Notch-Signaling in Retinal Regeneration and Müller Glial Plasticity

Home , Bipolar neuron, Gliogenesis, Neurite, Photoreceptor cell, Retinal regeneration

Notch-Signaling in Retinal Regeneration and Müller glial Plasticity

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Kanika Ghai, MS

Neuroscience Graduate Studies Program

The Ohio State University

2009

Dissertation Committee:

Dr. Andy J Fischer, Advisor

Dr. Heithem El-Hodiri

Dr. Susan Cole

Dr. Paul Henion

Kanika Ghai

2009

ABSTRACT

Eye diseases such as blindness, age-related macular degeneration (AMD),

diabetic retinopathy and glaucoma are highly prevalent in the developed world,

especially in a rapidly aging population. These sight-threatening diseases all involve

the progressive loss of cells from the retina, the light-sensing neural tissue that lines

the back of the eye. Thus, developing strategies to replace dying retinal cells or

prolonging neuronal survival is essential to preserving sight. In this regard, cell-based

therapies hold great potential as a treatment for retinal diseases. One strategy is to

stimulate cells within the retina to produce new neurons. This dissertation elucidates

the properties of the primary support cell in the chicken retina, known as the Müller

glia, which have recently been shown to possess stem-cell like properties, with the potential to form new neurons in damaged retinas. However, the mechanisms that govern this stem-cell like ability are less well understood. In order to better understand these properties, we analyze the role of one of the key developmental processes, i.e., the Notch-Signaling Pathway in regulating proliferative, neuroprotective and regenerative properties of Müller glia and bestow them with this plasticity.

The first part of this dissertation is a description of embryonic studies that verify the use of a pharmacological inhibitor of Notch-signaling, DAPT, as well as

RNA interference molecules that block downstream effectors of Notch – Hes1 and

Hes5. We find that inhibition of γ-secretase activity associated with Notch and

ii silencing of the bHLH effectors Hes1 and Hes5 have distinctly different outcomes on

cell-fate specification of cultured chicken retinal progenitors. Further, our studies

reveal that Notch-signaling plays a limited but important role during retinal

regeneration. Components of the Notch-signaling pathways are transiently

upregulated in proliferating Müller glia after damage in a chicken retina and blocking

Notch after damage enhances some neural regeneration from glial-derived progenitors.

In the second part of this dissertation, we analyze the role of the Notch pathway in the postnatal retina in the absence of damage. We find that components of the Notch-signaling pathway are expressed at low levels in most Müller glia in undamaged retina. Further, Notch-signaling influences the phenotype and function of

Müller glia in the mature retina; low levels of Notch-signaling diminish the neuroprotective capacity of Müller glia, but are required to maintain their ability to

become progenitor-like cells. We also find that there is cross-talk between Notch and

MAPK pathways – FGF2, a secreted protein that activates the MAPK pathway, also induces the expression of Notch pathway genes. Further, active Notch-signaling is required for the FGF2-mediated accumulation of p38 MAPK and pCREB in Müller glia. Our data indicate that Notch-signaling is down-stream of and is required for

FGF2/MAPK- signaling to drive the proliferation of Müller glia.

The third part of this dissertation describes the patterning of the immature zone of cells present at the retinal margin, called the circumferential marginal zone

(CMZ). Our data indicates that there is a gradual spatial restriction of this zone of progenitor cells during late stages of embryonic development and that there are

iii regional differences in the maturity of cells within the CMZ. Further, we find that retinal neurons adjacent to the CMZ in far peripheral regions of the temporal retina

remain immature and differentiate far more slowly compared to the neurons in central

regions of the retina and that the microenvironment at the periphery of the retina that

promotes the persistence of a zone of retinal progenitors may also keep some types of

neurons immature for extended periods of time.

The last part of this dissertation describes the morphological and mechanistic

properties of a unique subset of interneurons we discovered in the chicken retina,

called the serotonin-accumulating bipolar cells. Even though serotonin is synthesized

by amacrine cells, another type of interneuron present in the retina, it transiently

accumulates in this distinct type of bipolar neuron. The accumulation of endogenous

or exogenous serotonin by bipolar neurons is blocked by selective reuptake inhibitors.

Further, inhibition of monoamine oxidase (A) prevents the degradation of serotonin in bipolar neurons, suggesting that MAO(A) is present in these neurons. Our data

indicates that serotonin-accumulating bipolar neurons perform glial functions in the

retina by actively transporting and degrading serotonin that is synthesized in

neighboring amacrine cells.

Taken together, the data presented in this dissertation furthers understanding

of Müller glial plasticity. This information could be applied to stimulating neural

regeneration, harnessing Müller glia as a localized source of stem cells intrinsic to the

retina, developing pharmacological therapies targeted to the glia and countering

neuronal death in sight-threatening diseases. Additionally, our studies on serotonin-

accumulating bipolar cells have implications for understanding the mechanisms of

iv melatonin biosynthesis and retinal circadian rhythms, dysfunctions of which lead to photoreceptor degeneration and loss of vision.

DEDICATION

Dedicated to my dear grandfathers I.S. Chowdhary and R.L. Ghai

ACKNOWLEDGEMENTS

As I write this dissertation, I have several people to thank for supporting me through the long drawn-out ordeal that is the PhD. Undoubtedly on top of this list are my parents, brother, and grandparents who have always been my pillars of strength. I can’t thank them enough for keeping me emotionally grounded and for being my most ardent cheerleaders through this process.

I am grateful to my mentor, Dr. Andy Fischer for helping me develop my professional skills and teaching me how to be a better scientist. Andy’s passion for

science, coupled with his ability to multi-task experiments, grants/paper writing,

family, administrative duties and yet, maintaining his great sense of humor and being

there whenever I needed him is inspiring and mind-boggling. I couldn’t have asked

for a better advisor through graduate school.

Other important people I am grateful to: Dr. Georgia Bishop and Dr. Ning

Quan for their indispensable support and advice over the years; the fantastic members

of my Candidacy and Defense Committees including Dr. Heithem El-Hodiri, Dr. Paul

Henion, Dr. Susan Cole and Dr. Dick Burry for their constructive criticism about my

research projects; Dr. Vidita Vaidya at TIFR, Mumbai, who introduced me to the

exciting world of neuroscience; the Professors at the Life Science Department, St.

Xavier’s College, Mumbai especially Dr. Radiya-Pacha Gupta, Dr. Sujayakumari, Dr.

vii Donde, Dr. D’Silva, Dr. Mangalore and my excellent Biology teacher in high school

Ms. Vasanthy.

My dear husband, Rajat, thank you for waiting patiently for me to finish, encouraging and pushing me, and enduring a long-distance marriage! A big thank you to all my friends who know what it’s like to be in graduate school, especially Sirsha,

Bharath, Pushkar, Kavitha, Harini, Shwetank, Trupti and Arnaz – you were there

when I needed you and I am grateful for that. I also want to acknowledge some

colleagues and friends who have inspired me in little ways, including Khalid, Nupur,

Greg, Sharmila, Richa, Cynthia, Kristen, Leah, Leena and Litty.

It’s been such a pleasure working in a lab where doing science is a fun and

creative learning process. Chris deserves special mention here – my dissertation

would not have been possible without his excellent help with experiments and his

enthusiasm to work tirelessly and on weekends. Many thanks to Jennifer for patiently

walking me through lab techniques, sharing graduate school woes and all those

scientific discussions. I would also like to acknowledge the support and friendship of

other lab members including Eric, Melissa, Pat, Ami and Rachel.

In the end, I am grateful that I was given the opportunity to be part of the

Neuroscience Department at OSU. Columbus has been a welcoming home away from

home for me and I shall cherish the memories associated with this University and the

city for years to come.

viii

VITA

1980………………………………………...Mumbai, India

2001………………………………………...BS, Life Sciences, St. Xavier’s College, Mumbai

2003………………………………………..MS, Life Sciences, Mumbai University

2003 – 2009……………………………...... Graduate Research Associate Neuroscience Graduate Studies Program The Ohio State University

PUBLICATIONS

1. Ghai K, Zelinka C, Fischer AJ. Serotonin released from amacrine neurons is scavenged and degraded in bipolar neurons in the retina. J. Neurochem. 2009 Oct; 111(1):1-14

2. Ghai K, Stanke JJ, Fischer AJ. Patterning of the circumferential marginal zone of progenitors in the chicken retina. Brain Res. 2008 Feb 4;1192:76-89

3. Fischer AJ, Stanke JJ, Ghai K, Scott M, Omar G. Development of bullwhip neurons in the embryonic chicken retina. J Comp Neurol. 2007 Aug 1;503(4):538-49.

FIELDS OF STUDY

Major Field: Neuroscience

TABLE OF CONTENTS

Abstract …………………………………………………………………………..…...ii

Dedication ……………………………………………………………...…………….vi

Acknowledgments …………………………………………………………...……...vii

Vita ………………………………………………………...…………………………ix

List of Tables………………………………………………………….……………..xii

List of Figures ………………………………………………………………………xiii

List of Abbreviations …………………………………………...…………………..xvi

Chapters

1. General Introduction …………………………..…………………………………. .1 A brief insight……………………………………….……..………………………….1 The retina – structure, function and neural connectivity……………...…………..…..2 Müller glia – the Yin and the Yang…………………………………………..……….3 Retinal development and Müller glial plasticity – the role of cell-signaling pathways……………..…...……………………...…………………..…6 Notch-signaling pathway – a biological superstar………….…………………..…..…8 Notch-signaling Pathway– structure, function and mechanisms….…………..………9 Notch in the nervous system – cell-fate decisions and beyond………………...……11 Notch-signaling and regeneration……………………………………………....……14 Non-traditional roles of Notch in the nervous system…………………………....….15 Notch-signaling – beyond the nervous system……………………………..……..…16 Patterning the circumferential marginal zone…………………………………..…....16 Serotonin accumulating bipolar cells in the retina – novel cells, novel role…...... 17

2. Notch-signaling regulates aspects of regeneration in the adult chicken retina…....19 Introduction ………………………………………………………..……………...... 19 Materials and methods……………………………………………..……..……..…...22

x Results ………………………………………………………………..…..……..…...33 Discussion……………………………………………………………………..…...... 50 Conclusions…………………………………………….…………………………….58

3. Notch-signaling influences neuroprotective and proliferative properties of Müller glia...... …..…...59 Abstract……………………………………………………………..………………..59 Introduction ……………………………………………………………………....….60 Materials and methods…………………………………………………………....….62 Results …………………………………………………………………………….....71 Discussion………………………………………………………………………...... 90 Conclusions………………………………………………………..………………..102

4. Patterning of the circumferential marginal zone of progenitors in the chicken retina…………………………………………...……………………103 Abstract……………………………………………………………………..…..…..103 Introduction …………………………………………………………...…..………..104 Materials and methods………………………………………….…………..……....106 Results ……………………………………………………………………..……….109 Discussion…………………………………………………………………....……..129 Conclusions…………………………………………………………………..……..134

5. Serotonin released from amacrine neurons is scavenged and degraded in bipolar neurons in the retina…………..…………...……..….………………….136 Abstract…………..…………………………………………………………….…...136 Introduction …………………………………………….……………………….….137 Materials and methods…………………………………………………………..….139 Results ……………………………………………………………………………...144 Discussion…………………………………………………………………...…...…162 Conclusions………………………...…………………….…………………………166

6. Summary and Future Directions……..………………...……………..………….167 Summary of findings………………………………………………...... 167 Future directions for studies on Müller glia……………………………...... 170 - Introduction……………..………………………………………..…………..…...170 - Notch and the effects of other signaling pathways……….……………...... 171 - Notch ligands – where are they expressed?...... 172 - Implications of Notch-mediated neuronal survival…………………………..…...173 - Epigenetic mechanisms may control glial plasticity………………..………..…...175 Future directions for studies on serotonin-accumulating bipolar cells……..………177 Concluding remarks……………………………………………………………...…178

References…………………………………………………………………………..179

LIST OF TABLES

Table 2.1 Primer sets used for standard PCR followed by dsRNA synthesis..... 24

Table 2.2 Double-stranded oligo sequences used for shRNA plasmid

synthesis……………………………………………………………...27

Table 3.1 Primer sets used for standard and real-time PCR……………..……..66

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LIST OF FIGURES

Figure 1.1 Structure of the eye and retina.……………………………………..…3 Figure 1.2 Overview of the Notch-signaling Pathway and its inhibitors used in the current studies ………………………………………...…11

Figure 2.1 Map of the BLOCK-iT RNAi Entry Vector……………………...... 26

Figure 2.2 Paradigm used for RNA-interference studies in embryonic retinal cell cultures.… ……………….…...……………………….....29 Figure 2.3 Notch pathway gene expression increases after damage ……………34 Figure 2.4 Proliferating cells express cNotch1 after damage……………………36 Figure 2.5 Proliferating Müller glia express cNotch1 after damage……………38 Figure 2.6 DAPT increases percentage of cells differentiating as (calretinin+) amacrine and horizontal cells and (Islet1+) amacrine and bipolar cells, but not (visinin+) photoreceptors or (Lim3+) bipolar and horizontal cells, without depleting (BrdU+) proliferating progenitors……….…40 Figure 2.7 Transfected embryonic retinal cell cultures………………………….42 Figure 2.8 Effects of RNA interference-mediated silencing of Hes genes has varied effects on cell-fate…………………………………………….43 Figure 2.9 DAPT blocks Notch pathway gene expression after damage………..45 Figure 2.10 DAPT increases the number of proliferating cells in the central retina after damage………………………………………..….47 Figure 2.11 DAPT enhances neuronal differentiation in the central regions of the retina, and glial differentiation in temporal regions…………..49 Figure 2.12 Basic structure of bHLH-O subfamily domain structures …………..53 Figure 3.1 Components of the Notch-signaling pathway are expressed at low levels in the postnatal chicken retina………………73 xiii Figure 3.2 Inhibition of Notch-signaling reduces the expression of components of the Notch-signaling pathway in undamaged postnatal chicken retina…………………………………75 Figure 3.3 Inhibition of Notch-signaling prior to retinal injury decreases cell death and increases neuronal survival……………………...……78 Figure 3.4 cNotch1 is upregulated by proliferating Müller glia in retinas treated with insulin and FGF2……………………………………..…80 Figure 3.5 Notch and related genes are upregulated in retinas treated with insulin and FGF2………………………………………..82 Figure 3.6 FGF2 induces the proliferation of Müller glia in injured retinas…….85 Figure 3.7 Inhibition of Notch-signaling reduces levels of p38 MAPK and pCREB in Müller glia…………………………………………….….87 Figure 3.8 Inhibition of Notch-signaling blocks FGF2-induced proliferation of Müller glia in NMDA-damaged retina………………………………89 Figure 3.9 Model for the influence of the Notch and MAPK Pathways on retinal Müller glia……………………………………………..……101 Figure 4.1 BrdU-birthdating of the cells in the far periphery of the retina…….112 Figure 4.2 PCNA-expressing cells are gradually confined to the CMZ during late stages of embryonic development……………………...114 Figure 4.3 Transitin-expression is gradually confined to the CMZ during late stages of embryonic development……………………...117 Figure 4.4 N-cadherin is highly expressed in the far peripheral retina, CMZ and ciliary body of both developing and postnatal retina, and gets restricted over time in far peripheral regions of the retina…………………………………………………………120 Figure 4.5 Early markers of neuronal differentiation are expressed by cells directly adjacent to the embryonic and postnatal CMZ…………….123 Figure 4.6 Synaptophysin and protein kinase C are expressed by neurons at increasing levels with increasing distance away from the CMZ…...126 Figure 4.7 Photoreceptors and calbindin-expressing neurons remain immature

xiv in far peripheral regions of the temporal retina…………………….128 Figure 4.8 Schematic summary of the onset of different neuronal markers and gradient of maturity that persists in peripheral postnatal retina…….132 Figure 5.1 Exogenous serotonin transiently accumulates in bipolar neurons….146 Figure 5.2 Characterization of SAB cells………………………………………150 Figure 5.3 Bipolar neurons accumulate exogenous serotonin even when amacrine neurons are destroyed…………………………….………152 Figure 5.4 Depolarization stimulates depletion from amacrine neurons and accumulation of serotonin by bipolar neurons……………………...155 Figure 5.5 Degradation of serotonin in bipolar neurons is prevented by an MAO(A) inhibitor (clorgyline) but not by an MAO(B) inhibitor (pargyline)………………………………………………...158 Figure 5.6 Immunoreactivity and in situ hybridization indicate expression for vesicular monoamine transporter 2 (VMAT2) in amacrine cells in the retina…………………………………………………………161

LIST OF ABBREVIATIONS

ascl achaete-scute complex-like bHLH basic helix loop helix °C degrees Celsius CREB cAMP response element binding CNTF Ciliary neurotrophic factor DAPT N-[(3,5-Difluorophenyl)acetyl]-L-alanyl-2-phenyl]glycin e-1,1- dimethylethyl ester DMEM Dulbecco’s modified Eagle medium DMSO Dimethyl sulfoxide dsRNA double-stranded RNA Egr Early growth response ERK Extracellular signal regulated kinase EGF Epidermal growth factor FGF Fibroblast growth factor γ gamma GCL Ganglion cell layer GFP Green Fluorescent Protein Hes Hairy/enhancer of split Hey Hairy/Enhancer of Split related with YRPW IGF Insulin-like growth factor INL Inner nuclear layer IPL Inner plexiform layer KCl Potassium chloride LB Luria Broth

xvi µl microlitre µm micrometer MAOA Monoamine oxidase A MAPK Mitogen-activated protein kinase NMDA N-methyl-D-aspartate OLM Outer limiting membrane ONL Outer nuclear layer OPL Outer plexiform layer PCR Polymerase Chain Reaction PKC Protein kinase C RPE Retinal pigmented epithelium SAB Serotonin-accumulating bipolar SERT Serotonin transporter SEM Standard error of mean Shh Sonic hedgehog siRNA short interfering RNA shRNA short hairpin RNA SSC Standard sodium citrate TH Tyrosine hydroxylase TUNEL Terminal deoxynucleotidyl transferase dUTP nick end labeling VMAT2 Vesicular monoamine transporter 2

xvii

CHAPTER 1

General Introduction

A brief insight

This dissertation is focused on studies done in the retina, neural tissue lining the back of the eye. The eye has been an object of human curiosity since the time of

Aristotle (4th century B.C). However, it took about 1900 years for Johannes Kepler to

establish that the retina is responsible for vision. Since the 17th century, the

complexities of the retina have baffled many scientists and philosophers; a quote by

Santiago Ramón y Cajal sums it up best –

“In the study of this membrane [the retina] I for the first time felt my faith in

Darwinism (hypothesis of natural selection) weakened, being amazed and confounded

by the supreme constructive ingenuity revealed not only in the retina and in the

dioptric apparatus of the vertebrates but even in the meanest insect eye. ... I felt more

profoundly than in any other subject of study the shuddering sensation of the

unfathomable mystery of life.”

A quote like that coming from one of the greatest stalwarts in neuroscience is

both humbling and inspiring. In the last 100 years since the original “Structure of the

Mammalian Retina” by Cajal was published, much has been learnt about the

extraordinary function and mechanisms of cells that construct the retina. However,

1 several questions remain unanswered about this tissue that allows our eyes to be

windows to the world.

The retina – structure, function and neural connectivity

The retina is a complex multilayered structure that is an outward extension of

the brain. It serves as a good model system for studying neural function in an intact

system due to its accessibility and the fact that it can be easily stimulated by

projecting light onto it. The retina is organized into 7 distinct layers as shown in Fig.

1.1. The cells in the retina include neurons, rod and cone photoreceptors in the

photoreceptor layer (PRL), the interneurons - bipolar cells and amacrine cells in the

inner nuclear layer (INL) and the ganglion cells in the ganglion cell layer (GCL)

whose axons travel through the optic nerve to higher brain structures and eventually

the visual cortex in the occipital lobe where visual information is processed (Kandel

et al. 2000). The retina is arranged as an inverted structure with light entering from

the ganglion cell layer and penetrating through all intermediary layers before reaching

the photoreceptors. Apart from the neurons, the primary glial cell type in the retina is the Müller glia, with cell bodies in the INL and processes spanning the entire breadth of the retina (Livesey and Cepko 2001; Marquardt 2003). Other glial cell types present in the retina include astrocytes and other cells of non-neuroectodermal origin such as microglia.

Figure 1.1: Structure of the eye and retina. The retina is neural tissue present at the back of the eye that has seven distinct layers containing 6 different types of neurons and one glial cell type. The neuronal cell types are: rod and cone photoreceptors (red nuclei), horizontal cells (green nuclei), bipolar cells (pink nuclei), amacrine cells (blue nuclei) and ganglion cells (orange nuclei). The main glial cell type is the Müller glia (yellow nuclei). Abbreviations: RPE – retinal pigmented epithelium; PRL – photoreceptor layer; ONL – outer nuclear layer; OPL – outer plexiform layer; INL – inner nuclear layer; IPL – inner plexiform layer; GCL – ganglion cell layer; NFL – nerve fiber layer. Eye structure modified from http://thefutureofthings.com/articles/57/shedding-light-on-blindness.html

Müller glia – the Yin and the Yang

The retina is involved in the pathology of several eye diseases such as age- related macular degeneration (AMD), diabetic retinopathy and glaucoma, highly prevalent diseases in the developed world. With a rapidly aging population, the number of Americans suffering from impaired vision is projected to almost double by

2020 (Rein et al. 2006). These sight-threatening diseases all involve the progressive loss of cells from the retina (Bonnel et al. 2003; Bringmann and Reichenbach 2001).

For example, in glaucoma, there is a progressive loss of ganglion cells and a 3 deterioration of optic nerve fibers. AMD and diabetic retinopathy involve a loss of retinal cells in the macula, particularly the photoreceptors and retinal pigmented

epithelial cells. Photoreceptor degeneration is also hallmark in the pathogenesis of

retinal detachments and retinitis pigmentosa.

Thus, developing strategies to replace dying retinal cells or prolonging neuronal survival is essential to preserving sight. In this regard, cell-based therapies

hold great potential as a treatment for retinal diseases. One strategy is to stimulate

cells within the retina to produce new neurons. In several species, Müller glia have

proven to be a promising candidate for cell-based therapies, and possess stem cell-

like properties, with the potential to form new neurons in damaged retinas (Fischer

and Reh 2001). The idea that glia may possess progenitor-like qualities is not new.

Indeed, in the central nervous system, several studies have shown that radial glia can

generate neurons in the developing neocortex (Anthony et al. 2004; Ever and Gaiano

2005; Gotz and Barde 2005; Malatesta et al. 2008) and are a source of adult neural

stem cells in the mammalian telencephalon (Doetsch 2003; Merkle et al. 2004). Glial

cells in the postnatal and adult CNS can be manipulated to undergo the initial steps of

neurogenesis both in vitro and, after brain injury, in vivo (Buffo et al. 2005; Heins et

al. 2002; Kondo and Raff 2000; Laywell et al. 2000; Noctor et al. 2001; Zheng et al.

2004).

Astrocytes have also been shown to generate neurons from postnatal cortex in

the presence of Pax6 (Heins et al. 2002). Pax6 plays a crucial role in the generation of

neurons from astroglial stem cells in the adult sub-ependymal zone (SEZ) of the

telencephalon (Hack et al. 2005). Astrocytes from the subventricular zone (SVZ)

4 have also been shown to give rise to neurons in the olfactory bulb (Doetsch et al.

1999). Müller glia in the retina, being equivalent to astrocytes in the brain, exhibit

similar stem cell-like characteristics. Müller glia have been shown to be proliferative

and regenerative, producing new neurons in several species including fish (Fausett

and Goldman 2006; Fausett et al. 2008; Fimbel et al. 2007; Kassen et al. 2009;

Raymond et al. 2006; Thummel et al. 2008; Yurco and Cameron 2005), chickens

(Fischer and Reh 2001; Hayes et al. 2007) and mammals (Karl et al. 2008; Ooto et al.

2004).

Unfortunately, the plasticity of Müller glia that bestows them with stem cell-

like properties may also contribute to the pathology of retinal diseases. Müller glial

proliferation occurs in retinal detachments and proliferative retinopathies (Bringmann et al. 2006; Bringmann and Reichenbach 2001; Fletcher et al. 2008). Faulty glutamate degradation by Müller glia is seen prior to photoreceptor degeneration in retinitis

pigmentosa (Fletcher and Kalloniatis 1996). Vascular Endothelial Growth Factor

(VEGF) expression in Müller glia is increased during neovascularization in diabetic

patients, contributing to hypoxia-induced pathological angiogenesis (Aiello et al.

1994; Jingjing et al. 1999). Almost every disease of the retina is associated with reactive Müller cell gliosis which may either support the survival of retinal neurons or accelerate glial scar formation and the progress of neuronal degeneration. Thus, the goal of our research is to minimize these damaging effects of Müller glia, while at the same time, harnessing their plastic phenotype to promote regeneration and neuronal survival. However, the mechanisms that govern this stem cell-like ability are not well understood. In order to better understand these properties, Chapters 2 and 3 of this

5 dissertation describe the role of an important signaling system i.e., the Notch- signaling Pathway in regulating proliferative, protective and regenerative properties of Müller glia and bestowing them with this plasticity.

Retinal development and Müller glial plasticity – the role of cell-signaling pathways

Many intrinsic and extrinsic factors and signals control the development of the retina. Extrinsic signals are mediated via growth factors and intrinsic cues mediated by transcription factors such as Paired Homeobox 6 (Pax6), Mammalian Atonal

Homologue 5 (Math5) and Sine oculus Homeobox 3 (Six3) Growth factors such as

Fibroblast Growth Factor 2 (FGF2), Bone Morphogenetic Protein 4 (BMP4) and

Sonic Hedgehog (Shh) are secreted from cells, bind to receptors at the cell surface and activate second messenger signaling cascades. In early eye development,

Transforming Growth Factor (TGF)ß-like and FGF signaling act antagonistically to specify the retina pigmented epithelium (RPE) and neural retina (NR) precursors from the optic vesicle (Martinez-Morales et al. 2004), whereas opposing gradients of

BMP and Shh control the establishment of dorsoventral polarity of the optic cup

(Yang 2004). Additionally, different cell types within the neural retina are generated in a spatiotemporally controlled process that involves many extracellular signals such as Wnt, Shh, BMP and FGF (Yang 2004). These signaling molecules also control eye field formation, onset of retinal neurogenesis and control of retinal cell numbers

(Esteve and Bovolenta 2006).

Multipotent neural stem cells/progenitors produce all types of neurons and the

Müller glia within the retina (Turner and Cepko 1987; Turner et al. 1990). Retinal

6 histogenesis proceeds in a fixed pattern that is largely conserved across species

(Livesey and Cepko 2001; Marquardt 2003). The first cell type to be born in the retina is the ganglion cell, followed by overlapping waves of differentiation of cone photoreceptors, amacrine cells, bipolar cells, horizontal cells and rod photoreceptors.

The last cell type to emerge is the Müller glia. Combinations of homeodomain and

bHLH transcription factors determine the fates of the retinal cell types. As discussed ahead, Notch-Delta signaling is also important in this process of retinal histogenesis.

