. Total number of labeled cortical neurons counted were n=129 (TM E8.5), n=225

(TM E9.5), n=169 (TM E11.5). Scale bar: 50 µm (E).

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Figure S3 Progression of telencephalic phenotype of Gli2 mutants from E12.5 to E18.5.

Telencephalic sections were stained with Hemotoxylin and Eosin. At E14.5, 1 out of four

Gli2 mutants showed enlarged ventricles. However all mutant embryos (4/4) showed enlarged ventricles at E18.5.

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