Notch-signaling is high in the developing retina and decreases over time as retinal

cells differentiate into postmitotic neurons. During this process of histogenesis,

secreted factors such as FGF, Epidermal Growth Factor (EGF), Insulin-like Growth

Factor (IGF) and Shh regulate the proliferation and survival of retinal progenitors as

well as influence cell-fate decisions. For example, Shh signaling regulates retinal

ganglion cell production (Zhang and Yang 2001a; Zhang and Yang 2001b), promotes

progenitor proliferation and promotes survival of rods and Müller glia (Jensen and

Wallace 1997; Levine et al. 1997). Wnt2b signaling controls retinal cell

differentiation and determines peripheral fates of the chick eye (Cho and Cepko 2006;

Kubo et al. 2003). FGF signaling contributes to positioning the presumptive retinal

cells in the right place at the right time for the onset of retinal neurogenesis (Moody

2004). Further, EGF and FGF signaling stimulate neurogenesis from progenitor-like

cells in the mature ciliary body (Fischer and Reh 2003).

The same signaling pathways that sculpt the retina have also been found to

regulate Müller glial plasticity in a mature retina. Shh was shown to promote the stem

cell potential of Müller glia in the mammalian retina (Wan et al. 2007). Notch-

7 signaling was shown to be upregulated in Müller glia in a damaged retina and alters glial-derived progenitor cell-fates (Hayes et al. 2007). This report has also been compared to our observations regarding the role of Notch in retinal regeneration in

Chapter 2 of this dissertation. In the rodent retina, Wnt-signaling has been shown to stimulate the proliferation and generation of new retinal neurons from Müller glia in response to NMDA-induced damage (Osakada et al. 2007). In the chicken retina,

FGF, IGF and Ciliary Neurotrophic Factor (CNTF) expression levels were shown to increase after neural damage (Fischer et al. 2004) and these factors caused an increase in glial expression of GFAP and filamentous neuronal proteins. Further, insulin and

FGF2 treatment in the retina activate a neurogenic program in Müller glia (Fischer et al. 2002). These factors bind to receptor tyrosine kinases that activate the Mitogen- activated protein kinase (MAPK)-signaling pathways in Müller glia. Activation of

MAPK-signaling was shown to initiate Müller glial proliferation and protect retinal neurons against excitotoxicity. In Chapter 3, I have discussed my findings regarding the interactions between the Notch- and MAPK-signaling pathways in regulating

Müller glial neuroprotection and proliferation.

Notch-signaling pathway – a biological superstar

One of the signaling pathways that is crucial in the process of retinal development is the Notch pathway. At a symposium held at Janelia Farms, Virginia earlier this year, Dr. Spyros Artavanis-Tsakonas from Harvard Medical School, pioneer in Notch biology, said this –

8 “There are two kinds of people in the world, those that work on Notch and those who

haven't yet” – a simple and yet profound statement, highlighting the incontrovertible

importance of the Notch-signaling Pathway to biology. The gene encoding the Notch

receptor was discovered in fruitflies about 90 years ago where a partial loss of function (haploinsufficiency) resulted in irregular structures or notches in the wing margin of flies (Artavanis-Tsakonas et al. 1999; Mohr 1919). Comprehensive studies done by Poulson in the 1930s on embryonic lethal loss-of-function mutations in the

Notch locus determined that these mutations led to a switch in cell-fates – cells destined to become epidermis now gave rise to neural tissue (Artavanis-Tsakonas et al. 1999; Poulson 1937; Wright 1970). Further, molecular analysis of the Notch locus done in the 1980s and 1990s revealed that Notch homologs were present in a diverse range of species, from flies to worms and from frogs to humans (Artavanis-Tsakonas et al. 1991; Theodosiou et al. 2009). Despite the striking diversity of species in which

Notch was found to be present, the structure and function of this gene was highly conserved (Weinmaster 1997) – the Notch receptor along with its ligands Delta,

Jagged or Serrate, initiates signaling events that ultimately directs cell-fate and is essential during development of many organ systems, including the nervous system.

Notch-signaling Pathway– structure, function and mechanisms

Molecules involved in the Notch-signaling Pathway have been identified and extensively characterized in several species, including chicken (Fleming et al. 1997;

Henrique et al. 1995; Myat et al. 1996). Notch encodes a 300 kDa single-pass transmembrane receptor protein that has a large extracellular domain containing 36

9 EGF-like repeat domains and three cysteine-rich Notch/LIN-12 repeats. Six tandem ankyrin repeats,a glutamine-rich domain (opa), and a PEST sequence are found within the intracellular domain (ICD) (Artavanis-Tsakonas et al. 1999; Wharton et al.

1985). The ligands for Notch are found on adjacent cells during development, and

include two single-pass transmembrane proteins that belong to the Delta and Jagged

gene families in vertebrates. Receptor-ligand interactions trigger the γ-secretase

dependent cleavage of the Notch ICD (NICD). The NICD gets translocated into the

nucleus where it binds to the major downstream effector of Notch-signaling,

Suppressor of Hairless, also called CBF1, Su(H) and Lag1 (CSL) and the co- activator, mastermind. This NICD/CBF1 activator complex upregulates expression of basic helix-loop-helix (bHLH) transcriptional repressors of the hairy and enhancer of split (HES) and HES related (HEY) genes, which in turn antagonize the expression of proneural genes such as Achaete-scute (Asc1) and neurogenin (Louvi and Artavanis-

Tsakonas 2006).

Studies described in Chapters 2 and 3 of this dissertation heavily rely on the use of a small-molecule inhibitor of γ-secretase activity, DAPT, which effectively blocks Notch-signaling by inhibiting the γ-secretase activity associated with the activated receptor. γ-secretase inhibitors such as DAPT block the activity of presenilin/γ-secretase complex which cleave Amyloid Precursor Protein (APP) to form beta-amyloid peptides, which in turn form plaques in Alzheimer’s disease

(Roberson and Mucke 2006). Thus these compounds were developed as a means to treat Alzheimer’s disease (Dovey et al. 2001). Additionally, retinal cell culture studies described in Chapter 2 have utilized RNA-interference molecules to the downstream

10 effectors of Notch-signaling, Hes1 and Hes5, to inhibit specific effector-driven effects

on progenitor cell-fates. A simplified schematic of the Notch pathway and inhibitors

used in this study are shown in Fig. 1.2.

Figure 1.2: Overview of the Notch-signaling Pathway and its inhibitors used in the current studies. The Notch receptor binds to the ligand Delta on adjacent cells. This binding triggers the γ-secretase dependent cleavage of the Notch intracellular domain (ICD) which gets translocated into the nucleus where it binds to the major downstream effector of Notch-signaling Suppressor of Hairless (CSL). This NICD/CSL activator complex upregulates expression of bHLH transcriptional repressors of the hairy and enhancer of split (HES) and HES related (HEY) family genes, which in turn antagonize the expression of proneural genes such as Achaete- scute (Asc1) and neurogenin. DAPT inhibits γ-secretase activity associated with Notch-signaling, thereby blocking the transcription of its effectors. RNA interference molecules (siRNA) to the Notch effectors inhibit Hes1 and Hes5 specifically but do not directly inhibit upstream Notch activation.

Notch in the nervous system – cell-fate decisions and beyond

Notch-signaling is one of the essential developmental mechanisms that sculpt the complexity of the nervous system, including the retina. The most well studied role

11 for this pathway is in the maintenance of progenitors during development. The idea

that activated Notch inhibits cell-fate commitment was first proposed by Coffman et al who demonstrated that Notch homologs delay differentiation in Xenopus embryos

(Coffman et al. 1993). Soon after, Dorsky et al showed that Xotch (Xenopus

equivalent of Notch) inhibits cell differentiation in the Xenopus retinal cells, causing

them to maintain a neuroepithelial morphology (Dorsky et al. 1995). In the

developing forebrain, Notch-1 protein is asymmetrically inherited by the neuronal

daughter cell of mitotically active cortical progenitors (Chenn and McConnell 1995).

Several misexpression studies of Hes1, Hes5 and other effectors of the Notch pathway, and expression of constitutively active forms of Notch in the developing mammalian nervous system have been shown to inhibit neuronal differentiation

(Jadhav et al. 2006b; Magavi et al. 2000; Morrison et al. 2000; Vetter and Brown

2001; Yaron et al. 2006).

Some studies have shown that cells that are kept mitotically active for longer

durations due to overactive Notch-signaling eventually form radial glia (Louvi and

Artavanis-Tsakonas 2006). Further, transient Notch-signaling instructs neural crest progenitors to switch their competence from neurogenesis to gliogenesis (Morrison et al. 2000). Ectopic expression of Hes5 in postnatal mouse retinae promotes gliogenesis at the expense of neurogenesis (Hojo et al. 2000). Thus the vast body of literature suggests that the permissive effects of Notch in the CNS promote a proliferative progenitor-like phenotype at early stages of development, while later on, Notch- signals are instructive for producing late-born cell types such as Müller glia and astrocytes (Chenn 2009; Grandbarbe et al. 2003; Vetter and Moore 2001).

12 In the retina, the role of Notch in cell-fate decisions and in promoting glial cell-fates has been widely studied (Bernardos et al. 2005; Furukawa et al. 2000;

Gaiano and Fishell 2002; Jadhav et al. 2006a; Morrison et al. 2000; Vetter and Moore

2001; Wu et al. 2001). Notch and Delta engage in a specific form of signaling called lateral inhibition (Rapaport and Dorsky 1998), wherein, both Notch and Delta are expressed by adjacent members of uncommitted cells. When Notch activity is induced in a cell, it simultaneously inhibits adjacent cells from responding to Notch.

In a feedback loop, Notch activation inhibits the expression of Delta in that particular cell, whereas adjacent cells upregulate Delta. Thus, the Notch-expressing cell remains undifferentiated whereas adjacent cells can acquire a differentiated phenotype.

Further, Delta-Notch-signaling regulates competence by keeping progenitors from responding to differentiation cues (Rapaport and Dorsky 1998).

Notch-signaling is an integral component of the Competence Model put forth by Livesey and Cepko (Livesey and Cepko 2001) which explains how progenitors are restricted to produce certain cell types during certain times. Progenitor competence is regulated by intrinsic rather than extrinsic factors, as determined by experiments in retina model systems, where early progenitors were put into a late environment and still produced only early cell types. Environment on the other hand may determine the cell-fate produced at that particular point of time as progenitors are competent to produce multiple cell types. Livesey and Cepko propose that intracellular signaling via Notch may help to move progenitors from one competent state into another. In

Chapter 2 of this dissertation, I have discussed findings about the effects of blocking

13 Notch-signaling or its downstream effectors in vitro on cell-fate specification of

cultured embryonic retinal progenitors.

Notch-signaling and regeneration

Since Notch-signaling plays an important role in embryonic development, its

role has also been studied in another similar process – regeneration. Broadly defined,

regeneration is a process in which cells can de-differentiate, proliferate and acquire

pluripotent properties that allow them to replace lost or damaged tissues. Notch-

signaling has been implicated in the regeneration of several tissues such as skin

(Adolphe and Wainwright 2005), bone marrow (Caiado et al. 2008), pancreas (Su et

al. 2006), liver (Kohler et al. 2004), muscle (Luo et al. 2005) and inner ear (Ma et al.

2008; Stone and Rubel 1999). In these tissues, Notch-signaling is required during the

proliferative phase of regeneration and is thereafter downregulated. As suggested by

Hayes et al, the diversity of tissues that require active Notch-signaling suggests that

some mechanisms for regeneration are conserved (Hayes et al. 2007).

In fish, chicken and mammalian retinas, Müller glia have the capacity to regenerate the retina whereas in frogs and newts, retinal pigmented epithelial cells possess this capacity (Del Rio-Tsonis and Tsonis 2003; Hitchcock et al. 2004; Karl et

al. 2008). However, the ability of Müller glia to progress from an undifferentiated

progenitor-like cell to a fully differentiated neuron after retinal damage is restricted in

chicks and rodents. In Chapter 2 of this dissertation, I show that the Notch-signaling pathway is one of the contributing factors that modulates the regenerative potential of these cells.

14 Non-traditional roles of Notch in the nervous system

Apart from cell-fate decisions and glial instruction, Notch also plays a role in other aspects of the developing and mature nervous system. Notch-signaling may be crucial for neuronal migration, as is evidenced by the expression of Notch ligands in migrating neurons (Irvin et al. 2001; Solecki et al. 2001). There are some reports of

Notch expression in terminally differentiated glia (Hayes et al. 2007) in the retina and cortical neurons (Berezovska et al. 1998; Redmond et al. 2000; Sestan et al. 1999), suggesting that Notch may be required for maintaining a differentiated state of the cells. Notch has also been shown to affect neuronal morphology and reduce neurite extension (Berezovska et al. 1999; Qi et al. 1999; Redmond et al. 2000; Sestan et al.

1999). In mice, downregulation of the Notch1 pathway activity impaired long-term potentiation in the hippocampus (Wang et al. 2004) whereas Notch mutants exhibited deficits in spatial learning and memory (Costa et al. 2003), suggesting an important role for Notch in higher brain processes.

A recent study has shown that Notch is required for Schwann cell differentiation as well as postnatal myelination, thus highlighting the potential role of

Notch-signaling in neurodegenerative diseases (Woodhoo et al. 2009). In Chapter 3 of this dissertation, I discuss my own findings regarding the continued expression of

Notch in mature Müller glia as well as some previously undescribed roles of Notch in the postnatal retina, including its anti-neuroprotective role, its ability to regulate

FGF2-induced proliferation and its cross-talk with the MAPK-signaling pathway within the retina.

15 Notch-signaling – beyond the nervous system

Notch-signaling also has diverse functions even outside the nervous system. It

is involved in the development of a variety of tissues ranging from somite formation

where it exhibits oscillatory activity (Aulehla and Pourquie 2008; Shifley and Cole

2007) to angiogenesis and other aspects of vascular development (Gridley 2001), and

from T- and B-cell development (Stanley and Guidos 2009) to pancreatic

development (Murtaugh et al. 2003). Abnormal Notch-signaling has been implicated

in the development of cancers (Bolos et al. 2007), T-cell leukemias (Sharma et al.

2007) and diseases such as Multiple Sclerosis (Lasky and Wu 2005).

Patterning the circumferential marginal zone

Studies in the retina have revealed that this neural tissue is not uniformly

homogenous across all regions. A spatiotemporal gradient in maturity exists across its

length, with peripheral regions of the retina remaining more immature than central

regions of the retina (Prada et al. 1991; Prada et al. 1999). This gradient in maturity

presumably leads to a differential response of glial cells to stress and damage (Fischer

et al. 2002; Fischer et al. 2009). As described in Chapters 2 and 3, responses of

Müller glia to NMDA or growth factor treatment vary in central, temporal and nasal

regions of the retina. Müller glia are more responsive to damage (i.e. de- differentiative and proliferative) in peripheral regions than central regions of the retina. This ‘maturity gradient’ extends out to the outermost region of the retina to an area called the circumferential marginal zone (CMZ), which contains retinal stem cells. A CMZ containing retinal progenitors has been identified in most vertebrate

16 classes including frogs, fish and birds, with the exception of mammals. It is located at the transition between the multilayered retina and the non-pigmented epithelium

(NPE) of the ciliary body, a pseudostratified columnar monolayer of cells that lines the vitread surface of the ciliary body. In chickens, the CMZ adds relatively few retinal cells to the peripheral edge of the retina, but appears to persist into adulthood

(Fischer and Reh 2000; Kubota et al. 2002; Reh and Fischer 2001). Although the

CMZ of the chick retina has been well described in the postnatal eye, nothing is known about its formation and patterning in the embryonic eye. Thus, chapter 4 of this dissertation describes a study done to determine when the CMZ arises during embryonic retinal development. Using molecular markers for proliferation and stem cell markers, we characterized the formation and patterning of the CMZ in the embryonic chicken retina. We found that there is a gradual spatial restriction of progenitors into a discrete CMZ during late stages of embryonic development between E16 and hatching, at about E21. In addition, we found that retinal neurons remain immature for prolonged periods of time in far-peripheral regions of the retina.

Our studies reveal that the neurons that are generated by late-stage CMZ progenitors differentiate much more slowly than neurons generated during early stages of retinal development.

Serotonin accumulating bipolar cells in the retina – novel cells, novel role

Chapter 5 of this dissertation describes a novel cell-type, serendipitously discovered during an immunohistochemical characterization of bipolar cells in the chicken retina. Bipolar cells are interneurons, with cell bodies present in the same

17 layer of the retina (INL) as Müller glia. These cells transfer information from rod and

cone photoreceptors to ganglion cells. Cone bipolar cells are distinct from rod bipolar

cells, designated depending on the photoreceptor cell-type they synapse with. There

are approximately 10 different kinds of cone bipolar cells in the retina called ON or

OFF bipolar cells, depending on their responses to glutamate. ON bipolar cells

depolarize in the presence of light, while OFF bipolar cells hyperpolarize. This

information is passed on to the ganglion cells and is integral to generating the

diversity of ganglion cell responses seen in the retina. In our studies, we note that

some bipolar cells in the retina are weakly immunopositive for serotonin, even though

the enzymes for serotonin-synthesis are only present in photoreceptors and a subset of

amacrine cells (Chong et al. 1998; Cotton et al. 1988; Haan et al. 1987; Iuvone et al.

1999; Liang et al. 2004). We demonstrate that this distinct type of bipolar cell can

transiently accumulate serotonin and discuss some of the mechanisms, functions and purpose underlying serotonin accumulation in bipolar cells. A novel role for these

serotonin-accumulating bipolar neurons emerges, wherein these cells perform a

previously undescribed glial function in the retina by actively transporting and

degrading serotonin that is synthesized in neighboring amacrine cells.

CHAPTER 2

Notch-signaling regulates aspects of regeneration in the adult chicken retina

Introduction

Some of the earliest studies in retinal regeneration were done in lower vertebrates as early as the 1780s. A serendipitous discovery by French naturalist

Charles Bonnet revealed that the eyes of newts are capable of completely regenerating if the majority of this tissue is removed or damaged (Kelly 2006;

Morgan 1901). Research through the mid 1900s showed that retinal pigment epithelial cell grafts in salamanders and newts are capable of generating neural tissue in adult retinas (Stone 1950; Stone 1949; Stone and Steinitz 1957). With the advent of molecular biology and sophisticated biological techniques, a vast body of literature has accumulated in the field of retinal regeneration over the last 20 years, demonstrating that retinal regeneration can occur in a wide range of species including frogs (Del Rio-Tsonis and Tsonis 2003; Vergara and Del Rio-Tsonis 2009), fish

(Hitchcock et al. 1992; Otteson et al. 2002; Yurco and Cameron 2005), chicks

(Fischer and Reh 2001a) and mammals (Hitchcock et al. 2004; Karl et al. 2008; Ooto et al. 2004; Raymond and Hitchcock 1997).

Analysis of regeneration studies across species revealed 2 common principles:

(1) regeneration occurs more easily and effectively in invertebrates and lower vertebrates than in higher vertebrates and warm-blooded species (2) regeneration is 19 more effective and functionally restorative at embryonic stages of vertebrate species

rather than adult stages. The source of regenerative cells in the retina varies across

species and with age of the animals. In larval frogs, embryonic chick, mice and

urodeles, pigmented epithelial cells have been show to be a source of retinal

regeneration (Bumsted and Barnstable 2000; Fischer and Reh 2001b). In amphibians

and fish, the stem cells of the circumferential marginal zone (CMZ) respond to retinal

damage by increasing the production of new neurons (Lamba et al. 2008).

Additionally, a source of stem cells has been shown to exist in the inner nuclear layer

of adult teleost fish, which respond to retinal damage and regenerate photoreceptors

(Hitchcock and Raymond 1992; Raymond 1991).

In several vertebrate species, Müller glia, the primary support cells of the

retina, have the capacity to de-differentiate, proliferate and produce new neurons

(Fischer and Reh 2001a; Fischer and Reh 2003; Hitchcock et al. 2004; Lamba et al.

2009; Yurco and Cameron 2005). Although these cells can spontaneously de- differentiate into a retinal progenitor in fish, this stem-cell like potential of Müller glia is more restricted in birds and almost non-existent in mammals (Hitchcock et al.

2004; Karl et al. 2008). In chicks, Müller glia are the primary source of regenerating progenitor cells. After retinal damage by NMDA that kills amacrine and bipolar cells in the retina, Müller glia can de-differentiate, proliferate and a small percentage of these cells (~5%) go on to produce new neurons. However, most of these cells remain undifferentiated (~80%) while some produce new Müller glia (~20%) (Fischer and

Reh 2001a). Even though this de-differentiative potential in chicken Müller glia is small, the mechanisms that regulate the ability of these cells to acquire stem cell-like

20 properties remain unexplored. An understanding of the cues that promote this regenerative capacity of Müller glia and their ability to act as neuronal precursors holds not only the obvious significance of neuronal regeneration, but may also offer insights into controlling glial proliferation and reactive gliosis that may be detrimental to sight-threatening diseases of the retina.

As described in Chapter 1, the Notch-signaling pathway is one of the important regulators of differentiation and cell-fate specification and proliferation in the developing nervous system, including the retina (Livesey and Cepko 2001; Louvi and Artavanis-Tsakonas 2006). Notch is highly expressed in progenitor cells and inhibits neuronal differentiation by activating its downstream targets of the Hes/Hey family (including Hes1 and Hes5) which in turn inhibit the expression of proneural genes. Notch expression is high at embryonic stages of development and decreases over time in the retina. In addition to restricting neuronal differentiation, Notch is also instructive for Müller glial differentiation at later stages of histogenesis in the retina

(Satow et al. 2001; Vetter and Moore 2001).

Since the ability of Müller glia to progress from an undifferentiated progenitor-like cell to a fully differentiated neuron after retinal damage is restricted, we speculated that the Notch-signaling pathway may be one of the contributing factors in blocking the regenerative potential of these cells. Indeed, recent studies have revealed that Notch-pathway components are expressed in regenerating fish retinas (Yurco and Cameron 2007), in proliferating Müller glia (Raymond 1991;

Sullivan et al. 2003) and in regenerating newt retinas (Nakamura and Chiba 2007).

Thus we hypothesized that Notch expression and signaling is upregulated in Müller

21 glia, which antagonizes their ability to de-differentiate and produce new neurons and promotes glial differentiation in a damaged adult retina. Further, we hypothesized that blocking this Notch activity with a small-molecule inhibitor DAPT or by silencing cHes expression would promote neural regeneration and restrict glial differentiation.

To study the effects of Notch on retinal regeneration, we first characterized the expression of Notch and its associated signaling components in a damaged retina.

Next, we validated the use of Notch-pathway inhibitors, i.e, DAPT, short hairpin

RNA (shRNA) and double-stranded RNA (dsRNA) molecules to interfere with cHes1 and cHes5 in embryonic cell cultures to determine if these molecules can affect neuronal cell-fate. Lastly, we blocked Notch activity in NMDA-damaged retinas using DAPT and analyzed the outcomes on regeneration after injury. We compare and contrast our findings with a report by Hayes et al ( on the role of Notch-signaling during retinal regeneration in the chicken retina (Hayes et al. 2007).

Materials and Methods

Animals

The use of animals in these experiments was in accordance with the guidelines established by the National Institutes of Health and The Ohio State University. Eggs were obtained from the Department of Animal Sciences at the Ohio State University.

Chick embryos were staged according to guidelines established by Hamburger and

Hamilton (Hamburger 1951). Newly hatched leghorn chickens (Gallus gallus domesticus) were obtained from the Department of Animal Sciences at the Ohio State

22 University and kept on a cycle of 12-h light, 12-h dark (lights on at 7:00 a.m.). Chicks

were housed in a stainless steel brooder at about 28°C and received water and

PurinaTM chick starter ad libitum.

Tissue Culture

Retinas from embryonic day 6 (E6) chicken embryos were dissected in sterile

Hanks’ buffered saline solution (HBSS) added with 3% d -glucose and 0.01 M

HEPES buffer (HBSS+). Retinal cells were dissociated by mild trituration after incubation for 10 min at 37 °C in Ca2+ /Mg2+ -free HBSS plus 0.05% trypsin. Cell density was determined by using a hemacytometer. For RNA-interference studies, dissociated cells were centrifuged to form a dense cell pellet, which was resuspended in the nucleofection solution (See transfection procedure ahead). For all studies, between 100,000 and 200,000 cells were plated onto 12 mm glass coverslips that

were coated sequentially with poly-d-lysine and Matrigel (Collaborative Research)

diluted to 1:100 in HBSS. Cell cultures were maintained at 37 °C and 5% CO2 under

culture medium (DMEM:F12 without glutamate or aspartate; Invitrogen), added with

100 units/ml penicillin and 100 mg/ml streptomycin, 0.05 M HEPES and 2% fetal bovine serum; Gibco BRL). Cultures of chick-derived cells were maintained under

1 ml of DMEM-F12 per well (using 24-well plates) for up to 14 days with 50% of the medium replaced every 48 h. After one day in vitro, retinal cells were added with 0.5

µM DAPT in DMSO (N-[N-(3,5-Difluorophenacetyl-L-alanyl)]-S-phenylglycine t- butyl ester) (Sigma Aldrich), an inhibitor of γ-secretase activity associated with

Notch-signaling. Cultures were pulse- labeled with 1 µg/ml Bromodeoxyuridine 23 (BrdU) at 4 DIV. 5 hrs after BrdU treatment, cells were fixed and processed for immunocytochemistry.

Double-stranded RNA (dsRNA) synthesis

dsRNA for cHes1 and cHes5 was synthesized from the respective PCR products. Forward and reverse PCR primer sets were appended with T7 promoter regions. The sequences for these primers were as follows:

Gene Primers used Product size

cHes1 Fwd : 5’- taatacgactcactatagggagaggttcctagcgaggtccttt -3’

368bp Rev : 5’- taatacgactcactatagggagagtcaattcgccttcctcatc - 3’

cHes5 Fwd : 5’- taatacgactcactatagggagaccagagacaccaacccaact - 3’

277bp Rev : 5’- taatacgactcactatagggagagctggtgagtggaagtggat - 3’

Table 2.1: Primer sets used for Standard PCR followed by dsRNA synthesis. The table lists 43-45 bp forward and reverse primer sets for the appropriate genes appended with the T7 promoter region, along with the expected PCR product size for each primer set.

24 E7 RNA was extracted using standard methods of Trizol extraction. cDNA

template was synthesized from the total extracted RNA using the Superscript III First

Strand Synthesis System for RT-PCR (Invitrogen). Standard PCR reactions were

performed, in which cDNA was amplified for 35 cycles using standard protocols,

PlatinumTM Taq (Invitrogen) and an Eppendorf thermal cycler. The first 5 cycles were run at 60ºC (~5º higher than the Tm for the gene-specific region of the primer) and the

next 30 cycles were run at 69ºC (~5º higher than the Tm for the entire PCR primer).

The PCR products obtained were run on an agarose gel and stained with ethidium bromide to determine the predicted product sizes.

PCR products were extracted and cleaned using the QIAEX® II Gel Extraction

Kit (Qiagen Sciences, Maryland, USA). dsRNA was synthesized using the

Megascript® RNAi Kit (Ambion Inc. Texas, USA). Briefly, transcription reactions

were carried out in the purified linear PCR product using the T7 Enzyme Mix

provided and incubated overnight at 37ºC. The resulting RNA products were

annealed at 75ºC for 5 mins, and then cooled to room temperature. Nuclease digestion

was carried out using the 10X Digestion buffer provided to remove any remaining

DNA and ssRNA, followed by purification of the dsRNA using the 10X Binding

buffer and Filter cartridges provided. The dsRNA was eluted out in preheated Elution

solution, run on a gel, quantified and stored at -20ºC until their use.

25 shRNA plasmid synthesis

To maintain long-term expression of shRNA in cells, an alternative method was used

wherein double-stranded DNA oligos for cHes1 and cHes5 were cloned into the

BLOCK-iT™ U6 RNAi Entry Vector (Invitrogen, CA, USA) (Fig. 2.1).

Figure 2.1: Map of the BLOCK-iT RNAi Entry Vector The Gateway® Technology cloning method provides a rapid and efficient way to clone ds oligos encoding the desired shRNA target sequence into the pENTR™ entry vector containing an RNA Polymerase III expression cassette driven by a human U6 promoter. (http://tools.invitrogen.com/content/sfs/vectors/pentr_u6_rest.pdf)

26 The sequences for the double-stranded (ds) oligos were designed on the Invitrogen

BLOCK-iT RNAi Designer website (https://rnaidesigner.invitrogen.com/rnaiexpress/) and were ordered from Invitrogen. The sequences for the ds oligos were as follows:

Gene DNA oligos Product size Top strand :

5’- caccgcaaataccgggctggtttcacgaatgaaaccagcccggtatttgc -3’

cHes1 368bp Bottom strand :

5’- aaaagcaaataccgggctggtttcattcgtgaaaccagcccggtatttgc - 3’

Top strand :

5’- aaaaggaaatcctgacacccaaagattcgtctttgggtgtcaggatttcc - 3’

cHes5 277bp Bottom strand :

5’- caccggattactgcgaagggtatgccgaagcataccttcgcagtaatcc - 3’

Table 2.2: Double-stranded oligo sequences used for shRNA plasmid synthesis. The table lists 50 bp forward and reverse primer sets for the appropriate gene, along with the expected PCR product size for each primer set.

27 The top and bottom strands of these 50 bp oligos were annealed and ligated into the

vector using the manufacturer’s protocol. The resulting plasmid was used to

transform One Shot® TOP10 Competent E.coli cells and plated onto LB Agar plates containing 50 µg/ml kanamycin and incubated overnight at 37ºC. Plasmids extracted from the transformants were sequenced and stored at -20ºC until their use.

RNA interference in cell cultures

We interfered with Notch-signaling using RNA interference molecules to knockdown gene expression of cHes1 or cHes5. Retinal cell cultures from E6 chicken embryos were dissociated for cell cultures as described earlier. Co-transfection of the cHes1 (or cHes5) dsRNA along with pCAX-eGFP was carried out using the protocol modified from the Chicken Neuron Nucleofector® Kit (Amaxa Inc. USA). Briefly, 3

µg of pCAX-eGFP plasmid DNA and 3 µg of dsRNA were resuspended in 100 µl of

Amaxa Neuron Nucleofection Solution which contained the dissociated cell pellet at a final concentration of ~5x106 cells per 100 µl. This suspension was transferred to a

cuvette and inserted into the cuvette holder of the Amaxa Nucleofector. Program O-

03 or G-13 was used to transfect the cells. The cells were then resuspended in culture

medium and allowed to settle on cover slips in a 24-well plate. Similarly, co-

transfection of 3 µg of BLOCK-iT RNAi plasmids for cHes1 (or cHes5) dsRNA

along with pCAX-eGFP was carried out using the protocol described earlier. Cultures

were pulse-labeled with 1 µg/ml BrdU at 4 DIV. 5 hrs after BrdU treatment, cells

were fixed and processed for immunocytochemistry as described in Fig. 2.2.

Time (days)

0 1 23 4

Pulse-label Process cultures Dissociate E6 retinas, cultures for ICC transfect cells with for 5 hrs with dsHesRNA or BrdU shHesRNA vector

Figure 2.2: Paradigm used for RNA-interference studies in embryonic retinal cell cultures. Dissociated embryonic (E6) retinal cell cultures were transfected with double-stranded Hes1/5 RNA or short hairpin Hes1/5 RNA along with pCAX-EGFP. Transfected cell cultures were maintained for 4 days in culture. 5 hrs prior to fixing, the cultures were pulse-labeled with 0.1 µg BrdU per well.

In situ hybridization:

Partial length clones of cNotch1 (1043bp), cDelta1 (784bp) and cHes1 (540bp) were

kindly provided by Dr. Domingos Henrique (Inst. de Medicina Molecular, Lisboa,

Portugal). Riboprobes were synthesized using a kit provided by Roche and stored at -

80°C until use. Ocular tissues from P7 eyes were dissected in RNase-free Hanks

Balanced Salt Solution (HBSS; Invitrogen), fixed overnight at 4°C in 4%

paraformaldehyde buffered in 0.1 M dibasic sodium phosphate, and embedded in

OCT-compound (Tissue-Tek). Cryosections were processed for in situ hybridization

with digoxigenin-labeled RNA probes as described previously (Fischer et al. 2002a;

Fischer et al. 2004). Hybridization was detected by using Fab fragments to

digoxigenin that were conjugated to alkaline phosphatase (anti-DIG-AP; Roche)

diluted in MABT (0.05 M maleic acid buffer, 0.1% Tween-20) plus 10% normal goat

29 serum, 10 mM levamisole, and 10 mM glycine. Nitro-blue tetrazolium (NBT) and 5-

Bromo-4-Chloro-3 Indolyphosphate p-Toluidine (BCIP) in 0.1 M NaCl, 0.1 M tris-

HCl pH 9.5, 0.05 M MgCl2 and 0.01% Tween-20 were used to precipitate

chromophore from the anti-DIG-AP. With sufficient levels of chromophore

developed, the sections were washed in PBS, fixed in 4% paraformaldehyde in PBS

for 15 mins, washed again in PBS and processed for immunolabeling for 2M6, BrdU,

and vimentin as described ahead.

Fixation, Sectioning and Immunohistochemistry

Cultured cells on coverslips were fixed in 4% PFA in PBS for 20 mins,

washed 3 times in PBS and immunolabeled as described ahead. Coverslips were

covered with 200 µl primary antibody solution (diluted in PBS plus 5% normal goat

serum, and 0.2% Triton X-100) and incubated for about 24 hrs at 20°C. The

coverslips were washed 3 times in PBS, covered with 200 µl secondary antibody

solution and incubated for about 1 hour at 20°C. For in vivo studies, tissue dissection,

fixation, cryosectioning, and immunolabeling were performed as described previously

(Fischer et al. 2002b; Fischer et al. 2009). Working dilutions and sources of antibodies used in this study included; rat anti-BrdU at 1:200 (OBT0030S; Serotec); mouse anti-BrdU at 1:100 (G3B4; DSHB); mouse anti-vimentin at 1:100 (H5;

Developmental Studies Hybridoma Bank); mouse anti-2M6 at 1:100 (Dr. P. Linser,

University of Florida); mouse anti-visinin at 1:50 (Developmental Studies Hybridoma

Bank); mouse anti-HuD at 1:200 (Monoclonal Antibody Facility, University of

Oregon); rabbit anti-calretinin at 1:500 (Swant, Switzerland); mouse anti-islet at 30 1:100 (67.4E12; DSHB); mouse anti-Lim3 at 1:50 (40.2D6, DSHB); and rabbit anti- eGFP at 1:3000 (Dr. L. Berthiaume, University of Alberta). Secondary antibodies included donkey-anti-goat-Alexa488, goat-anti-rabbit-Alexa488, goat-anti-mouse-

Alexa488/568/647, goat-anti-rat-Alexa488, rabbit anti-goat Alexa488 and goat-anti- mouse-IgM-Alexa568 (Invitrogen) diluted to 1:1,000 in PBS plus 0.2% Triton X-100.

Cells were labeled with DAPI (Sigma) at 1 µg/ml in PBS prior to mounting the cells and coverslips in Fluoromount-G (Southern Biotech, Alabama).When double labeling with BrdU, the complementary label was fixed after the secondary antibody and then acid-treated before addition of the primary antibody to BrdU as described previously

(Ghai et al. 2008).

Photography, measurements, cell counts and statistical analyses

Photomicrographs were taken by using a Leica DM5000B microscope equipped with epifluorescence, FITC and rhodamine filter combinations and 12 megapixel Leica DC500 digital camera. Images were optimized for color, brightness and contrast and double-labeled images overlaid by using Adobe Photoshop™ 6.0.

For cell culture studies, cell counts were made on at least five fields of view from at least three coverslips per experimental treatment and means and standard errors calculated. Experiments were repeated three times. Data between experimental conditions were compared statistically with Student’s t-test. Images were optimized for color, brightness and contrast, and double-labeled images overlaid by using

Adobe PhotoshopTM6.0. For in vivo studies, cell counts were made from at least six

31 different animals, and means and standard deviations calculated on data sets. To

avoid the possibility of region-specific differences within the retina, cell counts were

consistently made from the same region of retina for each data set.

Real-time PCR

cDNA was synthesized as described using standard protocols as described

before (Fischer et al. 2004). mRNA was harvested from retinas, reverse-transcribed to

generate cDNA, and amplified using real-time PCR for 40 cycles. Real-time PCR was

performed using the StepOnePlus™ Real-Time PCR System (Applied Biosystems,

Foster City, CA, USA) according to the manufacturer's instructions. Reactions were

performed in triplicates, in 25 µl volumes with 0.5 µM primers and MgCl2 concentration optimized between 2-5 mM. Nucleotides, Taq DNA polymerase, and buffer were included in the SYBR® Green PCR Master Mix (Applied Biosystems,

Foster City, CA, USA). A typical protocol designed on StepOne Software v2.0

(Applied Biosystems, Foster City, CA, USA) included a holding stage for 10 min at

95°C, 15 s denaturation step followed by 40 cycles with a 95°C denaturation for 15 s and 60°C annealing for 1 min. The Melt Curve stage included 95°C for 15 s, 60 °C annealing for 1 min and 95 °C for 15 s. Measurements of the fluorescence were carried out at the end of the 60°C annealing period. Ct values obtained from real-time

PCR were normalized to GAPDH and the fold difference between control and treated samples was determined using the ∆Ct method and represented as a percentage change from baseline.

32 Results

Notch-pathway gene expression increases after damage

To determine whether components of the Notch-signaling pathway are upregulated in the postnatal chick retina after damage, we used in situ hybridization and real-time

PCR. We injected 2 µmol NMDA intraocularly in P7 chicks and harvested the retinas

2-7 days after damage. We found that cNotch1 expression was low in control retinas

(Fig. 2.3a), increased in cells of the inner nuclear layer (INL) within 2 days of

damage (Fig. 2.3b), peaked at 3 days after damage (Fig. 2.3c) and returned to baseline

levels at 7 days after damage (Fig. 2.3d). At 3 days post damage, cells that expressed

cNotch1 appeared to migrate proximally and distally across the INL and also to the

outer nuclear layer (ONL), a trend reminiscent of Müller glial migration after damage

(Fischer and Reh 2001a; Thummel et al. 2008). cDelta1 expression was low in

control retinas (Fig. 2.3e), increased 3 days post damage (Fig. 2.3f) and persisted at 7

days post damage (Fig. 2.3g).

Similarly, cHes1 expression was low in control retinas (Fig. 2.3h) and increased 3

days after damage in the INL (Fig. 2.2i). Taken together, these findings indicate that

Notch-pathway gene expression increases after damage in a postnatal chick retina.

Figure 2.3: Notch-pathway gene expression increases after damage. An intraocular injection of saline (left eye) or 2 µmol NMDA (right eye) was made at P7, and eyes were harvested 2-7 days after treatment as shown earlier. Vertical sections of the retina were hybridized with riboprobe to cNotch1 (a-d), cDelta1 (e-g) or cHes1 (h, i). Arrows indicate cNotch1, cDelta1 or cHes1 positive cells in the retina. The calibration bar (50 µm) in panel a applies to panels a-i. Abbreviations: P7 – postnatal day 7, ISH – In situ hybridization, ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer, GCL – ganglion cell layer

34 Proliferating Müller glia express cNotch1 after damage

To determine whether proliferating cells express cNotch1 after damage, we injected NMDA intraocularly followed by BrdU and harvested the eyes 24 hrs later.

We used a combination of in situ hybridization and immunohistochemistry to analyze the results. We found that numerous cNotch1+ cells were also positive for BrdU, indicating that proliferating cells expressed cNotch1 (yellow arrows; Fig. 2.4a-c).

Additionally we also found some cNotch1+ cells that did not label for BrdU (red arrows; Fig. 2.4a-c) and some BrdU+ cells that did not label for cNotch1 (white arrows; Fig. 2.4a-c).

Figure 2.4: Proliferating cells express cNotch1 after damage. Eyes were treated with 2 µmol NMDA in both eyes at P7. 1 µmol BrdU was injected 2 days later at P9 and harvested 24 hrs at P10. Vertical sections of the retina were hybridized with riboprobe to cNotch1 (a) and subsequently labeled with antibodies to BrdU (b). Panel c is a 3 fold merged magnification of panels a-c. Yellow arrows indicate cNotch1+BrdU+ cells; red arrows indicate cNotch1+BrdU- cells; white arrows indicate cNotch-BrdU+ cells in the retina. The calibration bar (50 µm) in panel b applies to panels a-b. The calibration bar (50 µm) in panel c applies to panel c alone. Abbreviations: P7 – postnatal day 7, ISH – In situ hybridization, ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer

36 Since glial cells are known to proliferate in retinas after damage, we sought to determine whether the cNotch1+BrdU+ cells in the retina are Müller glia. To do this, we injected NMDA intraocularly at P7 followed by BrdU at P9 and harvested the eyes 4 hrs later. We found that cells were positive for vimentin, a Müller glial marker, as well as cNotch1 and BrdU (yellow arrows; Fig. 2.5a-d). Additionally we also found vimentin+ cells that were cNotch1+ but BrdU- (red arrows; Fig. 2.5a-d) and vimentin+ cells that were cNotch1- but BrdU+ (white arrows; Fig. 2.5a-d).

Collectively, these findings indicate that proliferating Müller glia upregulate Notch expression after damage in a postnatal chick retina.

Figure 2.5: Proliferating Müller glia express cNotch1 after damage. Eyes were treated with 2 µmol NMDA in both eyes at P7. 1 µmol BrdU was injected 4 hrs before harvesting the eyes at P9. Vertical sections of the retina were hybridized with riboprobe to cNotch1 (a) and subsequently labeled with antibodies to vimentin (b) and BrdU (c). Panel d is a 4 fold merged magnification of panels a-c. Yellow arrows indicate cNotch1+BrdU+vimentin+ cells; red arrows indicate cNotch1+BrdU- vimentin+ cells; white arrows indicate cNotch-BrdU+vimentin+ cells in the retina. The calibration bar (50 µm) in panel b applies to panels a-b. The calibration bar (50 µm) in panel d applies to panel d alone. Abbreviations: P7 – postnatal day 7, ISH – In situ hybridization, ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer

38 The Notch-pathway inhibitor DAPT alter cell-fate in retinal cell cultures

To determine whether Notch-pathway inhibitors can alter cell-fate

specification, we first used the inhibitors in studies on embryonic retinal cell cultures,

based on the assumption that the process of regeneration employs similar mechanisms

as those seen during embryonic development. For this, we dissociated embryonic day

6 (E6) retinas and introduced 0.5µM DAPT, in DMSO into the cultures one day after

allowing the cell to settle. 4-day-old cell cultures were pulse-labeled with BrdU for 5 hrs, fixed and subsequently immunolabeled for markers of proliferation (BrdU), amacrine cells and horizontal cells (calretinin), bipolar cells (Islet1) and horizontal cells (Lim3) and photoreceptors (visinin). DAPT is a small-molecule inhibitor of γ- secretase activity that is required for active Notch-signaling. In separate cell cultures using the same experimental paradigm, cells were labeled for Islet1 that labels bipolar cells, ganglion cells and some amacrine cells (Edqvist et al. 2006; Galli-Resta et al.

1997) and Lim3 that labels bipolar cells (Edqvist et al. 2006). Cells were also counterstained for DAPI, a nuclear label (Fig. 2.6a-i). We found a 1-fold increase in the percentage of cells differentiating as (calretinin+) amacrine cells (Fig. 2.6j) in the presence of DAPT. We also saw a 0.75-fold increase in the percentage of cells differentiating as bipolar and ganglion cells (Islet1+) (Fig. 2.6k), but no change in

(visinin+) photoreceptors (Fig. 2.6j) or (Lim3+) bipolar or horizontal cells (Fig. 2.6k).

We did not observe any change in the percentage of (BrdU+) proliferating progenitors in the presence of DAPT (Fig. 2.6j). It is possible that Islet1 and Lim3 label different subsets of bipolar cells; Islet1+ bipolar cells prematurely differentiate in the presence of DAPT, whereas Lim3+ bipolar cells do not.

Figure 2.6: DAPT increases percentage of cells differentiating as (calretinin+) amacrine and horizontal cells and (Islet1+) amacrine and bipolar cells, but not (visinin+) photoreceptors or (Lim3+) bipolar and horizontal cells, without depleting (BrdU+) proliferating progenitors. Cell cultures were fixed and observed under bright field (BF) (a, e) and labeled with DAPI (b, f) and antibodies to BrdU (green; c, g), visinin (green; g) and calretinin (red; h). In other experiments using the same paradigm, cell cultures were labeled with Islet1 and Lim3 (labeling not shown). Panel d is a merged representation of panels a-c; panel i is a merged representation of panels e-h. (j and k) Histograms indicate the percentage of DAPI positive cells that are also positive for the antibody used to label the appropriate cell-type. The calibration bar (200 µm) in panel a applies to panels a-i 40 Effects of RNA interference-mediated silencing of Hes genes has varied effects on

cell-fate in embryonic retinal cell cultures

As DAPT is a generic γ-secretase inhibitor that may have pleiotropic effects, we sought to specifically inhibit downstream effectors of the Notch-pathway – the bHLH factors Hes1 and Hes5. To do this, we used short interfering RNA molecules or double- stranded RNA molecules to degrade the endogenous mRNA molecules for these bHLH factors. We dissociated E6 retinal cultures and transfected them with

RNAi molecules along with pCAX-EGFP, to identify transfected cells. 4-day-old cell cultures were pulse-labeled with BrdU for 5 hrs, fixed and subsequently immunolabeled for GFP to label transfected cells (Fig. 2.7b,d, f, h, j, l) proliferation

(BrdU) (Fig. 2.7k, l), amacrine cells and ganglion cells (HuD) (Fig. 2.7g, h), photoreceptors (visinin) (Fig. 2.7c, d) along with DAPI (Fig. 2.7a, e, i, d, h, l). In the presence of dsRNA-interference molecules to cHes1, we found a 0.66-fold increase in the percentage of cells differentiating as visinin+ photoreceptors, no change in HuD+ amacrine and ganglion cells and an overall decrease in the number of proliferating cells (Fig. 2.8a). Consistent with these results, in the presence of shRNA molecules to cHes1, we found a 0.5-fold increase in the number of cells differentiating as visinin+ photoreceptors and no change in HuD+ amacrine and ganglion cells (Fig. 2.8b).

Figure 2.7: Transfected embryonic retinal cell cultures. Embryonic retinal cells transfected with pCAX-eGFP or pCAX-eGFP+RNAi molecules were labeled with DAPI and antibodies to GFP/visinin, GFP/HuD and GFP/BrdU. Panel d is a merged representation of panels a-c; Panel h is a merged representation of panels e-g; panel l is a merged representation of panels i-k. Arrows indicate (marker+GFP) double- labeled DAPI positive cells. The calibration bar (200 µm) in panel a applies to panels a-l.

Surprisingly, we saw the opposite effects with RNAi molecules to cHes5. In the presence of dsRNA-interference molecules to cHes5, we found a 3-fold decrease in the percentage of cells differentiating as visinin+ photoreceptors, a 1.4-fold decrease in the percentage of cells differentiating as HuD+ amacrine and ganglion cells and no change in the number of proliferating cells (Fig. 2.8c). However, in the

42 presence of shRNA molecules to cHes5, we did not observe any changes in the percentage of cells differentiating as visinin+ photoreceptors or HuD+ amacrine and ganglion cells (Fig. 2.8d). Taken together, our results suggest that RNA-interference of cHes1 blocks differentiation of photoreceptors but not of amacrine or ganglion cells. Further, RNA-interference of cHes5 has the opposite effect wherein it suppresses neurogenesis of photoreceptors and amacrine and ganglion cells.

Figure 2.8: Effects of RNA interference-mediated silencing of Hes genes has varied effects on cell-fate. (a-d) Histograms indicate the percentage of DAPI+GFP+ cells that are also positive for the antibody used to label the appropriate cell-type.

43 DAPT blocks Notch-pathway gene expression after damage

To test whether Notch-pathway inhibitors enhance neurogenesis as we had seen in embryonic cell cultures, we injected the small-molecule inhibitor of γ- secretase activity, DAPT, into NMDA-damaged eyes. First, we used in situ hybridization and real-time PCR to determine whether DAPT influences Notch expression and activity. Eyes were treated with 2 µmol NMDA in both eyes followed by Saline/DMSO (left eye) or DAPT (right eye) 2 days after damage and harvested 24 hrs later. We found that DAPT blocked the expression of cNotch1 in NMDA- damaged retinas (Fig. 2.9a-b). In NMDA-damaged retinas, we observed a 1.8-fold increase in cNotch1 expression, 1.5-fold increase in cDelta1 expression, 4.2-fold increase in cHes1 expression and a 0.5-fold increase in cHes5 expression (Fig. 2.9c).

When DAPT was administered after damage, it effectively blocked cNotch1, cDelta1, and cHes1 and cHes5 expression (Fig. 2.9d). We observed a 2-fold decrease in cNotch1 expression, 1.5-fold decrease in cDelta1 expression, 3.75-fold decrease in cHes1 expression and a 2.5-fold decrease in cHes5 expression (Fig. 2.9d). These findings suggest that DAPT decreases Notch-pathway gene expression as well as function.

Figure 2.9: DAPT blocks Notch-pathway gene expression after damage. Eyes were treated with 2 µmol NMDA in both eyes followed by Saline/DMSO (left eye) or DAPT (right eye) 2 days after damage and harvested 24 hrs later. Vertical sections of the retina were hybridized with riboprobe to cNotch1 (a, b). Eyes were also harvested for QPCR. Notch-pathway gene expression (cNotch1, cDelta1, cHes1) increases 3 days after NMDA treatment (c). DAPT can reduce the Notch-pathway gene expression (cNotch1, cDelta1, cHes1, cHes5) when administered 2 days after NMDA treatment and harvested 24 hrs later (d). Arrows indicate cNotch1 positive cells in the retina. The calibration bar (50 µm) in panel b applies to panels a-b. Abbreviations: P7 – postnatal day 7, ISH – In situ hybridization, QPCR – quantitative PCR, ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer, GCL – ganglion cell layer.

45 DAPT increases the number of BrdU-labeled cells that remain in the central retina after damage

Next, we sought to determine whether blocking Notch-signaling in a damaged retina influences Müller glial de-differentiation and proliferation. We treated eyes with NMDA followed by DAPT and BrdU 2 days later to label proliferating cells and harvested the eyes 10-11 days later. We immunolabeled retinas for a Müller glial marker - 2M6 (Fig. 2.10a), a neuronal marker - HuD (Fig. 2.10b) and BrdU (Fig.

2.10c). We observed that proliferating cells label for glial and/or as neuronal markers

(Fig. 2.10a-d). We observed that the numbers of proliferating cells in central regions of the retina increased almost 2-fold when Notch-signaling was inhibited with DAPT after damage (Fig. 2.10e). However, no change was observed in the numbers of proliferating cells in nasal or temporal regions of the retina.

Figure 2.10: DAPT increases the number of proliferating cells in the central retina after damage. Eyes were treated with 2 µmol NMDA in both eyes at P7 followed by 1 µmol BrdU (control) or 1 µmol BrdU and 100µM DAPT (treated) at P9. Eyes were harvested at P19 or P20. Vertical sections of the retina were labeled with antibodies to 2M6 (a), HuD (b) and BrdU (c). Panel d is a 3 fold merged magnification of panels a-c. Yellow arrows indicate HuD+BrdU+ cells; white arrows indicate 2M6+BrdU+ cells in the retina. The calibration bar (50 µm) in panel a applies to panels a-c. The calibration bar (50 µm) in panel d applies to panel d alone. Abbreviations: P7 – postnatal day 7, ISH – In situ hybridization, ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer, GCL – ganglion cell layer, IHC – immunohistochemistry

DAPT enhances neuronal differentiation in the central regions of the retina, and

glial differentiation in temporal regions

Fischer and Reh have previously shown that of all proliferating Müller glia in

a damaged retina, 20% can differentiate again into Müller glia and about 5%

differentiate into neurons such as amacrine and bipolar cells. However, the majority

of proliferating cells (about 75%) remain undifferentiated. To determine whether

blocking Notch- signaling in damaged retinas can alter cell-fates of the proliferating

47 Müller glia, we treated eyes with NMDA followed by DAPT and BrdU 2 days later to

label proliferating cells and harvested the eyes 10-11 days later and immunolabeled them for glial, neuronal markers (Fig. 2.10a-d). We saw that the cell-fates of proliferating Müller glia varied depending on the region of the retina observed. In central regions of the retina, there was no change in the percentage of proliferating cells differentiating as 2M6+ Müller glia, a 2-fold increase in the percentage of proliferating cells differentiating as HuD+ amacrine or bipolar cells, and no change in the percentage of other proliferating glia (undifferentiated cells or other unidentified cell-types) in the presence of DAPT (Fig. 2.11a). In nasal regions of the retina, there

was no change in the percentage of proliferating cells differentiating as Müller glia, amacrine or bipolar cells or those glia that remained undifferentiated and/or unidentified in the presence of DAPT (Fig. 2.11b). In temporal regions of the retina, we observed a 10% increase in the percentage of proliferating cells differentiating as

Müller glia and no changes in the number of proliferating cells differentiating as amacrine cells, bipolar cells or undifferentiated / unidentified glia in the presence of

DAPT (Fig. 2.11c). Collectively, our results suggest that DAPT can alter cell-fates of proliferating Müller glia after damage in a region-specific context.

Figure 2.11: DAPT enhances neuronal differentiation in the central regions of the retina, and glial differentiation in temporal regions. Eyes were treated with 2 µmol NMDA in both eyes at P7 followed by 1 µmol BrdU (control) or 1 µmol BrdU and 100µM DAPT (treated) at P9. Eyes were harvested at P19 or P20. Histograms represent the number of 2M6, HuD or other BrdU+ cells per 2mm2 in central regions (a), nasal regions (b) or temporal regions (c) of the retina. Abbreviations: P7 – postnatal day 7, IHC – immunohistochemistry.

49 Discussion

The Notch-pathway is a key component in sculpting the nervous system of

species ranging from flies to humans. In the retina, this pathway regulates cell-fates

and proliferation during histogenesis (Livesey and Cepko 2001) and is instructive for

glial differentiation (Vetter and Moore 2001). Further, Notch expression is high during embryonic development and decreases once histogenesis is completed. Apart from embryonic development, studies have also implicated the involvement of the

Notch-pathway during regeneration of various tissues such as pancreas (Su et al.

2006), bone marrow (Han et al. 2000; Han et al. 2006), skin (Adolphe and

Wainwright 2005), muscle (Carey et al. 2007; Luo et al. 2005) and inner ear (Ma et al. 2008; Stone and Rubel 1999) Notch expression has been shown to be enhanced in regenerating retinas of fish (Wu et al. 2001; Yurco and Cameron 2007) and newts

(Nakamura and Chiba 2007). Elsewhere in the nervous system, Notch is expressed in certain regions of the adult mammalian brain that retain neurogenic and regenerative capability, such as the subventricular zone, dentate gyrus, and the rostral migratory stream (Berezovska et al. 1999; Costa et al. 2003; Stump et al. 2002; Wang et al.

2004), Notch is also known to persist in the peripheral region of the retina containing

the circumferential marginal zone (CMZ) (Fischer 2005), a region of the retina that is

known to maintain an immature phenotype as discussed in Chapter 4. A common

theme for the involvement of Notch-signaling emerges, wherein Notch appears to be

a critical component in ‘immature’ cells and tissues that retain or acquire the capacity to be pluripotent or stem cell-like.

50 In this study, we report that Notch-pathway gene expression increases in

Müller glia in the postnatal chicken retina after retinal damage. De-differentiating and

proliferating Müller glia in a damaged retina can go on to produce new neurons

(Fischer and Reh 2001a; Hayes et al. 2007). Since neurogenesis in the regenerating

retina may involve similar Notch-driven mechanisms as seen during early

development, we first validated the use of Notch inhibitors by testing their effects on

neurogenesis in embryonic retinal cell cultures. We used retinal cultures of E6 chick

embryos for our studies, a stage at which the main cell types that are being born are

amacrine cells and rod photoreceptors. We found that inhibition of γ-secretase activity associated with Notch using a small-molecule inhibitor DAPT enhances premature differentiation of amacrine and ganglion cells whereas RNA-interference molecules to cHes1 or cHes5 enhance or suppress respectively, premature

photoreceptor neurogenesis in embryonic retinal cultures. Applying the Notch- inhibitor DAPT in vivo in a damaged postnatal retina, we show that inhibition of

Notch-signaling alters cell-fates even in a regenerating retina, i.e., it enhances

neuronal differentiation in central regions and glial differentiation in temporal regions

of the regenerating retina.

Our findings indicate that inhibition of γ-secretase activity with DAPT and

silencing of bHLH factors Hes1 and Hes5 have distinctly different outcomes on cell- fate specification of cultured chicken retinal progenitors. Inhibition of γ-secretase

activity in dissociated E6 retinas leads to premature differentiation of amacrine cells

whereas inhibition of Hes1 leads to premature differentiation of photoreceptors. Thus,

our data suggests that the overall suppression of γ-secretase activity associated with

51 the Notch-pathway impact neurogenesis differently from inhibition of specific

downstream effectors and that RNAi-mediated effects on neurogenesis are more

specific than DAPT. Our findings are similar to reports showing that knockdown of

Notch-signaling at early stages of embryonic retinal development enhances the

number of ganglion cells being born (Austin et al. 1995) and that Notch1 inhibits

photoreceptor production in developing mammalian retinas (Jadhav et al. 2006; Kubo

et al. 2005; Yaron et al. 2006).

Curiously, we see that Hes1 normally inhibits the photoreceptor fate of embryonic chicken retinal cells whereas Hes5 may act in opposition to Hes1 with respect to photoreceptor differentiation. Additionally, Hes1 and Hes5 are differentially regulated during Notch activation after damage. After damage, Hes1 expression increases to a much greater extent than Hes5. In the presence of DAPT,

Hes1 is also suppressed more potently than Hes5. These antagonistic effects could be because Hes1 and Hes5 belong to two different classes of bHLH factors. There are several subfamilies of bHLH proteins as shown in Fig. 2.12 (Davis and Turner 2001).

Hes1 belongs to hairy subgroup while Hes5 belongs to E(spl) which lacks HC after the Orange domain. The role played by HC is unclear; however, evidence from the literature suggests that Hes1 and Hes5 may regulate different aspects of development in the nervous system and need not be interchangeable (Jacobsen et al. 2008;

Kageyama et al. 2008; Wu et al. 2003; Zine et al. 2001).

Figure 2.12: Basic structure of bHLH-O subfamily domain structures. The bHLH and Orange domains are present in all family members. The HC domain is specific to the hairy subfamily. Hairy-like proteins end with WRPW and in some cases, two additional amino acids. E(spl)-like proteins usually end with WRPW. All Hey-like proteins have a YXXW motif near the C-terminus, with a generally conserved 6 ± 10 amino acid extension (modified from Davis and Turner 2001).

Another reason for differential effects of Hes1 and Hes5 could be that these

two bHLH factors are influenced by pathways other than Notch. The initiation of

Hes1 expression in neuroepithelial cells occurs prior to the expression of Notch-

signaling components (Kageyama et al. 2008). Hes5 on the other hand is expressed

later, only after the expression of Notch-signaling components is established in the neuroepithelium (Kageyama et al. 2008). Further, it was recently shown that Hes1 is

downstream of the Shh signaling pathway in the retina (Wall et al. 2009). Elsewhere in the nervous system, Hes1 has been shown to bind to STAT3 to mediate crosstalk between the Notch and JAK-STAT pathways and maintain radial glial cells as well as promote astrocyte differentiation (Kamakura et al. 2004). Further, Hes1 but not Hes5, was found to promote astrocyte differentiation at the expense of oligodendrocyte differentiation (Wu et al. 2003). Studies done in Hes1 and Hes5 mice mutants reveal

53 that Hes1-null retinae have more severe developmental defects than Hes5-null retinae

(Davis and Turner 2001; Hojo et al. 2000; Tomita et al. 1996). Hes5 mutant mice

have a severe reduction in Müller glial cell production (Hojo et al. 2000), while Hes1

mutant mice have increased numbers of all interneurons and ganglion cells in

abnormal rosettes, as well as a decrease in Müller glial cells (Takatsuka et al. 2004;

Tomita et al. 1996).

Even though Hes1 and Hes5 are both required to promote gliogenesis in the

retina at the expense of neuronal fate (Furukawa et al. 2000; Hojo et al. 2000;

Kageyama et al. 1997), they may play redundant roles when expressed within the

same cells. Ohstuka et al have reported that activated Notch completely inhibits the

differentiation of neural precursors from wild-type mice or mice with targeted

disruptions with either Hes1 or Hes5 (Kageyama and Ohtsuka 1999). However, in

neuronal precursors with disruptions in both Hes1 and Hes5, activated Notch did not

efficiently inhibit neuronal differentiation (Davis and Turner 2001). Thus, these

studies indicate that even though Hes1 and Hes5 are both downstream of Notch-

signaling and have some redundant functions, they may have distinctly different

outcomes on cell-fate specification within the retina.

Some caveats exist in our studies on Notch-inhibition in embryonic cell

cultures. Appropriate scrambled controls for Hes1 and Hes5 ds- and shRNA have not

been used in our experiments. Further, transfection efficacy is low (~30% of cells are

GFP-positive in transfected cell cultures), hence the siRNA effect is partial.

Moreover, we have not gauged the extent of knockdown of the Hes factors in embryonic cell cultures. Ideally, testing this would require in ovo knockdown of Hes

54 factors, followed by in situ hybridization or QPCR in transfected embryonic eyes.

Another caveat to testing the effects of Notch-signaling in cell cultures is that dissociation of cells may by itself reduce Notch-Delta interactions and hence reduce the potency of Notch-signaling. Despite all these drawbacks, we still see significant effects of Notch on neurogenesis in cell cultures.

Even though the RNAi molecules to effectors of Notch-signaling worked well in retinal cell cultures, we have been unsuccessful in our numerous attempts to transfect postnatal chicken retinas in vivo with these molecules using electroporation, mainly due to the insulating barrier provided by the sclera as well as the jelly-like consistency of the vitreous humor that retards the movement of RNAi molecules when injected into the eye. Thus, the minimally invasive molecular tools that can be used to study regeneration in the adult chicken retina are currently restricted.

However, DAPT is a potent pharmacological inhibitor that blocks Notch-signaling.

Our findings indicate that there is a feedback mechanism by which Notch expression and function are closely linked, and that perturbation of Notch function downregulates its expression as well.

Our data sheds light on the cell-types that express Notch after damage in the retina. As described in Chapter 3, Notch expression is normally present at low levels in Müller glia in the absence of damage. After damage, the expression of Notch receptor, its ligand cDelta1 and the effectors Hes1 and Hes5 are increased in the retina, as determined by real-time PCR. In situ hybridization reveals that Notch- pathway gene expression is enhanced 2-7 days after damage. cNotch1 is expressed mainly by proliferating Müller glia, within 2 days after NMDA-damage. cNotch1

55 expression remains high and peaks 3 days after damage as Müller glial cells migrate

proximally and distally across breadth of the INL and into the OPL. Notch expression

returns to low levels within 7 days after damage. Although we have not definitively

identified the cell types that express cDelta1 and cHes1 (and cHes5) within the retina

after damage, expression of these genes is also seen in the INL within 3 days after

damage, in the layer where interneurons and Müller glial cell nuclei are present.

cDelta1 expression is also seen within the IPL and scattered in the GCL after damage.

As discussed in Chapter3, we speculate that the neurons adjacent to Müller glia,

ganglion cells, microglia or recently discovered novel inner retinal glia-like cells

(NIRGs), may upregulate cDelta1 in the retina.

Previous studies of retinal regeneration have revealed that the majority of the

Müller glial-derived progenitors do not differentiate into neurons, but remain as

undifferentiated progenitors (Fischer and Reh 2001a). We tested whether enhanced

expression and activation of Notch prevents these progenitors from differentiating into neurons or glia during regeneration. Our findings are consistent with the hypothesis that Notch is required to specify cell-fate of stem cell-like Müller glia in a damaged retina, but is not the only factor influencing the regenerative process.

Inhibiting the pathway with DAPT causes a small increase in the percentage of these glial-derived progenitors to differentiate into neurons in central regions and a small percentage to differentiate as glia in temporal regions of the retina. One surprising finding is that inhibition of Notch promotes glial differentiation to a small extent in our studies. Notch-signaling is known to be instructive for glial differentiation during development, so blocking Notch would be expected to suppress glial differentiation.

56 This suggests that the damaged environment in the postnatal retina can not be

considered equivalent to an embryonic retina and that the effects of Notch-signaling

are context-dependent.

We report an enhancement in proliferation of glial-derived progenitors when

Notch-signaling is blocked after damage. An increased number of BrdU cells could

mean that blocking Notch-signaling enhances proliferation of glial-derived

progenitors. Alternatively, it could imply that there is improved survival of BrdU-

labeled glial-derived cells. Our experiments do not distinguish between these two

possibilities. Our study is similar to one that was recently reported by Hayes et al

(Hayes et al. 2007) However, these authors report a decrease in proliferation with

DAPT after NMDA-damage to the retina. Differences in the paradigms used may

contribute to the opposing effects seen in the two studies. In our study, we blocked

Notch activity at 2 days after damage whereas Hayes et al applied DAPT 1 day after

damage. In addition to more proliferation, we also see more neurogenesis in central

regions of the retina and more gliogenesis in temporal regions of the retina with the

same paradigm used, whereas Hayes et al report more neurogenesis only when Notch

is blocked 3 days after damage (Hayes et al. 2007). Collectively, these findings highlight the fact that Müller glia change their phenotype rapidly after retinal damage,

proceeding from a de-differentiated cell 1 day after damage to a proliferative cell at 2

days after damage to a potentially neurogenic progenitor at 3 days after damage. The

stage at which the Notch-pathway is blocked after damage determines the outcome on

the cell-fate of glial-derived progenitors.

57 From our observations, it is clear that Notch-mediated effects on regeneration are limited. We speculate that additional signaling pathways influence this process.

For example, Osakada et al have shown that activation of the Wnt pathway in rodent

retinas stimulates proliferation of Müller glia and promotes their de-differentiation

into retinal progenitors (Osakada et al. 2007). Further, Shh signaling is known to be

an important modulator of retina regeneration (Spence et al. 2004) and has been

shown to enhance glial-derived regeneration in a mammalian retina (Wan et al. 2007).

Conclusions

We conclude that Notch-signaling plays a limited but important role during

retinal regeneration. Components of the Notch-signaling pathways are transiently

upregulated in proliferating Müller glia after damage in a chicken retina and blocking

Notch after damage alters the cell-fates of glial-derived progenitors. Further,

inhibition of γ-secretase activity associated with Notch and silencing of the bHLH

effectors Hes1 and Hes5 have distinctly different outcomes on cell-fate specification

of cultured chicken retinal progenitors.

CHAPTER 3

Notch-signaling influences neuroprotective and proliferative properties

of Müller glia

Abstract

Notch-signaling is known to play important roles during retinal development.

Recently, Notch-signaling has been shown to be active in proliferating Müller glia in

acutely damaged chick retina (Hayes et al. 2007). However, the roles of Notch in

mature, undamaged retina remain unknown. Thus, the purpose of this study was to

determine the role of the Notch-signaling pathway in the postnatal retina. Here we

show that components of the Notch-signaling pathway are expressed in most Müller

glia at low levels in undamaged retina. The expression of Notch-related genes varies

during early postnatal development and across regions, with higher expression in peripheral versus central retina. Blockade of Notch activity with a small molecule

inhibitor prior to damage was protective to retinal interneurons, amacrine and bipolar

cells, and projection neurons, ganglion cells. In the absence of damage, Notch is

upregulated in retinas treated with insulin and FGF2; the combination of these factors

is known to stimulate the proliferation and de-differentiation of Müller glia (Fischer

et al. 2002b). Inhibition of Notch-signaling during FGF2-treatment potently inhibits

levels of the downstream effectors of the MAPK- signaling pathway - p38 MAPK

59 and pCREB in Müller glia. Further, inhibition of Notch activity potently inhibits

FGF2-induced proliferation of Müller glia. Taken together, our data indicate that

Notch-signaling is downstream of and is required for FGF2/MAPK- signaling to drive the proliferation of Müller glia. In addition, our data suggest that low levels of

Notch-signaling in Müller glia diminish the neuroprotective activities of these glial cells.

Introduction

The Notch-signaling pathway is a highly conserved cell-signaling system that regulates many aspects of embryonic development (Louvi and Artavanis-Tsakonas

2006). In the developing vertebrate nervous system, including the retina, Notch- signaling maintains progenitor cells in an undifferentiated state during neurogenesis

(Dorsky et al. 1997; Gaiano and Fishell 2002; Henrique et al. 1997; Louvi and

Artavanis-Tsakonas 2006; Rapaport and Dorsky 1998). Further, the Notch pathway has been implicated in promoting glial cell fates at later stages of retinal histogenesis

(Furukawa et al. 2000; Gaiano and Fishell 2002; Gaiano et al. 2000; Vetter and

Moore 2001). Elsewhere in the developing nervous system, Notch has been shown to promote cell survival (Breunig et al. 2007; Mason et al. 2006; Oishi et al. 2004).

Notch encodes a 300kDa heterodimeric transmembrane receptor that binds to transmembrane ligands (Delta, Serrate or Jagged) on the surface of adjacent cells.

Receptor-ligand binding initiates the γ-secretase-dependent cleavage of the Notch intracellular domain which is translocated to the nucleus to affect transcription. The major effector of the Notch-signaling pathway is the DNA-binding protein

60 Suppressor-of-hairless, which upregulates a class of basic helix-loop-helix transcriptional repressors, Hairy and Enhancer of Split (HES) genes.Although the mechanisms of Notch-signaling are highly conserved across species, the functions of the signaling differs with the cellular context and stage of development.

The Notch pathway has been shown to play a role in retinal regeneration in lower vertebrates such as fish (Sullivan et al. 1997; Yurco and Cameron 2005) and newts (Nakamura and Chiba 2007). In addition, a recent study has shown that Notch- signaling is upregulated in Müller glia in acutely damaged avian retina (Hayes et al.

2007). In response to excitotoxic injury, Müller glia in the chick retina de- differentiate, proliferate and generate a few neurons (Fischer and Reh 2001). During this regenerative process, these glia upregulate Notch-signaling; this signaling is necessary for glial de-differentiation and proliferation (Hayes et al. 2007). Thereafter,

Notch-signaling inhibits the neuronal differentiation of newly-generated cells (Hayes et al. 2007). Surprisingly, the role of Notch-signaling in normal Müller glia in undamaged retina remains unknown.

In previous studies, we have shown that the Mitogen Activated Protein Kinase

(MAPK) pathway stimulates the proliferation and transdifferentiation of Müller glia

(Fischer et al. 2009a; Fischer et al. 2009b). The MAPK pathway is activated by secreted factors such as insulin, insulin-like growth factors (IGFs) and fibroblast growth cactors (FGFs) (Grewal et al. 1999). In the absence of damage, consecutive daily injections of the combination of insulin and FGF2, but not either factor alone, stimulate Müller glia to proliferate, transdifferentiate and produce a few new neurons

(Fischer et al. 2002b). Treatment with the combination of insulin and FGF2 induces a

61 response in Müller glia similar to that observed in NMDA-damaged retinas (Fischer and Reh 2003a). Intraocular injections of FGF2 activate MAPK signaling in the retina; this signaling is manifested as accumulations of pERK1/2, pCREB, cFos, Egr1 and p38 MAPK in Müller glia, but not other types of retinal cells (Fischer et al.

2009b). Small molecule inhibitors to ERK1/2 and the FGF receptor suppress the proliferation of Müller glia in NMDA-damaged retinas (Fischer et al. 2009b). Taken together, these findings indicate that active MAPK signaling is required in damaged retinas, and may be sufficient in undamaged retinas, to drive the proliferation and transdifferentiation of Müller glia. Although Notch and MAPK pathways are known to be important during glia-mediated retinal regeneration, interactions between these pathways remain unknown.

Here we investigate the expression patterns and roles of the Notch pathway components in the postnatal chicken retina. Further, we investigate the interaction of the Notch pathway with the MAPK pathway in regulating Müller glial proliferation and transdifferentiation.

Materials and Methods

Animals

The use of animals was in accordance with the guidelines established by the

National Institutes of Health and the Ohio State University. Newly hatched leghorn chickens (Gallus gallus domesticus) were obtained from the Department of Animal

Sciences at the Ohio State University and kept on a cycle of 12-h light, 12-h dark

62 (lights on at 7:00 a.m.). Chicks were housed in a stainless steel brooder at about 28°C

and received water and PurinaTM chick starter ad libitum.

Intraocular Injections

Intraocular injections were performed as described previously (Fischer et al.

1999; Fischer et al. 1998). In all experiments, 20µl of vehicle containing the test compound was injected into the experimental (right) eye, and 20µl of vehicle alone was injected into the control (left) eye. The vehicle was sterile saline (0.9% NaCl in dH2O) containing bovine serum albumin, 50 µg/mL, as carrier for NMDA,

colchicine, FGF2 and insulin, and 50% DMSO in sterile saline/BSA for DAPT. Test compounds included NMDA (N-methyl-D-aspartate; 1 µmol/dose; Sigma-Aldrich),

Colchicine (500ng/dose; Sigma-Aldrich), recombinant human FGF2 (250 ng/dose;

R&D Systems), purified bovine insulin (1 µg/dose; Sigma-Aldrich), DAPT (N-[N-

(3,5-Difluorophenacetyl-L-alanyl)]-S-phenylglycine t-butyl ester) (100µmol/dose;

Sigma Aldrich).

We used the following injection paradigms: (1) On posthatch day 5 (P5) the right eye received a single injection of DAPT and the left eye received vehicle.

Retinas were harvested 24 hrs later. (2) On P5 and P6 the right eye received an injection of DAPT and the right eye received NMDA or Colchicine. Retinas were harvested 1 day, 2 days or 10 days later. (3) On P4, P5 and P6 the right eye received an injection of 1µg insulin and 250ng FGF2 and the left eye received vehicle. On P7, both eyes received an injection of 2 µg BrdU, and retinas were harvested 24 hrs or 48

63 hrs later. (4) On P5 and P6 the right eye received an injection of 250ng FGF2 and the

left eye received vehicle. Retinas were harvested 24 hrs later. (5) On P4 both eyes

received 1 µmol NMDA. On P5 the right eye received 250ng FGF2 and the left eye

received vehicle. On P6 the right eye received 250ng FGF2 and 2µg BrdU and the left eye received 2µg BrdU alone. On P7, retinas were harvested 24 hrs later. (6) On

P4 both eyes received 1 µmol NMDA. On P5 the right eye received 250ng FGF2 +

100µM DAPT and the left eye received 250ng FGF2 alone. On P6 the right eye

received 250ng FGF2 + 100µM DAPT + 2µg BrdU and the left eye received 250ng

FGF2 + 2µg BrdU. On P7, retinas were harvested 24 hrs later. (7) On P4 and P5 the

right eye received 250ng FGF2 + 100µM DAPT and the left eye received 250ng

FGF2 alone. On P6 both eyes received 1 µmol NMDA. On P7 both eyes received 2µg

BrdU and retinas were harvested 24 hrs later.

Reverse Transcriptase PCR

After removing the eye from the eyecup, a blade was used to cut away the posterior portion of the eye containing the central retina (approximately 6mm diameter) from the anterior portion containing the peripheral retina (approximately within 5mm of the CMZ). mRNA was harvested from central peripheral regions of the retinas dissected in cold HBSS by using the RNeasy Kit (Quiagen) as per the manufacturer's instructions. Purified RNA was re-suspended in 100 µL RNAse-free water and DNA removed by using DNAse I (Ambion). cDNA was synthesized from mRNA by using oligo dT primers and SuperscriptTM III First Strand Synthesis System

64 (Invitrogen) according to the manufacturer's protocol. Control reactions were performed by excluding the reverse transcriptase to assess whether primers were amplifying genomic DNA.

PCR primers were designed by using the web-based program Primer3 from the Whitehead Institute for Biomedical Research (http://frodo.wi.mit.edu/). Primer sequences and sequence lengths are presented in Table 3.1. Standard PCR reactions were performed, in which cDNA was amplified for 20 cycles using standard protocols, PlatinumTM Taq (Invitrogen) or TITANIUMTM Taq (Clontech) and an

Eppendorf thermal cycler. The standard PCR products were run on an agarose gel and stained with ethidium bromide to determine the predicted product sizes.

Table 3.1: Primer sets used for Standard and Real-Time PCR. The table lists 19- 21 bp forward and reverse primer sets for the appropriate gene, along with the expected PCR product size for each primer set.

66 Real-time PCR

cDNA was synthesized as described using standard protocols as described before

(Fischer et al. 2004a). mRNA was harvested from retinas, reverse-transcribed to generate cDNA, and amplified using real-time PCR for 40 cycles. Real-time PCR was performed using the StepOnePlus™ Real-Time PCR System (Applied Biosystems,

Foster City, CA, USA) according to the manufacturer's instructions. Reactions were performed in triplicates, in 25 µl volumes with 0.5 µM primers and MgCl2 concentration optimized between 2-5 mM. Nucleotides, Taq DNA polymerase, and buffer were included in the SYBR® Green PCR Master Mix (Applied Biosystems,

Foster City, CA, USA). A typical protocol designed on StepOne Software v2.0

(Applied Biosystems, Foster City, CA, USA) included a holding stage for 10 min at

PCR were normalized to GAPDH and the fold difference between control and treated samples was determined using the ∆Ct method and represented as a percentage change from baseline.

In Situ Hybridization

A partial length clone of cNotch1 (1043bp) was kindly provided by Dr.

Domingos Henrique (Inst. de Medicina Molecular, Lisboa, Portugal). Riboprobes

67 were synthesized using a kit provided by Roche and stored at -80°C until use. Ocular

tissues from P7 eyes were dissected in RNase-free Hanks Balanced Salt Solution

(HBSS; Invitrogen), fixed overnight at 4°C in 4% paraformaldehyde buffered in 0.1

M dibasic sodium phosphate, and embedded in OCT-compound (Tissue-Tek).

Cryosections were processed for in situ hybridization with digoxigenin-labeled RNA

probes as described previously (Fischer et al. 2002a; Fischer et al. 2004a).

Hybridization was detected by using Fab fragments to digoxigenin that were conjugated to alkaline phosphatase (anti-DIG-AP; Roche) diluted in MABT (0.05 M maleic acid buffer, 0.1% Tween-20) plus 10% normal goat serum, 10 mM levamisole, and 10 mM glycine. Nitro-blue tetrazolium (NBT) and 5-Bromo-4-Chloro-3

Indolyphosphate p-Toluidine (BCIP) in 0.1 M NaCl, 0.1 M tris-HCl pH 9.5, 0.05 M

MgCl2 and 0.01% Tween-20 were used to precipitate chromophore from the anti-

DIG-AP. With sufficient levels of chromophore developed, the sections were washed

in PBS, fixed in 4% paraformaldehyde in PBS for 15 mins, washed again in PBS and processed for immunolabeling for 2M6, BrdU and PCNA as described ahead.

Fixation, Sectioning and Immunocytochemistry

Tissue dissection, fixation, cryosectioning, and immunolabeling were performed as described previously (Fischer et al. 2002b; Fischer et al. 2009b). Working dilutions and sources of antibodies used in this study included; goat anti-Egr1 was used at

1:1,000 (AF2818; R&D Systems); mouse anti-2M6 used at 1:100 (Dr. P. Linser,

University of Florida); anti-Sox9 used at 1:2,000 (AB5535; Chemicon); rabbit anti-

GFAP used at 1:2000 (Z0334; DakoCytomation); mouse anti-transitin used at 1:50

68 (EAP3; DSHB); mouse anti-vimentin used at 1:80 (H5; DSHB); mouse anti-Brn3a used at 1:50 (MAB1585; Chemicon); mouse anti-PCNA used at 1:1,000 (M0879;

DAKO); rat anti-BrdU used at 1:200 (OBT0030S; Serotec); mouse anti-BrdU used at

1:100 (G3B4; DSHB); rabbit anti-p38 MAPK used at 1:80 (12F8; Cell-signaling

Technologies); rabbit anti-pCREB used at 1:600 (87G3; Cell-signaling

Technologies); rabbit pERK1/2 used at 1:200 (137F5; Cell-signaling Technologies) and rabbit anti-Egr1 used at 1:400 (sc-110; Santa Cruz). Secondary antibodies included donkey-anti-goat-Alexa488, goat-anti-rabbit-Alexa488, goat-anti-mouse-

Alexa488/568/647, goat-anti-rat-Alexa488, rabbit anti-goat Alexa488 and goat-anti- mouse-IgM-Alexa568 (Invitrogen) diluted to 1:1,000 in PBS plus 0.2% Triton X-100.

To permeabilize retinas for whole-mount labeling procedures, samples were frozen (-

80°C) and thawed (20°C) three times prior to incubation with the antibody solution.

Both primary and secondary antibodies were incubated overnight.

TUNEL

To identify dying cells that contained fragmented DNA we used the TUNEL method.

We used an In Situ Cell Death Kit (TMR red; Roche Applied Science), as per the manufacturer's instructions.

69 Photography, Measurements, Cell Counts, and Statistical Analyses

Photomicrographs were taken by using a Leica DM5000B microscope equipped with

epifluorescence and a 12 megapixel Leica DC500 digital camera. Confocal

microscopy was done by using a Zeiss LSM 510 at the Hunt-Curtis Imaging Facility

in The Ohio State College of Medicine. Images were optimized for color, brightness

and contrast, and double-labeled images overlaid by using Adobe PhotoshopTM6.0.

Cell counts were made from at least six different animals, and means and standard

deviations calculated on data sets. To avoid the possibility of region-specific differences within the retina, cell counts were consistently made from the same region of retina for each data set. Immunofluorescence was quantified by using ImagePro

6.2. Identical illumination, microscope and camera settings were used to obtain images for quantification. Areas (800 × 200 pixels or 232 × 54 µm) were sampled

from 5.4 MP digital images. The areas were randomly sampled over the INL where

dying cells, activated glia or proliferating glia were found. Measurements were made

for pixels with intensity values (0 = black, 255 = saturated) of >72 for pCREB and

>65 for p38 MAPK; thresholds that included labeling of cells in control and treated

samples. The total area was calculated for regions with pixel intensities above

threshold and the density sum was calculated as the total of pixel values for all pixels

within thresholded regions. Measurements were made for IPL regions sampled from

at least six different retinas for each experimental condition.

70 Results

Components of the Notch-signaling pathway are expressed at low levels in the

postnatal chicken retina

To determine whether components of the Notch-signaling pathway are

expressed in the postnatal chick retina, we used in situ hybridization,

immunohistochemistry and RT-PCR. We found that some cells in the inner nuclear

layer (INL) of the P7 retina normally express cNotch1 mRNA at low levels. These

cells were immunoreactive for 2M6 (Fig. 3.1a-g), a monoclonal antibody that is

known to selectively label Müller glia in the chicken retina (Fischer et al. 2009b;

Linser et al. 1997). These findings suggest that Müller glia maintain low levels of

Notch expression in undamaged, mature retina.

Notch pathway gene expression varies over time in the adult retina

Consistent with the findings of ISH studies, RT-PCR detected mRNA for

cNotch1, cHes1 and cHes5 in undamaged postnatal chicken retina (Fig. 3.1h). The

expression of Hes1 and Hes5 is a read-out of active Notch-signaling (Kageyama and

Ohtsuka 1999; Nelson et al. 2006). These findings suggest that low levels of Notch-

signaling are sustained in mature Müller glia. PCR products were sequenced to verify

the specificity of the reactions. We next assayed for changes in expression levels of

Notch and related genes in central and peripheral regions of the postnatal retina and

embryonic retina using real-time PCR. We found that the expression levels of

cNotch1 are significantly higher in peripheral retina compared to levels in central retina at P0 and P14 (Fig. 3.1i). Further, expression levels of cNotch1 were reduced

71 over time in the peripheral retina, compared to expression levels at P0. Interestingly, expression of cHes1 and cHes5 followed a similar pattern of expression as cNotch1. cHes1 expression was significantly higher in peripheral retina compared to central retina at P0, P7 and P21. cHes1 expression in central retina remained relatively low and unchanged throughout the first 3 weeks of postnatal development (Fig. 3.1j). By comparison, cHes5 expression declined rapidly in peripheral retina from P0 through

P7, whereas cHes5 remained low, and unchanged, in central retina throughout postnatal development similar to cHes1 expression (Fig. 3.1k). As expected, expression levels of cNotch1, cHes1 and cHes5 were much higher in embryonic retina compared to postnatal retina (Figs. 3.1i-k). Interestingly, cHes5 expression in embryonic retinas was 64-fold higher than that in central retina at P0 (Fig. 3.1k). The differences in expression levels of cNotch1 and cHes1 in embryonic and postnatal retinas were modest compared to differences seen for levels of cHes5 (Figs. 3.1i and j).

Figure 3.1: Components of the Notch-signaling pathway are expressed at low levels in the postnatal chicken retina. Vertical sections of the P7 retina were hybridized with riboprobes to cNotch1 (a; blue) and labeled for the glial marker 2M6 (b; red). Panel c is a merged image of panels a and b. Panels d-g are 7-fold enlargements of corresponding boxed areas in panel c. Arrows in panel c indicate cNotch1 and 2M6 double-positive cells. The scale bar (50µm) in panel a applies to panels a - c. mRNA was harvested from retinas obtained from P7 chickens, reverse- transcribed to generate cDNA, amplified using (h) PCR or (i-k) real-time PCR. The PCR products were run on an agarose gel, and stained with ethidium bromide (h). Reactions minus the reverse transcriptase enzyme (RT-) were used as a negative control for each primer set. The percentage change in cDNA levels for cNotch1 (i), cHes1 (j) and cHes5 (k) were calculated from C(t) values that were normalized to GAPDH. Relative expression levels were measured for each gene at P0 in central retina (set as baseline) and those at P7, P14 and P21 in central and peripheral retina and E3 whole retina. Significance of difference (*p<0.05 between two timepoints within the same region , #p<0.05 between central and peripheral retina at the same timepoint) was determined by using a two-tailed, unpaired Student’s t-test. Abbreviations: RT – reverse transcriptase, INL – inner nuclear layer, IPL – inner plexiform layer. 73 Inhibition of Notch-signaling in the retina does not affect glial reactivity

To determine whether Notch-signaling influences Müller glia in mature retina,

we inhibited Notch-signaling by using a γ-secretase inhibitor, DAPT. Using conventional PCR (Fig. 3.2a) and real-time PCR (Fig. 3.2b), we observed that DAPT significantly reduced the expression levels of cNotch1, cDelta1, cDll4, cHes1 and cHes2 in the retina, whereas levels of cHes5 were unaffected (Fig. 3.2b). However, there was no apparent change in the expression levels of Sox9 (Fig. 3.2c, d), GFAP

(Fig. 3.2e, f), transitin (Fig. 3.2g, h) and vimentin (Fig. 3.2 i, j), genes that are normally expressed by Müller glia in the mature retina. Transitin and GFAP are known to be upregulated in reactive Müller glia (Fischer and Omar 2005).

Figure 3.2: Inhibition of Notch-signaling reduces the expression of components of the Notch-signaling pathway in undamaged postnatal chicken retina. mRNA was harvested from central and peripheral retinas obtained from P7 chickens that were injected with vehicle (Control) or DAPT (Treated), reverse-transcribed to generate cDNA, and amplified using (a) PCR or (b) real-time PCR. The PCR products were run on an agarose gel, and stained with ethidium bromide (a). C(t) values were normalized to GAPDH and the fold difference between control and treated samples was determined using the ∆Ct method and represented as a percentage change from baseline (b). (c-j) Representative images from retinas harvested 24 hrs after the last dose of vehicle or DAPT. Vertical sections of the P7 retina were labeled for Sox9 (c-d), GFAP (e-f), transitin (g-h) or vimentin (i-j). Abbreviations: RT – reverse transcriptase, ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer. The scale bar (50µm) in panel j applies to panels c – j.

75 Intraocular injections of a Notch inhibitor prior to retinal injury reduces cell death

Given that some of the Müller glia normally express cNotch1 at low levels and Notch-signaling is upregulated in damaged retinas (Hayes et al. 2007), we sought to determine whether blocking Notch-signaling influenced glial responses to acute damage. NMDA-induced retinal damage is known to stimulate the proliferation of

Müller glia in birds and rodents (Fischer and Reh 2001; Karl et al. 2008); this proliferation is thought to be an integral step in the transdifferentiation of Müller glia

(Fischer and Reh 2003b; Lamba et al. 2009). We found that inhibition of Notch- signaling in Müller glia prior to NMDA-treatment caused a decrease in the number of proliferating glia (data not shown).

To test whether inhibition of Notch-signaling influenced the ability of Müller glia to protect retinal neurons from damage, we injected DAPT prior to NMDA or colchicine treatment and harvested the retinas 1 day, 2 days or 10 days later. NMDA results in excitotoxic death of amacrine and bipolar cells (Fischer et al. 1998) and colchicine results in death of ganglion cells (Fischer et al. 1999; Morgan 1981;

Morgan and Spooner 1983). We found that two consecutive daily doses of 865 ng

DAPT significantly reduced NMDA-induced cell death by 53% in central regions of the retina with no change in cell death in nasal or temporal retina (Fig. 3.3 a-c). With a lower dose of NMDA (30 µg), DAPT pre-treatment lowered the abundance of

TUNEL+ cells, with a 50% decrease in cell death in central regions, a 43% decrease in cell death in temporal regions and no significant changes in cell death in nasal regions of the retina (Fig. 3.3d).

76 We next assayed for the survival of ganglion cells in colchicine-damaged

retinas by probing for Brn3a, a homeodomain transcription factor that is expressed by

~98% of the ganglion cells (Badea et al. 2009; Liu et al. 2000). DAPT prior to

colchicine-treatment resulted in 55%, 100% and 62% more surviving ganglion cells

in dorsal, nasal and temporal regions of the retina, respectively, at 4 days after injury

(Figs. 3.3e-g). However, in colchicine-treated retinas, numbers of dying ganglion cells are maximal between 3 and 4 days after treatment (Fischer and Reh 2002;

Fischer et al. 2008). Thus, we next assayed for ganglion cell survival at 10 days after treatment when the damaging effects of colchicine had subsided. With DAPT- pretreatment, elevated survival of ganglion cells was seen at 10 days after injury in the nasal retina (Fig. 3.3h). However, the most prevalent survival-promoting effects for DAPT were observed in temporal regions of the retina with a near 3-fold increase in surviving ganglion cells compared to retinas 10 days after treatment with colchicine alone (Fig. 3.3h). Collectively, our results suggest that DAPT enhances neuronal survival when administered prior to injury by inhibiting Notch-signaling in

Müller glia.

Figure 3.3: Inhibition of Notch-signaling prior to retinal injury decreases cell death and increases neuronal survival. Representative images of TUNEL-positive nuclei in retinas that were treated with (a) vehicle before NMDA or (b) DAPT before NMDA. (c, d) Histograms illustrate the number of dying cells in the INL in central, nasal or temporal regions of the retina. Representative images of whole-mounted retinas that were treated with (e) vehicle before colchicine or (f) DAPT before colchicine. Retinas were labeled with antibodies to Brn3a which is expressed by ganglion cells. (g, h) Histograms illustrate the mean number of ganglion cells per 0.55mm2 in central, dorsal, nasal or temporal regions of the retina. Data sets were obtained from 6 animals per treatment. Significance of difference (*p<0.05, **p<0.01) was determined by using a two-tailed, unpaired Student’s t-test. Abbreviations: ONL – outer nuclear layer, INL – inner nuclear layer. The scale bar (50µm) in panel b applies to panels a and b and that in panel f applies to panels e and f.

78 Notch is upregulated in retinas treated with growth factors insulin and FGF2

Previous studies have shown that Notch is upregulated in Müller glia after retinal injury (Hayes et al. 2007; Yurco and Cameron 2007). However, it remains

uncertain whether glial increases in Notch-signaling in response to damage participate

exclusively in the process of proliferation and transdifferentiation or in the process of

damage-induced reactive gliosis. Accordingly, we tested whether Notch is

upregulated in Müller glia when stimulated to proliferate and transdifferentiate in the

absence of injury. We have previously shown that three consecutive daily doses of

insulin and FGF2 result in a peripheral-to-central wave of proliferation within the

retina and that these dividing cells are Müller glia (Fischer et al. 2002b). In response

to insulin and FGF2, cNotch1 was upregulated in proliferating Müller glia (Fig. 3.4a-

g). This effect was also seen in retinas treated with four consecutive daily doses of

IGF1 and FGF2 (data not shown). Saline injections did not influence cNotch1

expression or the proliferation of cells (Fig. 3.4h-j).

Figure 3.4: cNotch1 is upregulated by proliferating Müller glia in retinas treated with insulin and FGF2. Eyes were injected with three consecutive daily doses of insulin and FGF2 or saline (control) staring at P4, BrdU at P7 and harvested at P8. Vertical sections of the P8 retina were hybridized with riboprobes to cNotch1 (a; blue) and subsequently labeled for PCNA (b; red) and BrdU (c; green). Panel d is a merged image of panels a - c. Panels e-g are 3.5-fold enlargements of corresponding boxed areas in panel d. Control retinas did not show an upregulation of cNotch1 (h), PCNA (i) or BrdU (j). White arrows in panel b indicate PCNA positive cells. Yellow arrows in panel c and d indicate BrdU-positive cells. * in panels e-g indicate cNotch1-positive PCNA or BrdU-positive cells. Abbreviations: ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer. The scale bar (50µm) in panel a applies to panels a – c and h – j whereas the scale bar in panel d (50µm) applies to panel d alone.

80 Using real-time PCR, we observed that three consecutive daily doses of insulin and FGF2 induced an upregulation of cNotch1 and related genes. We found a

3.5-fold increase in cNotch1, 15-fold increase in cDelta1, 12-fold increase in Dll4,

4.5-fold increase in cHes5, but no change in cHes1 (Fig. 3.5a). We also found significant increases in the expression of other Notch-related genes such as a 2-fold increase in the ligand cJagged, 4-fold increase in cHey1, 12-fold increase in cHey2, but no change in the proneural gene ascl1a (Fig. 3.5a). To determine whether levels of expression of Notch and associated genes were influenced by FGF or insulin alone, we injected either growth factor separately and harvested retinas 24 hrs after the last injection. Two consecutive daily doses of FGF2 induced a 6-fold increase in cNotch1 expression, 4.5-fold increase in cDll4, 5.7-fold increase in cJagged, 3-fold increase in cHes5 and cHey1, 5-fold increase in cHey2 and a small decrease in cHes1 expression, but did not influence levels of cDelta1 and ascl1a (Fig. 3.5b). Two consecutive daily doses of insulin, however, did not affect levels of cNotch1, and resulted in modest but significant decreases in expression levels of cDelta1 (0.3-fold), cHes1 (0.55-fold), cHes5 (0.25-fold), cHey2 (0.3-fold) and ascl1a (0.15-fold) (Fig. 3.5c). Interestingly, insulin induced small but significant increases in expression levels of cDll4 (0.2- fold), cJagged (0.7-fold) and cHey1 (0.7-fold) (Fig. 3.5c).

Figure 3.5: Notch and related genes are upregulated in retinas treated with insulin and FGF2. mRNA was harvested from retinas obtained from eyes treated with saline (control) or insulin and FGF2 (a), saline (control) or FGF2 (b) and saline (control) or insulin (c) mRNA was reverse-transcribed to generate cDNA and amplified using real-time PCR. Ct values were normalized to GAPDH and the fold difference between control and treated samples was determined using the ∆Ct method and represented as a percentage change (a-c). Significance of difference (*p<0.03) was determined by using a two-tailed, unpaired Student’s t-test. Abbreviations: RT – reverse transcriptase

82 FGF2 upregulates effectors of MAPK signaling in Müller glia and stimulates

proliferation in damaged retinas

We have previously shown that the damage caused by 2µmol NMDA

upregulates retinal levels of FGF1 and FGF2 (Fischer et al. 2004), induces the

accumulation of pCREB in Müller glia at 2-3 days after injury, and that inhibition of

FGF receptors and MEK suppresses the proliferation of Müller glia (Fischer et al.

2009a). In the absence of damage, FGF2 induces the accumulation of MAPK

effectors, including pERK1/2, cFos, Egr1, pCREB and p38 MAPK in Müller glia,

whereas retinal neurons appear unaffected. Further, FGF2 is neuroprotective and

enhances Müller glial proliferation when administered prior to NMDA (Fischer et al.

2009b). It is also known that 1µmol of NMDA causes some cell death, but fails to

induce Müller glial proliferation as is elicited by the maximum dose (2µmol) (Fischer

et al. 2009b). Hence, we tested whether FGF2 induced MAPK effectors and Müller

glial proliferation in the retina after moderate levels of damage. We administered

1µmol NMDA into eyes followed by two daily doses of saline (control) or FGF2

(treatment) and BrdU to label proliferating cells. We found that FGF2 administered

after NMDA greatly enhanced Müller glial proliferation (Fig. 3.6 a- i). Sox9-positive

Müller glia expressed high levels of PCNA and accumulated BrdU in retinas treated

with FGF2 following NMDA (Fig. 3.6 e-h). Although low levels of PCNA were

detected in Müller glia in retinas treated with NMDA alone, few of these cells

accumulated BrdU and levels of PCNA appeared much lower than those seen in

Müller glia in retinas treated with NMDA and FGF2 (Fig. 3.6a-h). Thus, the Müller

glia with low levels of PCNA likely were not proliferating, and only those cells

83 labeled for BrdU and high levels of PCNA were counted as proliferating. We also

found that FGF2 increased levels of p38 MAPK and pCREB in Müller glia (Fig. 3.6

j-u), but not pERK, Egr1 and cFos (data not shown). Thus, FGF2 induces active

MAPK signaling in Müller glia and induces proliferation of these cells after moderate levels of injury.

Figure 3.6: FGF2 induces the proliferation of Müller glia in injured retinas. Eyes were treated with 1000 nmol of NMDA at P4 followed by two consecutive daily doses of saline (control) or 250ng FGF2 (treated) at P5 and P6, BrdU at P6 and harvested 24 hrs later at P7. Vertical sections of the P7 retina were labeled for Sox9 (a, e; magenta), BrdU (b, f; green) and PCNA (c, g; red). Panel d is a merged image of panels a - c. Panel h is a merged image of panels e-g. Panels d and h are 3.5-fold enlargements of panels a-c and e-g respectively. The histogram in panel i represents the mean number of BrdU and PCNA positive cells per 15000µm2 in control and treated retinas (j-u). Vertical sections from control and treated retinas were labeled for 2M6 (red) and p38 MAPK (green; j-o) or pCREB (green; p-u). Significance of difference (**p<0.001) was determined by using a two-tailed, unpaired Student’s t- test. Arrows indicate 2M6 positive Müller glia labeled for p38 MAPK or pCREB. Abbreviations: INL – inner nuclear layer. The scale bar (50µm) in panel g applies to panels a – c, e – g and that in panel j applies to panels i – l. 85 Inhibition of Notch reduces FGF2-induced accumulations of p38 MAPK and pCREB in Müller glia

Some studies have indicated that interactions exist between the Notch and the

FGF2-MAPK pathway in the developing nervous system (Faux et al. 2001; Ota and

Ito 2006; Saravanamuthu et al. 2009; Wahl et al. 2007). However, such an interaction has not been demonstrated in the retina. Since components of the Notch-signaling pathway were upregulated by FGF2 (Fig. 3.4, 3.5), we tested whether Notch reciprocally influences FGF2-MAPK signaling. Specifically, we tested whether

Notch activity is required for the upregulation of pCREB and p38 MAPK in damaged retinas treated with FGF2. After treatment with 1µmol NMDA, two consecutive daily doses of FGF (control) or FGF and DAPT (treatment) were injected into eyes that were harvested 24 hrs after the last dose. The FGF-induced accumulations of pCREB

(Fig. 3.7a, c) and p38 MAPK (Fig. 3.7e) in Müller glia in damaged retina were significantly reduced by DAPT (Fig. 3.7b, d-f and h-j). We did not observe any change in expression levels of FGFR1 or FGFR3 mRNA in this paradigm (data not shown), suggesting that DAPT-treatment acts on downstream components of the

FGF-MAPK pathway rather than inhibiting receptor-expression levels. It should be noted that we were unable to detect significant levels of FGFR2 in the retina consistent with previous reports (Fischer et al. 2009a), and that FGFR4 may not exist in chicken retina (Scott and Fischer, unpublished observation).

Figure 3.7: Inhibition of Notch-signaling reduces levels of p38 MAPK and pCREB in Müller glia. DAPT suppresses the accumulation of p38 MAPK and pCREB in Müller glia that results from NMDA+FGF2 treatment. Images were obtained using identical camera exposures and microscope illumination settings. Retinas were processed for immunolabeling after treatment with 1000 nmol of NMDA and two consecutive daily doses of 250ng FGF2 (control) or 250ng FGF2 + 100µM DAPT (treated). As described in the Methods, ImagePro 6.2 was used to obtain measurements of total area for pixel intensities >72 for pCREB and >65 for p38 MAPK (0 = black, 255 = saturated green), and the density sum. The small numbers and red areas in panels c, d, g and h indicate the pixels designed by ImagePro 6.2 that met the threshold criteria. Means and standard deviations are displayed in histograms for total area (e and i) and density sum (f and j) for areas with pixels above threshold. Significance of difference (**p<0.001) was determined by using a two-tailed, unpaired Student’s t-test. Abbreviations: ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer. The scale bar (50µm) in panel b applies to panels a – h.

87 Inhibition of Notch activity reduces FGF-induced proliferation of Müller glia

Since DAPT can inhibit the accumulation of p38 MAPK and pCREB in

Müller glia (Fig. 3.7), we tested whether DAPT influenced FGF2-induced glial proliferation in damaged retinas. To do this, we injected 1µmol NMDA into eyes at

P4 followed by 2 injections of FGF2 (control) or FGF2 and DAPT (treatment) at P5 and P6, and BrdU at P6 to label proliferating cells. Eyes were harvested at P7, 3 days after the excitotoxic insult. As expected, FGF2 stimulated Müller glial proliferation after NMDA-injury (Fig. 3.8a-d) to levels similar to those seen 3 days after 2µmol

NMDA alone (Fig. 3.8i). Interestingly, DAPT blocked the FGF2-induced proliferation of Müller glia (Fig. 3.8e-h), suggesting that FGF2/MAPK-signaling requires active Notch-signaling to stimulate the proliferation of Müller glia in damaged retinas.

In another paradigm, we injected FGF2 (control) or FGF2 and DAPT

(treatment) at P4 and P5 followed by 1µmol NMDA into eyes at P6 and BrdU at P7 to label proliferating cells. Eyes were harvested 4 hrs after the BrdU injection. Similar to the results seen when FGF2 and DAPT were injected after NMDA, we observed that DAPT blocked the proliferation-inducing effects of FGF2 when applied before

NMDA (Fig. 3.8i).

Figure 3.8: Inhibition of Notch-signaling blocks FGF2-induced proliferation of Müller glia in NMDA-damaged retinas. Eyes were injected with 1000nmol NMDA before or after 2 daily doses of 250ng FGF2 (control) or 250ng FGF2 and 100µM DAPT (treated), BrdU at P6 (or P7) and harvested at P7. Vertical sections of the P7 retina were labeled for Sox9 (a, e; purple), BrdU (b, f; green) and PCNA (c, g; red). Panel d is a merged image of panels a - c. Panel h is a merged image of panels e-g. Panels d and h are 4-fold enlargements of panels a-c and e-g respectively. The histograms in panel i represent the number of BrdU- and PCNA-positive cells per 15000µm2 area of the control and treated retinas. Significance of difference (**p<0.001) was determined by using a two-tailed, unpaired Student’s t-test. Abbreviations: INL – inner nuclear layer. The scale bar (50µm) in panel g applies to panels a – c and e – g.

89 Discussion

We report here that components of the Notch-signaling pathway are expressed in many Müller glia at low levels in undamaged postnatal retina where this pathway regulates the neuroprotective and proliferative properties of the glia. This pathway has been extensively studied in the developing retina, but few studies have addressed its function in the mature retina. During embryonic development, Notch and its effectors have been shown to be highly expressed by retinal progenitors in the chick retina at E4-E4.5 (Austin et al. 1995; Nelson et al. 2006) and E5.5 (Kubo and

Nakagawa 2009) and in the rat retina from E12.5-P0 (Bao and Cepko 1997).

Over time, Notch expression is confined to the peripheral retina (Bao and

Cepko 1997) and is thought to be completely absent in central retina, although some expression in the adult retina has been reported (Ahmad et al. 1995; Hayes et al.

2007). Our findings indicate that low levels of Notch expression are maintained in postnatal chick retina, long after the completion of retinal histogenesis which is completed about 10 days before hatching (Ghai et al. 2008; Prada et al. 1991).

Consistent with previous reports, our findings indicate large decreases in expression levels of cNotch1 and cHes5 between early embryonic and postnatal stages of development. Although we find that levels of cHes1 are decreased during retinal development, the magnitude of this decrease is a fraction of the decreases seen for cNotch1 and cHes5. Further, we find elevated expression of Notch and its downstream effectors in peripheral compared to central regions of postnatal retina.

This is consistent with reports indicating that peripheral retina matures more slowly than central retina (Ghai et al. 2008; Harman and Beazley 1989).

90 Our findings indicate that most, if not all, of the Notch expression in postnatal chick retina is restricted to Müller glia. A single cell gene expression analysis of

Müller glia in rodents revealed highly enriched expression of genes associated with

Notch-signaling, such as Nrarp, Notch1, Notch2, Cntn1 and Hes1(Roesch et al.

2008). The cell types that express Notch ligands in the postnatal retina remain unknown. In the developing retina, Notch ligands are expressed by various cell types.

Delta1 is expressed primarily in progenitor cells, Dll4 is expressed in newly differentiated neurons (Nelson and Reh 2008) whereas Jagged is expressed in postmitotic ganglion cells (Valsecchi et al. 1997). In the postnatal chick retina, these ligands are likely expressed by neurons. Microglia may also express Notch ligands; these cells have been shown to interact with Müller glia during retinal injury and influence photoreceptor survival (Harada et al. 2002). Another possibility is that

Notch ligands are expressed by Müller glia. Consistent with this hypothesis, FGF2 increases retinal levels of Delta1, Dll4 and Jagged (current study) and FGF2 acts directly on Müller glia (Fischer et al. 2009a). Further studies are required to unambiguously identify the retinal cell-types that express Notch ligands

Low levels of Notch-signaling in Müller glia in mature retina may function to suppress the neuroprotective properties of these glia. When Notch-signaling is blocked, there is decreased cell death and increased survival of neurons in ‘stressed’ retinas. We used two different injury paradigms to determine the role of Notch- signaling in Müller glia on neuronal survival. NMDA is an excitotoxin that destroys amacrine and bipolar cells in the INL (Fischer et al. 1998), whereas colchicine inhibits microtubule polymerization and hence prevents axonal transport, leading to

91 the death of ganglion cells (Fischer et al. 1999; Morgan 1981). Thus, NMDA and colchicine have different modes of action and cause different types of damage in the retina. Regardless of the type of injury, however, blocking Notch-signaling in Müller glia prior to retinal damage increased neuronal survival. An interesting observation

was that we found region-specific differences in the survival of retinal neurons with inhibition of Notch-signaling. It is possible that no significant changes in cell death

were observed in peripheral regions of the retina in the NMDA-injury paradigm

because DAPT was injected at P5 and P6, when Notch expression is relatively low in

both peripheral and central retina. Thus, there was little Notch-signaling for DAPT to

inhibit in peripheral retina when applied at or after P5. However in the colchicine-

injury paradigm, DAPT was applied early, at P0 and P1. Hence, the DAPT-mediated

survival-effect in this paradigm was relatively large in peripheral regions of the

retina, where Notch expression is higher during early postnatal development.

It is known that Müller glia protect neurons against excitotoxic cell death via

the uptake and degradation of glutamate, release of neurotrophic factors, and the

secretion of the antioxidant, glutathione (Bringmann et al. 2006). The Notch-

dependent mechanisms that regulate the ability of Müller glia to support neuronal

survival remain uncertain. Our observations indicate that Notch-signaling in Müller

glia impacts amacrine and bipolar cell survival with NMDA injury. Interestingly, we

find that inhibition of Notch significantly enhances ganglion cell survival with

colchicine injury. To our knowledge, the relationship between colchicine-induced

ganglion cell death and the responses of Müller glia has not been studied. One report

suggests that colchicine-treatment enhances IGF1 immunoreactivity in Müller glia

92 (Hansson et al. 1989). In addition, it is known that ganglion cell death stimulates

gliosis in Müller cells (Bringmann and Reichenbach 2001; Inman and Horner 2007)

but there currently is no evidence that Notch-signaling influences the gliotic

phenotypes of Müller cells. Our results indicate that Notch-signaling plays an

important role in Müller glia-mediated neuronal survival during injury to ganglion

cells. This may have important implications in the development of therapies to treat

glaucoma, where visual acuity is lost because of the death of ganglion cells.

The mechanisms that regulate the ability of Müller glia to protect or render

retinal neurons vulnerable to injury remain uncertain. Pro-survival factors such as

Ciliary Neuro Trophic Factor (CNTF) and Glial cell line-Derived Neurotrophic

Factor (GDNF) have been shown to influence primarily Müller glia and support the

survival of different types of retinal neurons (Fischer et al. 2004b; Harada et al. 2003;

Harada et al. 2002; Joly et al. 2007; Kassen et al. 2009; van Adel et al. 2005).

Similarly, FGF is known to directly act at Müller glia and protect retinal neurons

from different types of damage and degeneration (Chaum 2003; Fischer et al. 2009a;

Russell 2003). The factor-mediated changes in glial activity that effect neuronal

survival in the mature retina are yet to be determined. On the other hand, Müller glia

have been shown to play a role in several degenerative disorders of the retina and, in

certain contexts, may exacerbate neuronal cell death (Bringmann and Reichenbach

2001). It is possible that active Notch-signaling may render Müller glia less

neuroprotective in a degenerating retina. Thus, a delicate balance exists between

neuroprotective and neurodegenerative activities of Müller glia that impact the

survival of retinal neurons and maintenance of vision.

93 It is known that Notch-signaling is increased in damaged retinas and that this

signaling stimulates proliferation and regeneration from Müller glia (Hayes et al.

2007; Yurco and Cameron 2007). These studies indicate that widespread neuronal

cell death is followed by Müller glial upregulation of Notch between 1-5 days after

the initial insult. These cells then go on to proliferate and become progenitor-like

cells in the chick retina (Hayes et al. 2007). The link between Notch-signaling and

neuroprotection has not been established in damaged retinas, but remains a distinct

possibility. For example, it remains uncertain whether the upregulation of Notch-

signaling in Müller glia in damaged retinas, in part, influences the neuroprotective

properties of the glia. Our current findings are consistent with the hypothesis that

normal baseline Notch-signaling in Müller glia is required to maintain these glia in a progenitor-like state which detracts from their neuroprotective ability. We speculate that in a mature retina, the ability of Müller glia to protect neurons could be incompatible with the progenitor-like phenotype of these cells.

The link between Notch-signaling and FGF-MAPK signaling has not been established previously in the retina. We have shown that FGF2 stimulates neuroprotection and proliferation of Müller glia in damaged retinas (Fischer et al.

2009a). Our findings suggest that the effects of FGF2/MAPK upon the proliferation of Müller glia requires Notch-signaling. However, FGF2-mediated neuroprotection by Müller glia is not influenced by blocking Notch-signaling (data not shown). This indicates that the Notch-dependent mechanism by which FGF2 stimulates glial proliferation is distinct from the mechanism by which FGF2 enhances neuroprotection through Müller glia. Further, the interaction of MAPK- and Notch-

94 signaling in stimulating glial proliferation may involve a feed-forward loop. For

example, the expression of Notch and related genes is increased in Müller glia, in the

absence of damage, by insulin and FGF2. Consistent with our findings, interactions

between FGF2/MAPK and Notch are known to exist in the developing nervous

system. Faux and colleagues have demonstrated that FGF2 upregulates Notch

expression in neuroepithelial precursor cells during development and that FGF2 acts

synergistically with Notch to inhibit differentiation in these cells (Faux et al. 2001).

Ota and colleagues have demonstrated that FGF2 promotes gliogenesis and

suppresses ngn2 expression in trunk neural crest cells by increasing expression of

Notch1 and Delta1, (Ota and Ito 2006). In the mouse, FGF/MAPK signaling has been shown to control the oscillating expression of Notch pathway genes in presomitic

mesoderm during somitogenesis (Wahl et al. 2007). In the eye, crosstalk between

FGF2 and Notch pathways was shown in lens-fiber differentiation, where FGF2

induces expression of Jag1 and Notch2 (Saravanamuthu et al. 2009). DAPT

decreased the FGF2-induced Jag1 expression, demonstrating the dependence of

FGF2-signaling on Notch for normal lens development (Saravanamuthu et al. 2009).

Collectively, these reports support our findings that FGF2/MAPK-signaling requires

Notch to stimulate the de-differentiation and proliferation of mature Müller glia.

Previous findings indicate that intraocular injections of FGF alone are insufficient to stimulate the proliferation of Müller glia (Fischer et al. 2002b). The proliferation of Müller glia is potently stimulated by FGF2 only when combined with insulin (Fischer et al. 2002b), IGF1 (current studies) or low levels of NMDA-induced damage (Fischer et al. 2009a). Thus, FGF and MAPK-signaling facilitate the

95 transdifferentiation of Müller glia, but are not sufficient to drive this process. Further we find that insulin or FGF2 alone have different effects upon Notch and associated genes. FGF2 increased expression of Notch and associated downstream effectors whereas insulin does not. However, the combination of insulin and FGF2 act synergistically to dramatically increase levels of genes associated with the Notch pathway; an effect not seen with either factor alone. Expression of Notch ligands varied greatly between treatments. cDelta1 expression is not affected by FGF2- treatment and is significantly decreased by insulin-treatment or IGF1-treatment (data not shown). By contrast, the combination of insulin and FGF2 greatly enhances its expression. Interestingly, expression levels of cDll4 and cJagged are enhanced by both FGF2 and insulin and by the combination of insulin and FGF2. We also see a significant upregulation of cDll4 with IGF-FGF (data not shown).

Additionally, we find that the influence of FGF2/MAPK signaling on the expression of Notch and related genes in Müller glia is somehow enhanced by insulin and IGF1. In addition, Hes5, but not Hes1, expression increases with FGF2-treatment whereas expression of both Hes1 and Hes5 are reduced by insulin treatment.Our data suggests that the Notch pathway genes can be upregulated by FGF/MAPK signaling, but the Notch-enhancing effects of FGF2 are somehow amplified by co-application of insulin/IGF1. These differences could be explained by different sites of action within the retina. FGFR1 may be expressed by Müller glia but the insulin receptor is not detectable in the retina (Fischer et al. 2009a). We have recently proposed that insulin acts through IGF1R present on novel inner retinal glia-like cells (NIRGs), that are stimulated by insulin and IGF1 (ref in press). The NIRG cells influence Müller glia

96 through unidentified mechanisms. Thus, FGF-induced Notch-signaling may be a

direct effect on Müller glia whereas Insulin/IGF1 may inhibit Notch-signaling in

Müller glia through intermediate cells and signals. Together, insulin and FGF2 have

combined effects on Notch-signaling through direct effects on Müller glia as well as

indirect effects through NIRGs and/or microglia. We propose that the synergistic

influence of FGF2 and insulin/IGF1 on the expression of Notch and related genes in

Müller glia is needed to stimulate proliferation without neural damage.

Insulin and FGF2 are known to activate MAPK signaling in Müller glia

(Fischer et al. 2009a). Growth factor-mediated activation of ERK1/2 and p38 MAPK pathways are involved in proliferation and differentiation during development (Irving and Bamford 2002). In the retina, Müller glia nuclei accumulate pCREB and transiently express immediate early genes such as cFos and Egr1, downstream

effectors of MAPK signaling, in response to acute retinal damage (Fischer et al.

2009b). Moreover, application of FGF2, that activates MAPK signaling, prior to

NMDA protects retinal neurons against excitotoxicity (Fischer et al. 2009a). There is no known precedence for interactions between downstream effectors of MAPK signaling with the Notch pathway in the retina. Since we do not see any change in the expression level of the FGF receptors in the presence of DAPT-treated retinas (data not shown), it is likely that these pathways intersect downstream of receptor

activation. Some interactions between Notch and MAPK pathways have been shown

in other developing systems. Notch induces activity of MKP-1, a MAPK phosphatase

that inhibits p38 MAPK, thereby inhibiting myogenesis (Kondoh et al. 2007). In

specification of cardiac and muscle progenitors in the Drosophila embryonic

97 mesoderm, Ras/MAPK induces Notch and Delta expression which in turn feeds back to upregulate Ras (Carmena et al. 2002). Our data suggests that Notch-signaling may impact FGF2/MAPK by influencing the accumulation or phosphorylation of p38

MAPK and CREB. Alternatively, cHes1 and cHes5 may inhibit phosphatases associated with MAPK activity, thereby enhancing MAPK signaling. The interactions that may take place between Notch-signaling effectors and genes downstream of, or associated with, the MAPK pathway which influence neuroprotection and glial proliferation need to be elucidated.

Our findings indicate that the stimulation of Müller glial proliferation by

FGF/MAPK requires Notch-signaling. Our results suggest an interaction between the

Notch pathway, p38 MAPK and pCREB, but not ERK1/2, Egr1 or cFos. However, pERK and associated immediate early genes cFos and Egr1 accumulate soon (<2 days) after acute damage in the retina, whereas p38 MAPK and pCREB accumulate later (>2 days) (Fischer et al. 2009b). Hence, changes in ERK1/2, cFos and Egr1 expression may have been missed because our studies focused on the effects of

DAPT at 3 days after damage. We find that DAPT blocked the FGF2-mediated accumulation of pCREB and p38 MAPK in the nuclei of Müller glia in NMDA- damaged retina. The consequence of this is a significant inhibition of proliferation of

Müller glia, indicating that Notch-signaling is required for FGF2 to stimulate glial proliferation.

Collectively, our findings indicate that MAPK-signaling alone is not sufficient to induce glial proliferation, but requires Notch activity and damage-derived signals.

Interestingly, Notch activity is required for proliferation-enhancing effects of

98 FGF/MAPK signaling, but decreases neuroprotection in the retina, unlike

FGF/MAPK signaling. Inhibition of Notch activity increased the neuroprotective capacity of Müller glia, similar to activation of FGF2-MAPK signaling (Fischer et al.

2009a). Inhibition of Notch combined with activation of FGF/MAPK-signaling did not additively protect retinal neurons against NMDA-treatment (data not shown).

Notch-signaling in the postnatal retina promotes progenitor-like characteristics

(potential to proliferate) in Müller glia, similar to FGF/MAPK-signaling. These findings suggest that the neuroprotective capacity and shifts towards or away from a

progenitor-like phenotype are independent glial activities. We propose a model for

the interaction between MAPK and Notch pathways which exert independent effects

as well as collectively impact the functions of Müller glia in a context-dependent

manner (Fig. 3.9).

Figure 3.9: Model for the influence of the Notch and MAPK pathways on retinal Müller glia. This figure draws from several studies to construct putative modes of action for insulin/IGF, FGF2 and neuronal damage on Müller glia in the chick retina (Fischer et al. 2002b; Fischer et al. 2004a; Fischer et al. 2009a; Fischer et al. 2009b) (a) Effects of insulin/IGF1 alone: Insulin/IGF1 signaling via the NIRG cells and/or microglia secondarily enhances effectors of MAPK-signaling, i.e p38 MAPK (IGF1- mediated) and cFos, Egr1 and pERK (Insulin-mediated) in Müller glia in addition to increasing neuronal cell death (purple arrows). It also modestly decreases Notch and related genes (purple dashed arrows). Low levels of Notch-signaling in Müller glia inhibits the neuron-supporting functions of Müller glia, exacerbating neuronal death (black arrows) during damage (orange arrows). In addition, low levels of Notch- signaling enhance de-differentiation and progenitor-like properties (black arrows), promoting Müller glial proliferation during damage. (b) Effects of FGF2 alone: FGF2 has Notch-dependent and Notch-independent effects. (1) Notch-independent effects – FGF/MAPK-signaling induces the accumulation of pERK1/2, p38 MAPK, pCREB, cfos and Egr1 in Müller glia, which may stimulate the Müller glia to become neuroprotective and provide support to neurons in damaged retinas (blue arrows). pERK1/2 and Egr1 promote Müller glial de-differentiation and proliferation in a Notch-independent manner (green arrows). (2) Notch-dependent effects - FGF/MAPK-signaling upregulates expression of Notch and associated genes (pink dashed arrows). Low levels of Notch-signaling in an undamaged or moderately damaged retina ‘prime’ the glia to proliferate (black arrows). FGF2-mediated Müller glial proliferation requires some retinal damage and active Notch-signaling, which promotes accumulation of p38 MAPK and pCREB (red arrows), which may make the glia more progenitor-like, inducing Müller glial proliferation during damage (orange arrow). (c) Combined effects of insulin/IGF1 and FGF2: insulin/IGF1 signaling in the NIRG cells/microglia and FGF/MAPK-signaling in the Müller glia together upregulate expression of Notch and its downstream effectors (pink arrows). Insulin/IGF1- signaling, FGF/MAPK-signaling and upregulated Notch-signaling together induce the de-differentiation and proliferation of Müller glia in the absence of damage.

100

Figure 3.9

101 Conclusions

We conclude that Notch-signaling influences the phenotype and function of

Müller glia in the mature retina. Low levels of Notch-signaling diminish the neuroprotective capacity of Müller glia, but are required to maintain the ability of

Müller glia to become progenitor-like cells. In addition, we conclude that there is cross-talk between Notch and MAPK pathways. FGF2 induces the expression of

Notch pathway genes and active Notch is required for the FGF2-mediated accumulation of p38 MAPK and pCREB in Müller glia. Further, Notch-signaling is essential for the proliferation-stimulating effects of FGF/MAPK-signaling in Müller glia in damaged retinas.

102

CHAPTER 4

Patterning of the circumferential marginal zone of progenitors

in the chicken retina2

Abstract

A circumferential marginal zone (CMZ) of retinal progenitors has been identified in most vertebrate classes, with the exception of mammals. Little is known about the formation of the CMZ during late stages of embryonic retinal histogenesis.

Thus, the purpose of this study was to characterize the formation and patterning of the

CMZ in the embryonic chicken retina. We identified progenitors by assaying for the expression of proliferating cell nuclear antigen (PCNA), N-cadherin and the nestin- related filament transitin, and newly generated cells by using BrdU-birthdating. We found that there is a gradual spatial restriction of progenitors into a discrete CMZ during late stages of embryonic development between E16 and hatching, at about

E21. In addition, we found that retinal neurons remain immature for prolonged periods of time in far peripheral regions of the retina. Early markers of neuronal differentiation (such as HuC/D, calretinin and visinin) are expressed by neurons that are found directly adjacent to the CMZ. By contrast, genes (protein kinase C,

2 Published Manuscript: Ghai, K Stanke, JJ Fischer, AJ. Brain Research (2008) 103 calbindin, red/green opsin) that are expressed with a delay (7–10 days) after terminal mitosis in the central retina are not expressed until as many as 30 days after terminal mitosis in the far peripheral retina. We conclude that the neurons that are generated by late-stage CMZ progenitors differentiate much more slowly than neurons generated during early stages of retinal development. We propose that the microenvironment within the far peripheral retina at late stages of development permits the maintenance of a zone of progenitors and slows the differentiation of neurons.

Introduction

Post-embryonic retinal growth results from on-going neurogenesis and the addition of new neurons to the peripheral edge of the retina. A zone of proliferating retinal stem cells exists at the far peripheral edge of the retinas of fish, frogs and birds

(Fischer et al. 2005; Hitchcock et al. 2004; Otteson and Hitchcock 2003; Reh and

Levine 1998). This zone of stem cells has been termed the ciliary or circumferential marginal zone (CMZ). In the normal mammalian retina, there is no evidence for the persistence of neural progenitors that are organized into a CMZ (Close et al. 2005;

Kubota et al. 2002; Moshiri and Reh 2004). The CMZ is a relatively narrow band

(<100 µm in diameter) of cells that line the periphery of the neural retina. It is located at the transition between the multilayered retina and the non-pigmented epithelium

(NPE) of the ciliary body, a pseudostratified columnar monolayer of cells that lines the vitread surface of the ciliary body. The far periphery of the retina narrows and

104 tapers down to the CMZ, which further tapers and is continuous with the NPE of the ciliary body. In the frog eye, Perron et al. (Perron et al. 1998) have demonstrated that

there is a gradient of maturity that extends through the CMZ, with retinal stem cells

residing in the most anterior region of the CMZ, restricted-fate progenitors residing in

middle, and postmitotic differentiating neurons residing in the posterior region of the

CMZ.

In comparison to the CMZ of frogs and fish, the avian CMZ adds relatively few

retinal cells to the peripheral edge of the retina, but appears to persist into adulthood

(Fischer and Reh 2000; Kubota et al. 2002; Reh and Fischer 2001). Although the

CMZ of the chick can be discretely identified with progenitor-specific markers such

as transitin (Fischer and Omar 2005), Notch1/cHairy (Fischer 2005), and the

combination of Pax6/Chx10 (Fischer and Reh 2000), progenitor- like cells may

extend into the NPE of the ciliary body and retain the ability to proliferate and

generate new neurons (Fischer and Reh 2003). Under normal conditions, the

progenitors in the CMZ are relatively quiescent, but can be stimulated to proliferate

by intraocular injections of insulin, insulin-like growth factor-I, epidermal growth

factor or sonic hedgehog, and by increasing rates of ocular growth via form

deprivation (Fischer et al. 2002a; Fischer and Reh 2000; Kubota et al. 2002; Moshiri

et al. 2005). Conversely, the proliferation of progenitors can be suppressed by

intraocular injections of glucagon or glucagon-like peptide 1 (Fischer 2005). The

quiescent state of the postnatal CMZ may, in part, result from glucagon-mediated

inhibitory input from a unique type of retinal neuron that has been termed “bullwhip

cell” (Fischer et al. 2005; Fischer et al. 2006). Furthermore, under normal conditions,

105 the progenitors in the postnatal avian CMZ do not generate photoreceptors (Fischer

and Reh 2000), or ganglion cells unless stimulated by the combination of insulin and fibroblast growth factor-2 (Fischer et al. 2002a).

Although the CMZ of the chick retina has been well described in the postnatal eye, nothing is known about the formation and patterning of the CMZ in the embryonic eye. Accordingly, the purpose of this study was to characterize the development of the CMZ in the embryonic chicken eye. We find that the CMZ is formed late during embryonic development and that there is a gradual restriction of progenitor cells to the CMZ during this time. In addition, we find that the differentiation of retinal neurons near the postnatal CMZ is much slower compared to that of neurons found in central regions of the retina. We propose that the microenvironment that permits the persistence of retinal progenitors in the CMZ also slows the differentiation of neurons in the periphery of the retina.

Materials and methods

Animals

The use of animals in these experiments was in accordance with the guidelines established by the National Institutes of Health and the Ohio State University.

Fertilized eggs and newly hatched Leghorn chickens (Gallus gallus domesticus) were

obtained from the Department of Animal Sciences at the Ohio State University.

Newly hatched chicks were kept on a cycle of 12 h light, 12 h dark (lights on at 7:00

106 am) in a stainless steel brooder at about 30 °C and received water and Purina™ chick starter ad libitum.

In ovo BrdU injections

Chick embryos were staged according to guidelines established by Hamburger and Hamilton (Hamburger and Hamilton 1992). At embryonic days 8, 10, 12 or 14, we delivered 50 µg of BrdU, diluted in 20 µl of sterile saline, into the yolks of fertilized eggs. Embryos were incubated and hatched, and retinas harvested and processed for immunolabeling procedures at postnatal day 5 (P5).

Fixation, sectioning, immunohistochemistry and photography

Tissues were fixed, sectioned and immunolabeled as described elsewhere

(Fischer et al. 1998; Fischer and Stell 1999). In short, enucleated eyes were hemisected equatorially and the gel vitreous removed from the posterior eye cup. Eye cups were fixed (4% paraformaldehyde plus 3% sucrose in 0.1 M phosphate buffer, pH 7.4, 30 min at 20 °C), washed three times in PBS (phosphate-buffered saline; 0.05

M sodium phosphate, 195 mM NaCl, pH 7.4), cryoprotected in PBS plus 30% sucrose overnight, immersed in embedding medium (OCT-compound; Tissue-Tek), and mounted onto sectioning blocks. Vertical sections, 12 µm thick, were cut consistently in the nasotemporal plane, and thaw-mounted onto SuperFrost Plus™ slides (Fisher Scientific). Sections were air-dried and stored at −20 °C until use.

Sections were thawed, ringed with rubber cement, washed three times in PBS, covered with primary antibody solution (200 µl of antiserum diluted in PBS plus 5% normal goat serum, 0.2% Triton X-100, and 0.01% NaN3), and incubated for about

107 24 h at 20 °C in a humidified chamber. The slides were washed three times in PBS, covered with secondary antibody solution, and incubated for at least 1 h at 20 °C in a humidified chamber. Finally, samples were washed three times in PBS, rubber cement removed from the slides, and coverglass mounted in 4:1 (v:v) glycerol to water. Working dilutions and sources of antibodies used in this study included: mouse anti-BrdU raised to 5-bromo-2′-deoxyuridine-BSA and used at 1:50 (G3G4;

Developmental Studies Hybridoma Band (DSHB)), rat anti- BrdU raised to BrdU and

used at 1:100 (OBT0030S; Serotec) mouse anti-proliferating cell nuclear antigen

raised to rat PCNA–protein-A fusion protein obtained from vector pC2T and used at

1:1000 (PCNA; Dako), mouse anti-transitin raised to live neural crest cells derived

from stage 11–12 quail embryos (Henion et al. 2000) and used at 1:600 (7B3A5; Dr.

P. Henion, Ohio State University), mouse anti-N-cadherin raised to N-cadherin

isolated from 11-day chick embryo heart extract and used at 1:20 (6B3; DSHB),

rabbit anti-calretinin raised to full length recombinant human calretinin at 1:1000

(7699/4; Swant Immunochemicals), mouse anti-visinin raised to visinin purified from

bovine eye and used at 1:80 (7G4; DSHB), mouse anti-HuC/D raised to a peptide

(QAQRFRLDNLLN, common to human HuC and HuC/D) conjugated to keyhole

limpet hemocyanin and used at 1:200 (16A11; Invitrogen), mouse anti-synaptophysin

raised to amino acids 205–307 of rat synaptophysin and used at 1:100 (611880; BD

Biosciences Pharmingen), mouse anti-PKC raised to PKC purified from bovine brain

and used at 1:100 (554207; BD Biosciences Pharmingen), mouse anti-calbindin

raised to calbindin D-28k purified from chicken gut and used at 1:800 (300; Swant

Immunochemicals), and rabbit anti-red/green opsin raised to recombinant human

108 red/green opsin and used at 1:400 (AB5405; Chemicon). We evaluated the specificity of primary antibodies by comparison with published examples of results and assays for specificity (Cole and Lee 1997; Fischer et al. 2002b; Fischer and Reh 2000;

Fischer and Reh 2001; Kelly et al. 1995; Kubota et al. 2004; Marusich et al. 1994).

Fluorescence within tissue sections was not caused by non-specific binding of the secondary antibody because sections labeled with secondary antibodies alone were devoid of fluorescence. Secondary antibodies included goat-anti-rabbit-

Alexa488/568, goat-anti-mouse- Alexa488/568 and goat-anti-mouse-IgM-Alexa568

(Invitrogen) diluted to 1:1000 in PBS plus 0.2% Triton X-100. Photomicrographs were taken by using a Leica DM5000B microscope equipped with epifluorescence and a 12megapixel Leica DC500 digital camera. Images were optimized for color, brightness and contrast, and double-labeled images overlaid by using Adobe

Photoshop™ 6.0.

Results

Few BrdU-labeled cells are observed within the postnatal CMZ when BrdU was applied between E8 and E14

Prada et al. utilized tritiated thymidine to determine the birthdates of cells in the chick retina and described gradients of maturation from central to peripheral and nasal to temporal retinal regions (Prada et al. 1991). This study, however, did not characterize the production of retinal neurons in far peripheral regions of the retina and the CMZ. To determine when the cells in the periphery of the retina and the CMZ

109 are generated, BrdU was injected into the yolk of time-staged embryos at E8, E10,

E12 or E14. The embryos were allowed to hatch and the eyes were fixed at postnatal

day 5. Cells with nuclei that were robustly labeled for BrdU-immunofluorescence

were likely generated by progenitors that went through terminal S-phase shortly after

the delivery of the BrdU. Cells with nuclei that were partially labeled with BrdU and contained only puncta of fluorescence were likely generated subsequent to one or more divisions, and dilutions of the BrdU, before terminal mitosis. Cells without nuclear BrdU-labeling were postmitotic prior to the delivery of BrdU or were labeled with BrdU and divided enough times to render the remaining BrdU undetectable.

In the far nasal regions of the postnatal retina, a single dose of BrdU applied at

E8 labeled cells in the GCL within 600 µm of the CMZ, numerous cells across the

ONL, and many cells in the INL (Fig. 4.1a). Most of the BrdU-labeled cells in the

INL were at least 400 µm into the neural retina, away from the postnatal CMZ (Fig.

4.1a). In the far temporal regions of the postnatal retina, by comparison, a pulse of

BrdU at E8 did not completely label any cells within 600 µm of the postnatal CMZ

(Fig. 4.1b). In far nasal regions of the retina, BrdU delivered at E10 labeled the nuclei of a few cells in the ONL (presumptive photoreceptors) within 300 µm of the CMZ, many cells in the INL within 800 µm of the CMZ, and a few cells in the GCL within

400 µm of the CMZ (Fig. 4.1c). In the far temporal regions of the postnatal retina, by comparison, BrdU delivered at E10 fully labeled the nuclei of a few cells in ONL that were at least 400 µm away from the CMZ (Fig. 4.1d). In addition, we observed some scattered cells that appeared completely labeled with BrdU in the inner and outer

INL, likely amacrine and horizontal cells, respectively (Fig. 4.1d).

110 In peripheral regions of the nasal retina, BrdU-treatment at E12 labeled a few nuclei scattered in the distal half of the INL (bipolar and/or Müller glia; Fig. 4.1e).

Most of these cells were within 300 µm of the postnatal CMZ. In peripheral regions of the temporal retina, BrdU-treatment at E12 labeled numerous nuclei scattered in the distal half of the INL (bipolar and/or Müller glia; Fig. 4.1f); most of these cells were within 2000 µm of the postnatal CMZ. In addition, we observed a high density of BrdU-labeled cells in the ONL and INL within 400 µm of the temporal CMZ (Fig.

4.1f). This “band” of BrdU-labeled cells is reminiscent of the “bands” of H3- thymidine-labeled cells observed in the birthdating studies in frog retinas of

Hollyfield over 30 years ago (Hollyfield 1968; Hollyfield 1971).

BrdU delivered at E14 failed to label photoreceptors in the ONL (Fig. 4.1g and h), indicating that the production of photoreceptors in the far peripheral regions of the retina is completed by E12 or E13. In the far periphery of the nasal retina,

BrdU delivered at E14 labeled only a few cells that were usually found within 200 µm of the CMZ (Fig. 4.1g). In the temporal retina, BrdU delivered at E14 labeled numerous cells and nearly all of these cells were found in the distal INL and a few were scattered in the proximal INL (Fig. 4.1h). Regardless of when the BrdU was delivered to the embryo, we rarely observed BrdU-labeled cells within the CMZ (Fig.

4.1).

111

Figure 4.1: BrdU-birthdating of the cells in the far periphery of the retina. Vertical sections of the nasal (a, c, e and g) and temporal (b, d, f and h) retina and CMZ were labeled with antibodies to BrdU. BrdU was delivered to embryos at E8 (a and b), E10 (c and d), E12 (e and f) or E14 (g and h), and retinas were harvested at P5. Arrow-heads indicate BrdU-positive nuclei (e–h). Asterisk indicates a “band” of proliferating cells in the ONL and INL within 400 µm of the CMZ (f). The calibration bar (50 µm) in panel f applies to panels e and f, and the bar in panel h applies to panels a–d, g and h. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; CMZ, circumferential marginal zone.

112 PCNA-expressing cells are gradually confined to the CMZ during late stages of

embryonic development

Proliferating cell nuclear antigen (PCNA) is known to be expressed by the progenitors in the postnatal chicken CMZ (Fischer and Reh 2000). The PCNA antibody labels a subunit of DNA polymerase delta that is expressed at high levels in cells in G1 and S phases of the cell cycle (Kurki et al. 1988). Thus, we assayed for the formation and patterning of the CMZ in the embryonic retina by using PCNA immunolabeling. As development proceeded, we observed a progressive restriction of

PCNA-expression within far peripheral regions of the retina. At E12, all nuclei within the far peripheral retina, presumptive CMZ and presumptive NPE were immunoreactive for PCNA (Fig. 4.2a and b). Four days later at E16 there was a significant reduction in the region of cells that were PCNA-positive (Fig. 4.2c).

Between 500 and 1000 µm away from the CMZ, PCNA-immunoreactivity was observed in fusiform nuclei near the middle of the INL (Fig. 4.2c and c′). These nuclei were likely those of differentiating Müller glia.

Within 500 µm of the CMZ, we observed numerous PCNA positive nuclei scattered across distal layers of the retina; these cells were concentrated at the peripheral edge of the retina (Fig. 4.2c and c″). In addition, we observed PCNA immunoreactivity in pigmented cells that were distal to the CMZ and the NPE of the ciliary body (Fig.

4.2c and c″). At E18, in addition to cells within the CMZ, we found PCNA- immunoreactivity in the nuclei of NPE cells and pigmented cells overlying the NPE of the pars plana (Fig. 4.2d). Further, PCNA-immunoreactivity was observed in the nuclei of presumptive Müller glia, with fusiform nuclei in the middle of the INL; this

113 immunoreactivity decreased in intensity with increasing distance from the CMZ (Fig.

4.2d). At P3 we found PCNA-positive cells that were largely confined to the CMZ

(Fig. 4.2e), consistent with previous reports (Fischer and Reh 2000).

Figure 4.2: PCNA-expressing cells are gradually confined to the CMZ during late stages of embryonic development. Vertical sections of the peripheral retina were labeled with antibodies to PCNA. Tissues were obtained from animals at E12 (a and b), E16 (c), E18 (d) and P3 (e). The calibration bar (50 µm) in panel c applies to panels a–c, the bar in panel d applies to panel d alone, and the bar in panel e applies to panel e alone. The boxed-out areas in panel c are enlarged 2-fold in the underlying panels. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone.

114 Transitin-expression is gradually confined to the CMZ during late stages of

embryonic development

The nestin-related intermediate filament transitin is expressed by neural

progenitors as well as differentiating or reactive Müller glia in the embryonic and

postnatal chicken retina (Close et al. 2005). We have shown previously that transitin-

expression is gradually down-regulated by post-mitotic Müller glia in central regions

of the retina from E12 to E18 and becomes restricted to peripheral retina sometime

shortly before hatching, at E21(Close et al. 2005). To determine exactly when

transitin expression becomes confined to a CMZ, we looked at its distribution at two

time points late in embryonic development. At E16, widespread distribution of

transitin persists in the temporal retina and ciliary body (Fig. 4.3a), with

immunoreactivity in vertically oriented processes that span all layers in the peripheral

retina (Fig. 4.3a′). Transitin-immunoreactivity was more intense in the presumptive

CMZ than in peripheral regions of the retina at E16 (Fig. 4.3a and a′). In peripheral

regions of the nasal retina, transitin is restricted to filamentous structures in the GCL

and NFL (Fig. 4.3b and b′). These structures likely were the endfeet of differentiating

Müller glia. At E16, intense

transitin-immunoreactivity persists across the CMZ and ciliary body (Fig. 4.3b and

b″). Within a short span of time, between E16 and E18, transitin-immunoreactivity

declines in peripheral regions of the temporal retina (Fig. 4.3c and c′) and is almost

completely absent from peripheral regions of the nasal retina (Fig. 4.3d and d′). In addition, there is further spatial restriction of transitin-immunoreactivity to the far periphery of the temporal retina and to the CMZ (Fig. 4.3c, c″, d and d″). At E18,

115 transitin-immunoreactivity in the nasal peripheral retina is similar to that seen at P3, persisting only in the CMZ (Fig. 4.3e and f). Consistent with patterns of PCNA- expression in the peripheral retina, we find that there is a gradual spatial restriction of transitin-positive progenitors to the far peripheral edge of the retina and into the CMZ during the last quarter of embryonic development.

116

Figure 4.3: Transitin-expression is gradually confined to the CMZ during late stages of embryonic development. Tissues were obtained from animals at E16 (a and b), E18 (c and d), and P3 (e, f). Vertical sections of the retina were labeled with antibodies to transitin. Photomicrographs were taken of central (a′ and d′), and far peripheral regions of the retina (a″ and d″) where the presumptive or well-defined circumferential marginal zone is present. The calibration bar (50 µm) in panel d applies to panels a–d, and the bar in panel f applies to panels e and f. The boxed-out areas in panels a–d are enlarged 5-fold in the underlying panels. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone.

117 N-cadherin is gradually excluded from the neural retina and becomes confined to the CMZ and NPE of the ciliary body

The calcium-dependent adhesion molecule N-cadherin is known to be

diffusely distributed on plasma membranes of retinal stem cells and progenitors in the

CMZ and ciliary body, as well as on progenitors in the zebrafish retinal primordium

(Liu et al. 2001; Liu et al. 2002; Raymond et al. 2006). In the chick embryo, Wohrn et

al. have shown a gradual down-regulation of N-cadherin expression in the central retina through development (Wohrn et al. 1998). Given that N-cadherin is expressed by progenitor cells, we wanted to determine whether it is present in the progenitor rich CMZ in late stages of retinal development.

In the E16 peripheral retina, N-cadherin-immunoreactivity is restricted to the

OPL and the distal ONL adjacent to the outer limiting membrane with very faint immunoreactivity in the IPL (Fig. 4.4a and a′). However, there is intense N-cadherin- immunoreactivity within the far periphery of the retina, including the presumptive

CMZ and the NPE of the ciliary body (Fig. 4.4a and a″). At E18, N-cadherin-

immunoreactivity is diminished in the peripheral distal retina compared to that seen at

E16 (Fig. 4.4a′, a″, b′ and b″). However, N-cadherin-immunoreactivity in the CMZ and NPE of the ciliary body continues to be high (Fig. 4.4b″). In P3 eyes, N-cadherin-

expression is absent from the far peripheral regions of the retina but continues to be

maintained in the CMZ and NPE of the ciliary body (Fig. 4.4c and f). This pattern of expression is maintained until at least P15 (data not shown). Taken together, these results indicate that spatial restriction of N-cadherin to the CMZ and NPE of the

118 ciliary body occurs gradually during late stages of embryonic development and persists into postnatal development.

To study N-cadherin expression relative to mature neurons, P3 tissue sections were labeled with antibodies to N-cadherin and calretinin, a calcium-binding protein present in the cytoplasm and neurites of horizontal, amacrine and ganglion cells (Ellis et al. 1991; Fischer et al. 1999). We observed a mutually exclusive distribution of immunoreactivities, with calretinin-positive cells found posterior and adjacent to the

CMZ, while N-cadherin positive cells remain confined to the CMZ and the NPE of the ciliary body (Fig. 4.4c–e). Further, we observed N-cadherin-positive processes that wrap around the calretinin-positive neurons adjacent to the CMZ (Fig. 4.4c-e). At

P3, there is a distinct overlap of transitin and N-cadherin immunoreactivity in the

CMZ (Fig. 4.4f–h), clearly demarcating the CMZ and the transition from neural retina to the NPE of the ciliary body. These findings suggest that transitin- and Ncadherin- positive cells are involved in patterning the CMZ at late stages of embryonic retinal development despite having differing temporal and spatial patterns of expression.

119

Figure 4.4: N-cadherin is highly expressed in the far peripheral retina, CMZ and ciliary body of both developing and postnatal retina, and gets restricted over time in far peripheral regions of the retina. Tissues were obtained from animals at E16 (a), E18 (b), and P3 (c–h). Vertical sections of the retina were labeled with antibodies to N-cadherin (a and b) and calretinin (d) or transitin (g). Photomicrographs were taken of central (a′, b′), and far peripheral regions of the retina (a″, b″, c–h) where the CMZ is forming. The calibration bar (50 µm) in panel a″ applies to panels a′ and a″, the bar in panel b applies to panels a and b, the bar in panel b″ applies to panels b′ and b″, and the bar in panel h applies to panels c–h. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone.

120 Neuronal differentiation is slowed in retinal regions that are adjacent to the CMZ

Early markers of neuronal differentiation are expressed by cells directly

adjacent to the embryonic and postnatal CMZ. To identify the onset of neuronal

differentiation we used markers that are known to be expressed soon after exit from

the cell cycle. In the chick retina, the formation of cone photoreceptors may begin as

early as E6 with the onset of visinin-expression (Bruhn and Cepko 1996). Calretinin is expressed by amacrine, horizontal and ganglion cells early during development, shortly after these cells exit the cycle and well before morphological differentiation is complete (Ellis et al. 1991). During early stages of retinal development in the chick embryo, HuC/D is expressed shortly after the terminal mitosis of amacrine and ganglion cells as these cells migrate into inner layers of the retina (Fischer and Omar

2005). Not surprisingly, we found that visinin, calretinin and HuC/D are expressed by neurons that are found directly adjacent to the CMZ at E16 and P3 (Fig. 4.5). At the periphery of the retina, however, many visinin-positive cells had the morphology of immature photoreceptors with short outer segments and without distinct axon terminals (Fig. 4.5a, d and e). In addition, we found that many bipolar cells in peripheral regions of P3 retina were weakly immunoreactive for visinin (Fig. 4.5b and c), suggesting that these cells were still in the process of differentiating. Visinin is transiently expressed at low levels by a few subtypes of bipolar and amacrine cells in central regions of the embryonic chick retina as they differentiate (unpublished observations). By making consecutive daily intraocular injections of BrdU from P0 through P4 we failed to find evidence of newly generated photoreceptors, consistent

with previous reports (Fischer and Reh 2000). This finding is in agreement with the

121 findings of our birthdating studies indicating that the generation of photoreceptors is

completed in the temporal retina by E13.

Similar to the visinin-immunoreactive photoreceptors, the calretinin- and

HuC/D-immunoreactive cells in peripheral regions of the retina had immature

morphology with oblong somata and poorly defined neurites (Fig. 4.5f–i). In the E16

retina, the region of immature calretinin-expressing cells extended about 800 µm into the retina away from the CMZ (Fig. 4.5f). In the P3 retina, by comparison, the region of immature calretinin-immunoreactive cells extended only about 200 µm away from the CMZ (compare Fig. 4.5f and g). Similar to the pattern of calretinin- immunolabeling, HuC/D was expressed by neurons that are directly adjacent to the

CMZ at E16 and P3 (Fig. 4.5h and i).

122

Figure 4.5: Early markers of neuronal differentiation are expressed by cells directly adjacent to the embryonic and postnatal CMZ. Tissues were obtained from animals at E16 (a, f and h) and P3 (b–e). Vertical sections of the retina were labeled with antibodies to visinin (red; a–e), BrdU (green; d), calretinin (green; f and g), and HuC/D (red; h and i).Tissues were obtained from chicks at E16 (a, f and h) or P3 (b–e, g and i). Arrows in panels b–d indicate visinin-positive bipolar cells. The calibration bar (50 µm) in panel a applies to panel a alone, the bar in panel d applies to panels b–d, the bar in panel e applies to panel e alone and the bar in panel i applies to panels f–i. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone.

123 To further assess whether the far periphery of the retina remains immature in

the postnatal retina we assayed for the expression of synaptophysin. Synaptophysin is

a glycoprotein present in the pre-synaptic vesicles of most neurons (Brandstatter et al.

1996). In the developing chick retina immunoreactivity for synaptophysin is first

detected at E7 in the IPL (Hering and Kroger 1996). Synaptophysin has a diffuse

distribution during early development and becomes confined to discrete lamina as

development proceeds, indicating a gradual maturation of synapses within the

plexiform layers of the retina. In the P3 eye, the CMZ is completely void of synaptophysin-immunoreactivity (Fig. 4.6a and a″). Immunoreactivity for synaptophysin begins in the IPL at about 200 µm into the retina and gradually increases with increasing distance from the CMZ (Fig. 4.6a and b). Immunoreactivity for synaptophysin in the IPL is relatively diffuse within 800 µm of the CMZ, and gradually becomes concentrated into discrete laminae (Fig. 4.6b). By comparison, synaptophysin-immunoreactivity in the OPL does not appear until about 600 µm into the retina and gradually increases in the outer lamina of the OPL with increasing distance from the CMZ (Fig. 4.6a and a′). These findings suggest that in far peripheral regions of the retina, the synaptic connections in the IPL mature more rapidly than those in the OPL.

To better characterize the gradual maturation of retinal neurons in far peripheral regions of the retina, we applied antibodies to PKC, calbindin and red/green opsin, markers that are known to be expressed by differentiating neurons during late stages of development. In the chicken retina, PKC is known to be expressed by many bipolar cells in the distal INL, as well as a few types of amacrine

124 cells in the proximal INL (Fischer et al. 1998). In the embryonic chick retina, PKC- immunoreactivity first appears in presumptive amacrine cells at about E9 and in differentiating bipolar cells at about E16 (Caminos et al. 1999). Consistent with previous reports, we did not detect immunoreactivity for PKC in central regions of the embryonic retina until E16 (Fig. 4.6d), about 8 days after these cells are generated

(Prada et al. 1991). At E16 there was no expression of PKC within 2 mm of the CMZ

(Fig. 4.6c). In temporal regions of the P3 retina, the onset of PKC expression occurred at about 300 µm from the CMZ and steadily appeared in increasing numbers of bipolar cells with increasing distance from the CMZ (Fig. 4.6f). By comparison, nasal regions of the retina contained PKC-immunoreactive bipolar cells within 50 µm of the CMZ (data not shown). In temporal regions of the P15 retina, we found PKC- immunoreactive bipolar cells within 50 µm of the CMZ (Fig. 4.6g). These findings suggest a gradual maturation of PKC-positive bipolar cells in far peripheral regions of the temporal retina.

125

Figure 4.6: Synaptophysin and protein kinase C are expressed by neurons at increasing levels with increasing distance away from the CMZ. Tissues were obtained from animals at E16 (c and d), P3 (a, b, e and f) and P15 (g). Vertical sections of the peripheral retina and CMZ were labeled with antibodies to synaptophysin (a–b) or PKC (c–g). Panels f and g; arrow-heads indicate the onset of PKC expression in P3 and P15 retinas. The calibration bar (50 µm) in panel g applies to panels a, c–g. The boxed-out areas in panel a are enlarged 2-fold in the underlying panels. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone.

126 Calbindin is known to be expressed during late stages of embryonic development in photoreceptors and bipolar cells (Ellis et al. 1991). Similarly, opsin- expression in photoreceptors begins at E15, about 10 days after terminal mitosis

(Bruhn and Cepko 1996). Consistent with these prior reports, we observed low levels of expression of calbindin and opsin in photoreceptors in central regions of the E16 chick retina (Fig. 4.7a and a′). By contrast, we failed to find immunoreactivity for calbindin or opsin in peripheral regions of E16 retina within 2- 3 mm of the CMZ

(Fig. 4.7b). At P3, we failed to find calbindin or opsin expression in far peripheral photoreceptors within approximately 300 µm of the temporal CMZ (Fig. 4.7c and c″).

In nasal P3 retina, calbindin and opsin were expressed within 50 µm of the CMZ

(data not shown). The “gap” in calbindin and opsin expression is decreased to less than 100 µm between P3 and P15 (Fig. 4.7e and e′), unlike visinin-immunoreactivity which remains in photoreceptors that are found adjacent to the CMZ from P3 through to P15 (data not shown).

127

Figure 4.7: Photoreceptors and calbindin-expressing neurons remain immature in far peripheral regions of the temporal retina. Vertical sections of the retina were labeled with antibodies to calbindin (green) and red/green opsin (red). Tissues were obtained from animals at E16 (a, b), P3 (c, d) and P15 (e). Panels c″ and e′; arrow-heads indicate the onset of calbindin expression while arrows indicate the onset of red/green opsin expression in immature photoreceptors of the peripheral retina. The calibration bar (50) µm in panel e applies to panels a–c and e and the bar in panel e′ applies to panels a′, c′, c″, d and e′. Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone.

128 Discussion

We report here that progenitors gradually become confined to the CMZ during

late stages of embryonic development, between E16 and the time of hatching, at

about E21. Proteins expressed by progenitors, such as PCNA, transitin and N-

cadherin, are present in broad domains in peripheral regions of the late embryonic

retina and these domains become reduced during the last third of retinal development.

For example, the zone of progenitors in the temporal retina is about 300 µm wide at

E16, and over the following 8 days of development, this zone of progenitors is reduced to about 50 µm in width in the postnatal (P3) retina.

We found that the nasal CMZ becomes spatially defined several days before the temporal CMZ. The progenitor markers PCNA, transitin and N-cadherin are confined to a narrow region of CMZ in the nasal retina several days before these markers become restricted to the CMZ in the temporal retina. BrdU-birthdating studies indicate that progenitors in CMZ proliferate and add new neurons to the edge of the retina through late stages of development. Numbers of BrdU-labeled cells in the nasal retina are reduced at E12 and E14, when significantly more cells are BrdU- labeled in temporal regions of the retina. These studies indicate that cells in the far peripheral regions of the nasal retina are generated before those found in the far peripheral regions of the temporal retina. We propose that the cues that maintain and pattern the CMZ in the nasal retina are reduced compared to the “CMZ cues” found in the temporal retina. Consistent with this hypothesis, we have reported previously

that numbers of proliferating progenitors in the temporal CMZ are twice as abundant

as those in the nasal CMZ (Fischer and Reh 2000).

129 We failed to observe significant numbers of BrdU-labeled cells within the postnatal

CMZ when the BrdU was applied between E8 and E14 (see Fig. 4.1). Instead, we

found BrdU-labeled cells within far peripheral regions of the retina, where

postmitotic neurons reside. The absence of BrdU-labeled cells within the CMZ likely

resulted from continued proliferation of CMZ progenitors and dilution of BrdU

applied at or before E14.

We found that retinal neurons in far peripheral regions of the postnatal retina differentiate more slowly that those generated during embryonic development in the central retina. Fig. 4.8 is a schematic diagram illustrating the regions of the far peripheral postnatal retina where different neuronal markers are expressed, indicating different stages of neuronal differentiation and maturation. In temporal regions of the retina, our BrdU-birthdating studies indicate that the last photoreceptors are added to the retina between E12 and E13 (see Fig. 4.1). The photoreceptors in the far peripheral regions of the temporal retina do not mature and differentiate to express opsin or calbindin even at P15 (see Fig. 4.7), which is about 24 days after terminal mitosis. Taken together, these findings indicate that photoreceptors at the far peripheral edge of the retina that are generated at E12 remain undifferentiated for more than 3 weeks after being generated. By comparison, cone photoreceptors in central regions of the retina are generated starting at about E5 (Prada et al. 1999), and begin to express opsins about 10 days later at E15 (Bruhn and Cepko 1996). In line with these findings, a report in the primate retina has demonstrated that photoreceptors in far peripheral regions of the retina are maintained with an immature morphology into adulthood (Fischer et al. 2001). Furthermore, we found that

130 synaptophysin- immunoreactivity in the OPL (indicating mature, functional synapses

between photoreceptors and inner retinal neurons) was not present in the peripheral regions of the retina. The gradual onset of synaptophysin-expression with increasing distance from the CMZ further indicates that the photoreceptors differentiate slowly and eventually establish synaptic connections, perhaps after the onset of opsin- and calbindin-expression. Taken together these findings suggest that the photoreceptors that are generated near the postnatal CMZ differentiate much more slowly than those generated in central regions of the retina.

131

Figure 4.8: Schematic summary of the onset of different neuronal markers and gradient of maturity that persists in peripheral postnatal retina. Each arrowhead and accompanying number indicates the region where a marker begins to be expressed by retinal cells. The timeline below the retina indicates the regions of postnatal peripheral retina that correspond to embryonic developmental stages (the gradient of maturity). The zone of the retina closest to the CMZ is the most immature (equivalent to the central retina present in E4–6 chick) while the zone of the retina furthest away from the CMZ is the most mature (equivalent to central retina present in E17–21 chick). Abbreviations: ONL, outer nuclear layer; INL, inner nuclear layer; IPL, inner plexiform layer; OPL, outer plexiform layer; GCL, ganglion cell layer; CMZ, circumferential marginal zone; PE, pigmented epithelium; CB, ciliary body; NPE, non-pigmented epithelium; PKC, protein kinase C.

132 Similar to the stunted maturation of photoreceptors in far peripheral regions of

the retina, the bipolar cells in the peripheral regions of the retina appear to

differentiate much more slowly that those in central regions of the retina. For

example, we found that the bipolar cells within 400 µm of the CMZ are generated

between E12 and E14 (see Fig. 4.1f), and these cells express PKC sometime after P3,

at least 14 days after terminal mitosis (see Fig. 4.6g). By comparison, the bipolar cells

in central regions of the retina are generated between E7 and E9 (Prada et al. 1991),

and begin to express PKC at E16 (see Fig. 4.6d), about 8 days after terminal mitosis.

Taken together these findings suggest that the microenvironment in the far

peripheral retina may be involved in serving to maintain a zone of progenitors and to

maintain retinal neurons in an immature state. Additionally, there may be different rates of cellular maturation of central versus peripheral retinal neurons because the neurons arise from different pools of progenitors in different regions of the eye. There are several possible explanations to account for the delayed maturation of neurons in the far peripheral regions of the retina: (1) there are factors in the far peripheral retina

that inhibit neuronal maturation, (2) there is a paucity of factors that stimulate the

maturation of retinal neurons, (3) the late-born neurons in the retinal periphery are

innately predisposed to mature slowly, or (4) a combination of these aforementioned

possibilities.

Secreted factors in the microenvironment at different stages of retinal

development presumably play a role in patterning the CMZ and maintaining an

immature neuronal phenotype in far peripheral regions of the retina. For example,

FGFs and FGF receptors have been shown to be involved in the central to peripheral

133 wave of differentiation of ganglion cells in the chick retina (McCabe et al. 1999).

Antagonistic interactions between FGFs secreted in the developing retina and sonic hedgehog (Shh) in the ventral forebrain may also play a role in patterning the edge of the retina, given that complex interactions between these two signaling pathways have been implicated during development and regeneration (Bertrand and Dahmane

2006; Spence et al. 2004). In addition, polypeptide hormones like insulin, insulin-like growth factor-I and glucagon have been shown to influence the proliferation of CMZ

progenitors (Fischer et al. 2002a; Fischer et al. 2005; Fischer and Reh 2000; Fischer

and Reh 2002), and may participate in the patterning of the CMZ and peripheral

retina.

Conclusions

A gradient of maturity is known to exist within the CMZ in the retinas of lower vertebrates, with the most primordial cells residing in the anterior CMZ and more mature cells residing in posterior regions of the CMZ (Hitchcock et al. 2004;

Marquardt 2003). Reports also indicate a centroperipheral gradient of maturity across the retina in teleosts (Hitchcock et al. 2004; Peng and Lam 1991; Vecino 1998). In

the chicken retina, we propose that a gradient of maturity extends through the zone of

progenitors within the CMZ and into the neural retina, similar to that seen in teleosts.

The studies presented here indicate that: (1) there is a gradual spatial

restriction of progenitors to form a CMZ during the late stages of embryonic

development, (2) the CMZ at the nasal edge of the retina forms before the CMZ at the

134 temporal edge of the retina, (3) the retinal neurons in far peripheral regions of the

nasal retina differentiate and mature before those in far peripheral regions of the

temporal retina and (4) the retinal neurons adjacent to the CMZ in far peripheral

regions of the temporal retina remain immature and differentiate far more slowly

compared to the neurons in central regions of the retina. The factors that keep retinal

neurons in an immature state remain to be identified. The microenvironment at the

periphery of the retina that promotes the persistence of a zone of retinal progenitors may also keep some types of neurons immature for extended periods of time.

135

CHAPTER 5

Serotonin released from amacrine neurons is scavenged and degraded in bipolar

neurons in the retina1

Abstract

The neurotransmitter serotonin is synthesized in the retina by one type of amacrine neuron but accumulates in bipolar neurons in many vertebrates. The mechanisms, functions and purpose underlying of serotonin in bipolar cells remain unknown. Here, we demonstrate that exogenous serotonin transiently accumulates in a distinct type of bipolar neuron. KCl-mediated depolarization causes the depletion of serotonin from amacrine neurons and, subsequently, serotonin is taken-up by bipolar neurons. The accumulation of endogenous or exogenous serotonin by bipolar neurons is blocked by selective reuptake inhibitors. Exogenous serotonin is specifically taken- up by bipolar neurons even when serotonin-synthesizing amacrine neurons are destroyed; excluding the possibility that serotonin diffuses through gap junctions from amacrine into bipolar neurons. Further, inhibition of monoamine oxidase (A) prevents the degradation of serotonin in bipolar neurons, suggesting that MAO(A) is present in these neurons. However, the vesicular monoamine transporter (VMAT2) is present only in amacrine cells suggesting that serotonin is not transported into synaptic vesicles

1 Published Manuscript: Ghai, K Zelinka, C Fischer, AJ. Journal of Neurochemistry (2009) 136 and re-used as a transmitter in the bipolar neurons. We conclude that the serotonin-

accumulating bipolar neurons perform glial functions in the retina by actively

transporting and degrading serotonin that is synthesized in neighboring amacrine cells.

Introduction

Our current understanding of the use of transmitters by neurons is based on the

axiom that a neuron synthesizes the transmitters that are released at presynaptic

terminals. Once released from a terminal, the fate of the transmitter is limited to four

possibilities; (1) the transmitter binds and activates pre- and postsynaptic receptors, (2)

the transmitter is degraded by enzymes within or surrounding the synapse, (3) the

transmitter is taken-up by the neuron that released it (re-uptake), or (4) the transmitter is

taken-up, and subsequently degraded, by glia with processes that flank the synapse.

Here we describe a fifth possibility which involves synthesis of the transmitter in one cell type that subsequently is released and actively transported into a neighboring neuron where it is degraded.

Serotonin is an amino acid-derived transmitter that is synthesized by discrete types of neurons in the central nervous system. However, a few studies have suggested that serotonin can accumulate in neurons that do not synthesize it. Lebrand and colleagues have shown that during mouse embryonic development, thalamic neurons do not synthesize serotonin but transiently take-up exogenous serotonin through high affinity transporters located on thalamocortical axons and terminals, and that glutamatergic neurons may co-release serotonin as a ‘borrowed’ transmitter (Lebrand et al. 1996). Similarly, Upton and colleagues have shown that retinal ganglion cells take-

137 up serotonin during embryonic and early postnatal development, even though these

cells do not synthesize serotonin (Upton et al. 1999). In addition, Whitworth and

colleagues have shown that serotonin transporters (SERTs) are transiently expressed in thalamocortical neurons during development (Whitworth et al. 2002). Taken together, these findings indicate that during neuronal development, serotonin is transiently transported into distinct types of neuronal cells that do not synthesize this transmitter.

Serotonin-immunoreactive bipolar neurons have been described in the retinas of several vertebrate species. In the skate retina, Schuette and Chappel have demonstrated that a subset of bipolar cells show increased serotonin-immunoreactivity in the presence of high potassium-Ringer in in vitro eye-cup preparations. Further, a SERT inhibitor blocks the uptake of exogenous serotonin by bipolar cells, but not by amacrine cells

(Schuette and Chappell 1998). The same authors provide evidence that OFF bipolar neurons acquire serotonin from large amacrine neurons in Xenopus retina (Schutte

1994). Similarly, in the chicken retina, a population of bipolar neurons is weakly

immunoreactive for serotonin during late-stages embryonic development (Rios et al.

1997). However, the roles and mechanisms of serotonin accumulation in retinal bipolar

neurons remain unknown.

In the current study, we demonstrate that a distinct type of bipolar cell in the

mature chicken retina actively transports serotonin that is injected into the eye or is

synthesized and released by amacrine cells. We determine the morphological

characteristics and immunohistochemical profile of the serotonin-accumulating bipolar

cells. We also provide evidence that serotonin is not synthesized by the bipolar neurons,

but is specifically taken-up and degraded in these cells. A distinct type of amacrine

138 neuron is the solitary source of serotonin in the retina, whereas the accumulation of

serotonin in bipolar neurons relies upon active transport.

Materials and Methods

Animals

The use of animals was in accordance with the guidelines established by the

National Institutes of Health and the Ohio State University. Newly hatched leghorn chickens (Gallus gallus domesticus) were obtained from the Department of Animal

Sciences at the Ohio State University and kept on a cycle of 12 hours light and 12 hours dark (lights on at 7:00 am). Chicks were housed in a stainless steel brooder at about

28oC and received water and Purinatm chick starter ad libitum.

Intraocular injections

Chickens were anesthetized via inhalation of 2.5% isoflurane in oxygen at a

flow rate of 1.5 l/min. Injections were made using a 25 µl Hamilton syringe and a 26-

gauge needle with a beveled, curved tip. Penetration of the needle was consistently

made through the upper eyelid into the dorsal quadrant of the vitreous chamber. In all

the experiments, 20 µl of the vehicle containing the test compound was injected into the

experimental (right) eye, and 20 µl of the vehicle alone was injected into the control

(left) eye. The vehicle was sterile saline containing bovine serum albumin, 50 µg/ml,

as the carrier. Test compounds included serotonin (5-Hydroxytryptamine; 1 µg per

dose; Sigma-Aldrich), NMDA (N-methyl-D-aspartate; 1000 nmol per dose; Sigma-

139 Aldrich), Quis (200 nmol per dose; Sigma-Aldrich), KCl (potassium chloride; 3.5M per

dose; Sigma-Aldrich), Zimelidine dihydrochloride (200 ng per dose; Sigma-Aldrich), 6-

Nitroquipazine maleate (200 ng per dose; Sigma-Aldrich), Sertraline hydrochloride

(90ng per dose; Sigma-Aldrich), Clorgyline (200 ng per dose; Sigma-Aldrich) and

Pargyline (200 ng per dose; Sigma-Aldrich). Assuming a total volume of about 1ml for a P14 eye, the initial maximum concentration of the injected compounds are 4.7µM for serotonin, 1mM for NMDA, 0.2mM for Quis, 70mM for KCl, 0.512µM for

Zimelidine dihydrochloride, 0.534µM for 6-Nitroquipazine maleate, 0.262µM for

Sertraline hydrochloride, 0.648µM for Clorgyline and 1.022µM for Pargyline.

We used 4 different injection paradigms: (1) Paradigm A – on post-hatch day 14

(P14), the right eye received a single injection of serotonin and the left eye received vehicle. Retinas were harvested at various time points 10 mins-24 hrs later. (2)

Paradigm B – on P14, the right eye received an injection of the test compound (NMDA or Quis) and the left eye received vehicle. On P14, both eyes received a single injection of serotonin. Retinas were harvested 2.5 hrs later. (3) Paradigm C – on P14, the right eye received a single injection of the test compound (KCl and/or SSRI, Clorgyline or

Pargyline) and the left eye received vehicle. Retinas were harvested 0.5 hrs or 2 hrs later. (4) Paradigm D – on P14, the right eye received a single injection of the test compound + serotonin and the left eye received serotonin alone. Retinas were harvested

2 hrs later.

140 Fixation, sectioning and immunocytochemistry

Tissues were fixed, sectioned, and sequential double-immunolabeled with

primary antibodies raised in the same species was performed as described elsewhere

(Fischer et al. 2007); (Fischer et al. 2008) Working dilutions and sources of antibodies

used in this study included; rabbit anti-serotonin polyclonal antibody (Courtesy Dr. G.

Bishop originally developed by Dr. R. Ho, The Ohio State University), mouse anti-

Lim3 used at 1:50 (67.4E12; Developmental Studies Hybridoma Bank (DSHB),

University of Iowa), mouse anti-Islet1 used at 1:50 (402.D6; DSHB), rabbit anti-Prox1

used at 1:400 (Novus), mouse anti-PKC used at 1:100 (BD Pharmingen), rat anti-

glycine used at 1:1000 (Dr. D. Pow; Univeristy of Newcastle), mouse anti-calbindin

used at 1:150 (Swant), rabbit anti-calretinin used at 1:1000 (Swant), and rabbit anti-

VMAT2 used at 1:1000 (Affinity Bio-Reagents). None of the observed labeling

appeared to be due to secondary antibody or fluorophore because sections labeled with

secondary antibodies alone were devoid of fluorescence. Secondary antibodies

included goat-anti-rabbit-Alexa488, goat-anti-mouse-Alexa488/568, goat-anti-rat-IgG-

Alexa488, goat-anti-mouse-IgG-Alexa568 (Invitrogen) diluted to 1:1000 in PBS plus

0.2% Triton X-100.

Reverse transcriptase PCR

Retinas from 3 chicks at P7 were pooled and placed in 3.0 ml of TRI Reagent®

(Sigma-Aldrich). Total RNA was isolated as described elsewhere (Fischer et al. 2008).

141 The web-based program Primer3 from the Whitehead Institute for Biomedical Research

(http://frodo.wi.mit.edu/) was used to design primers for PCR. Primer sequences were

as follows: VMAT2 Forward 5’ ACG ATG AAG AGA GAG GCA AC 3’, VMAT2

Reverse 5’ CAC CTA TGG GAT AGG ACT GG 3’. Predicted product size was 881

base pairs. PCR reactions were performed by using standard protocols and an

Eppendorf thermal cycler. 20 –mer T7 and T3 RNA polymerase initiation sites were

added to the 5’ ends of the forward and reverse primers respectively in order to

generate probes off the PCR product directly for in situ hybridization. PCR products

were run on an agarose gel to verify the predicted product sizes and purified using the

ChargeSwitch-Pro PCR clean-up kit (Invitrogen).

In situ hybridization

Standard procedures were used for in situ hybridization, as described elsewhere

(Fischer and Reh 2002); (Fischer et al. 2004). Digoxigenin-labeled riboprobes were

generated from the purified PCR product synthesized by using a kit provided by Roche

(Alameda, CA) and stored at -80°C until use. Postnatal (P14) eyes were dissected in

RNase-free Hanks’ balanced salt solution (HBSS), fixed overnight at 4°C in 4% PFA

buffered in 0.1 M dibasic sodium phosphate (pH 7.4), and embedded in OCT

compound. Cryosections were processed for in situ hybridization as described

previously (Fischer and Reh 2002); (Fischer et al. 2004).

142 Photohistogramy, Measurements, Cell Counts, and Statistical Analyses

Photomicrohistograms were obtained using a Leica DM5000B microscope equipped with epifluorescence and Leica DC500 digital camera. Confocal microscopy was done with a Zeiss LSM 510 at the Hunt-Curtis Imaging Facility at the Department of Neuroscience at the Ohio State University. Confocal stacks of images were obtained for 1 µm-thick optical sections by using a 20× objective (0.75 NA) and multi-track, narrow-pass emission filter settings to exclude the possibility of fluorescence bleeding across channels. Images were optimized for color, brightness, and contrast, and double- labeled ones were overlaid by using Adobe PhotoshopTM6.0. Cell counts were made

from at least five different animals, and means and standard deviations calculated on data sets. To avoid the possibility of region-specific differences within the retina, cell counts were consistently made from the central region of retina for each data set.

Immunofluorescence was quantified by using ImagePro 6.2. Identical illumination, microscope and camera settings were used to obtain images for quantification. Areas (1000 × 150 pixels or 290 × 43.5 µm) were sampled from 5.4 MP digital images. These areas were randomly sampled over the INL where the nuclei of the bipolar and amacrine neurons were observed. Measurements were made for regions

containing pixels with intensity values of 72 or greater (0 = black, 255 = saturated

green); a threshold that included labeling in the bipolar or amacrine neurons. The total

area was calculated for regions with pixel intensities >72. The average pixel intensity

was calculated for all pixels within threshold regions. The density sum was calculated

as the total of pixel values for all pixels within threshold regions. These calculations

143 were determined for INL regions sampled from six different retinas for each experimental condition.

Results

Serotonin is known to be synthesized and released by a subset of amacrine neurons in most vertebrate species (Jacobsen et al. 2008; Millar et al. 1988; Pourcho

1996; Rios et al. 1997). Additionally, there are a few reports of accumulation of serotonin in bipolar neurons of the chicken retina during development, with the levels declining after hatching (Rios et al. 1997). Studies in lower vertebrate species such as frog (Schuette and Chappell 1998) and skate (Schutte 1994) have suggested that serotonin is transported into bipolar neurons. However, this notion has not been unambiguously tested. Using the post-hatch chicken eye as a model system, we tested whether serotonin is merely being scavenged by bipolar neurons and degraded, or is actively taken-up by transporters and subsequently used as a transmitter by the bipolar neurons.

Exogenous serotonin is taken up by bipolar neurons

Serotonin-immunoreactivity was observed at high levels in a small population of amacrine cells with large somata found near the middle of the INL and large dendritic fields in the IPL (Fig. 5.1a), consistent with previous reports (Zhu et al. 1992);

(Schutte 1994). In addition, we observed weak serotonin-immunoreactivity in the somata of presumptive bipolar neurons located near the middle of the INL (Fig. 5.1a).

The levels of serotonin-immunoreactivity were not different in the amacrine or bipolar cells in retinas obtained under day-light or night-time illumination (data not shown).

144 For all of the following data, procedures were performed during the middle of the day.

We tested whether intraocular delivery of serotonin would increase serotonin- immunoreactivity in bipolar cells. Accordingly, 1µg of serotonin was injected into the vitreous chamber of the eye and retinas were harvested at various time-points after

injection (Fig. 5.1a-g). Serotonin-immunoreactivity was found to be higher within 10

mins of injection (Fig. 5.1a), peaked within 2.5 hrs (Fig. 5.1e), declined rapidly, and

returned to baseline levels by 4.5 hrs (Fig. 5.1f) and beyond until 24hrs (Fig. 5.1g). It is

expected that an injection of serotonin would result in transient accumulation and

degradation as is evident when we plot the area and intensity of the relative serotonin-

immunofluorescence within the distal INL (bipolar cell bodies and dendrites) across the

different timepoints (Fig. 5.1h, e). The quantitative immunofluorescence supports this

notion and verifies the utility of this method to measure relative levels of

immunolabeling. These findings indicate that exogenous serotonin is transiently taken-

up by a population of bipolar neurons and is cleared rapidly thereafter. There was an

increase in serotonin-immunoreactivity in the small population of amacrine cells too, within 2.5 hrs of serotonin delivery (Fig. 5.1b-e). In addition, neurites in the inner plexiform layer (IPL), presumably from both the amacrine and the bipolar neurons, appeared to transiently accumulate serotonin following an intraocular injection (Fig.

5.1a-g).

145

Figure 5.1: Exogenous serotonin transiently accumulates in bipolar neurons. Intraocular injections of vehicle or 1µg of serotonin were made at P14, and the retinas were harvested at various timepoints between 10 mins to 24 hours after the injection. Vertical sections of the retina were labeled with antibodies to serotonin (a-g). As described in the methods, Image Pro 6.2 was used to obtain measurements of total area for pixel intensities >72 (0 = black, 255 = saturated) and the intensity sum. Serotonin- immunofluorescence was measured in approximately 12600 µm2 of the distal INL containing the somata and dendrites of bipolar neurons. The somata of amacrine cells were excluded from the measurements. Histograms indicate the total area of labeling (h) and the total intensity of labeling (i) per timepoint measured. Arrows indicate bipolar neurons and arrow-heads indicate amacrine neurons. The calibration bar (50µm) in panel g applies to panels a-g. Abbreviations : ONL - outer nuclear layer, INL - inner nuclear layer, IPL - inner plexiform layer.

146 Immunofluorescence was not measured in the terminals of bipolar neurons in the

IPL because it was impossible to distinguish the axon terminals from the processes of amacrine neurons. Although it is likely that the uptake of serotonin is prevalent at the terminals of bipolar neurons, elevated levels of serotonin-immunofluorescence in the somata and dendrites of bipolar neurons occurred, presumably, following uptake at the axon terminals.

Serotonin-accumulating bipolar (SAB) cells have a distinct morphology and immunohistochemical profile

In order to characterize the type of bipolar cell that accumulates serotonin, we injected eyes with serotonin, harvested the retinas 2 hours later and labeled the retinal sections with antibodies to well-known markers of bipolar neurons - Lim3, Prox1,

Islet1, PKC, calretinin, calbindin, and glycine, along with serotonin. Members of the

Lim-domain family of transcriptional factors are known to be constitutively expressed by bipolar neurons in the retina (Edqvist et al. 2006; Fischer et al. 2008). We observed that a subset of Lim3-positive bipolar neurons accumulated serotonin; these cells had somata located near the middle of the INL, vitread to most other types of bipolar cells

(Fig. 5.2a-d). All SAB cells (n=121) counted were positive for Lim3. The SAB cells were weakly positive for Prox1 (Fig. 5.2e-h). None of the SAB cells were positive for glycine, Islet1, calretinin, calbindin or PKC (Fig. 5.2i-n). We compared the morphology of SAB cells to the bipolar cell-types described by Cajal by using Golgi- impregnation (Fig 5.2o). The SAB cells have somata that were located in the center of the INL and bistratified axon terminals in both the distal and proximal IPL (Fig. 5.2o).

147 Their dendritic terminals branched out and established endings adjacent to calbindin-

positive photoreceptor pedicles in the distal OPL (Fig. 5.2l and n). The SAB cells do

not form a Landolt’s club (the enlarged apical dendrite of a bipolar cell) unlike most

other types of bipolar neurons. However, we cannot exclude the possibility that

serotonin fails to accumulate within the Landolt’s club. The SAB cells are distinctly

different from the types of bipolar cells described by Cajal in the avian retina (Fig.

5.2o).

SAB cells take-up exogenous serotonin when the endogenous source is ablated

It is possible that exogenous serotonin accumulates in bipolar neurons because

of active transport and diffusion through gap junctions from the serotonergic amacrine

cells to the bipolar cells. NMDA and Quis are known to destroy the serotonergic

amacrine cells, in addition to many other types of amacrine neurons, whereas bipolar

cells escape excitotoxic cell death. Consistent with a previous report (Fischer et al.

1998), we found that NMDA and Quis destroyed the vast majority of serotonin- producing amacrine neurons within 7 days of the treatment (Fig. 5.3b, c and f).

Occasionally, we observed a residual serotonin-positive amacrine neuron with abnormal dendrites remaining in the damaged IPL (not shown). Bipolar neurons in damaged retinas had little or no serotonin-immunoreactivity (not shown).

148

Figure 5.2: Characterization of SAB cells. Immunoreactivity for Lim3 and Prox1 is present in the nuclei of SAB cells whereas Islet1, PKC, calretinin, calbindin and glycine is in different subsets of bipolar cells. An intraocular injection of serotonin was made at P14, and the retina was harvested 2 hours after the injection. Vertical sections of the retina were labeled with antibodies to serotonin (green; a, b, d, e, f, h, i, j, k, l, m, n) and Lim3 (red; a, c and d), Prox1 (red; e, g and h), glycine (red; i), Islet1 (red; j), calretinin (red; k), calbindin (red; l and n) and PKC (red; m). The box in panel a has been enlarged in panels b-d, and the channels have been separated to show to different labels used. Similarly, the box in panel e has been enlarged in panels f-h. Panel n is a threefold enlargement of the box in panel l. The typical morphology of the two SAB cells (indicated by asterisks) is shown in panel o along with other types of bipolar neurons as described by Cajal. These SAB cells are a color inversion grayscale conversion of the two adjacent SAB cells indicated by asterisks in panel l. Arrows indicate serotonergic bipolar neurons that express Lim3 (b-d) or Prox1 (f-h). Blue arrows indicate serotonergic bipolar neurons that do not express glycine, Islet1, PKC, calbindin or calretinin (i-m). Arrowheads indicate non-serotonergic bipolar neurons immunoreactive for Lim3, Prox1, Islet1, PKC, calretinin, calbindin or glycine. Double- arrows indicate serotonergic amacrine neurons (b, d, i, j). The calibration bar (50µm) in panel k applies to panels a, e, i-k. The calibration bar (50µm) in panel m applies to panels b-d, f-h, l and m. Abbreviations : ONL – outer nuclear layer, INL – inner nuclear layer, IPL – inner plexiform layer, OLM – outer limiting membrane, ON and OFF – On and Off layers of the IPL, OPL – outer plexiform layer, GCL – ganglion cell layer

149

Figure 5.2

150 Thus, either the serotonin-immunoreactive bipolar neurons were destroyed by the excitotoxins, or these bipolar neurons survived but were no longer detected because their source of serotonin, the amacrine neurons, was ablated. To test these possibilities, we injected serotonin into NMDA and Quis-treated eyes. Intense serotonin- immunoreactivity was seen in bipolar neurons in NMDA and Quis-treated retinas (Fig.

5.3b and c), whereas few amacrine cells were detected (Fig. 5.3f). In the absence of serotonin-producing amacrine neurons, exogenous serotonin was taken-up by the bipolar neurons, suggesting that these cells express serotonin transporters and survive

NMDA and Quis treatments. With the majority of serotonergic amacrine cells destroyed, we were able to distinguish the axon terminals of the bipolar neurons that stratify in both distal and proximal strata of the residual IPL.

151

Figure 5.3: Bipolar neurons accumulate exogenous serotonin even when amacrine neurons are destroyed. Intraocular injections of vehicle (a), Quis (b) or NMDA (c) were made at P14. 7 days post treatment (dpt), one µg of serotonin was injected into both eyes and the retinas were harvested 2.5 hours later. Vertical sections of the retina were labeled with antibodies to serotonin. As described in the methods, Image Pro 6.2 was used to obtain measurements of total area for pixel intensities >72 (0 = black, 255 = saturated), average pixel intensity and the density sum. Serotonin- immunofluorescence was measured in approximately 12600 µm2 of the distal INL containing the somata of bipolar neurons. The somata of amacrine cells were excluded from the measurements. Histograms indicate the mean intensity of labeling (d), total area of labeling (e), mean numbers of amacrine neurons (f), mean numbers of bipolar neurons per 4800µm2 (g). Arrowheads indicate serotonergic amacrine cells and arrows indicate SAB cells. Significance of difference (*p < 0.05, **p < 0.01) was determined using a Student’s t-test. The calibration bar (50µm) in panel c applies to panels a - c. Abbreviations: INL - inner nuclear layer, IPL - inner plexiform layer.

152 Depolarization induces the accumulation of serotonin in bipolar neurons with a loss

of serotonin from amacrine neurons

KCl is known to cause membrane depolarization and neurotransmitter release

from neurons. We hypothesized that neurons would release serotonin with KCl- mediated depolarization, hence decreasing serotonin-immunoreactivity. To test this

hypothesis, we injected saline or KCl into eyes and harvested the retinas 2 hours later.

Although we observed a significant decrease in serotonin-immunofluorescence in

amacrine neurons, their levels were significantly enhanced in bipolar neurons (Fig.

5.4a-b). This finding suggests that KCl-mediated depolarization stimulates amacrine

neurons to release serotonin, thereby depleting stores and reducing

immunofluorescence. It is also possible that KCl-mediated depolarization reverses the

direction of SERT in amacrine cells to deplete stores of serotonin. Subsequent to KCl- stimulated release from amacrine cells, serotonin is taken-up by nearby bipolar neurons.

Alternatively, KCl causes an increase in the rate of transport of serotonin, or depolarization of inhibitory amacrine cells may feed back to inhibit the depolarization

of bipolar neurons, thereby preventing the release of serotonin from bipolar neurons.

The percentage change in the intensity of serotonin-immunofluorescence is shown in

Fig 5.4c.

153

Figure 5.4: Depolarization stimulates depletion from amacrine neurons and accumulation of serotonin by bipolar neurons. Uptake of exogenous serotonin uptake by bipolar cells is blocked by a selective serotonin transport inhibitor (zimelidine). KCl-induced serotonin uptake by bipolar neurons is blocked by zimelidine. Intraocular injections of saline (a) or 20µl of 3.5M KCl (b) were made at P14 and the retinas were harvested 2 hours later. Vertical sections of the retina were labeled with antibodies to serotonin. As described in the methods, Image Pro 6.2 was used to obtain measurements of total area for pixel intensities >72 (0 = black, 255 = saturated), average pixel intensity and the density sum. Immunofluorescence in 12600µm2 of the distal INL containing the somata of bipolar neurons was measured. Amacrine cell somata were excluded from the measurements. The histogram in panel c indicates the percentage change in mean serotonin-fluorescence intensity from the baseline (saline) to the treatment (KCl) condition after 2 hours. Intraocular injections of 1µg of serotonin (d, f, h) or 1µg of serotonin and 25µM zimelidine (e, g) or 1µg of serotonin and 9µM sertraline were made at P14 and the retinas were harvested 2 hours later. The small numbers and red outlines in panels f and g indicate the areas designed by ImagePro 6.2 for each object that met the threshold criteria. Histograms in panels j- m indicate the mean area (j, l) and intensity (k, m) of the serotonin labeling respectively. Intraocular injections of 20 µl of saline (n), 3.5M KCl (o) or 3.5M KCl and 25µM zimelidine (p) were made at P14 and the retinas were harvested 30 min later. The histogram in panel q illustrates the percentage change in mean fluorescence intensity of the serotonin labeling from the saline-treated to the KCl-treated condition. The histogram in panel r indicates the percentage change in mean fluorescence intensity of the serotonin labeling from the KCl-treated condition to the KCl + zimelidine-treated condition. Images in panels a and b were obtained using confocal microscopy and images in panels d-i and n-p were obtained using wide-field epifluorescence microscopy. Significance of difference (**p < 0.01) was determined using a Student’s t-test. Arrowheads indicate serotonergic amacrine cells and arrows indicate SAB cells. The calibration bar (50µm) in panel e applies to panels d and e, that in panel h applies to panels h and i, and the one in panel p applies to panels a, b, n, o and p. Abbreviations: INL - inner nuclear layer, IPL - inner plexiform layer, GCL - ganglion cell layer.

154

Figure 5.4

155 Exogenous uptake of serotonin by bipolar neurons is blocked by selective SERT

inhibitors

Accumulation of serotonin in bipolar neurons may occur through serotonin

transporters. Accordingly, a specific SERT reuptake inhibitor, zimelidine, was injected

with serotonin into the eye. Zimelidine significantly reduced the serotonin-

immunofluorescence within bipolar neurons (Fig. 5.4d-g, j, k), while causing no change

in serotonin-immunofluorescence in amacrine neurons (Fig. 5.4d-g, j, k). Similar

results were seen with two additional SERT inhibitors, sertraline (Fig. 5.4h, i, l, m) and

nitroquipazine maleate (data not shown). Since KCl induced an accumulation of

serotonin in bipolar neurons, we sought to determine whether this accumulation was

due to SERT-mediated uptake of the serotonin. KCl was injected with or without zimelidine and retinas were harvested 30 minutes later. Serotonin-immunofluorescence was significantly diminished in both amacrine and bipolar neurons treated with KCl and zimelidine (Fig. 5.4n-r). Our results are consistent with the hypothesis that the

SAB cells normally take-up serotonin that is released from amacrine neurons.

Degradation of serotonin in bipolar neurons is prevented by an MAO(A) inhibitor

Monoamine oxidases (MAO) catalyze the oxidative deamination of

monoamines to inactivate and clear transmitters. MAO(A) specifically oxidizes

serotonin, norepinephrine and epinephrine while MAO(B) oxidizes phenethylamine.

We tested whether MAO inhibitors influence the endogenous levels of serotonin-

immunofluorescence in retinal cells. Eyes were injected with MAO(A) inhibitor

(clorgyline) or MAO(B) inhibitor (pargyline) and retinas were harvested 2 hours later.

156 We observed that clorgyline (Fig. 5.5a-b), but not pargyline (Fig. 5.5c-d), increased the

serotonin-immunofluorescence in bipolar neurons and amacrine neurons, suggesting that degradation via MAO(A) was inhibited. Further, neurites in the IPL were more

intensely immunoreactive for serotonin in the presence of clorgyline (Fig. 5.5b). The total area and intensity of serotonin-immunofluorescence in the somata of SAB cells in the distal INL was significantly increased by the MAO(A) inhibitor, but not with the

MAO(B) inhibitor (Fig. 5.5e-h). This finding suggests that the enzymes for serotonin degradation are present in the SAB cells.

157

Figure 5.5: Degradation of serotonin in bipolar neurons is prevented by an MAO(A) inhibitor (clorgyline) but not by an MAO(B) inhibitor (pargyline). Intraocular injections of saline (a, c), 200 ng clorgyline (b) or 200 ng pargyline (d) were made at P14 and the retinas were harvested 2 hours later. Vertical sections of the retina were labeled with antibodies to serotonin. As described in the methods, Image Pro 6.2 was used to obtain measurements of total area for pixel intensities >72 (0 = black, 255 = saturated green), average pixel intensity and the density sum. Serotonin- immunofluorescence was measured in 12600µm2 of the distal INL containing the somata of bipolar neurons. Amacrine cell somata were excluded from the measurements. Histograms in panel e illustrate the total area and intensity of the serotonin labeling per treatment condition. Significance of difference (**p < 0.01) was determined using Student’s t-test. Arrowheads indicate serotonergic amacrine cells and arrows indicate SAB cells. The calibration bar (50µm) in panel d applies to panels a-d. Abbreviations : INL - inner nuclear layer, IPL - inner plexiform layer. 158 Expression of vesicular monoamine transporter 2 (VMAT2) in the retina

To determine whether SAB cells utilize serotonin as a neurotransmitter, we

labeled retinas for VMAT2, a protein that is required to load synaptic vesicles with

serotonin. VMAT2 has been shown to be present in the IPL processes of the rat retina

(Ostergaard et al. 2007). We found that VMAT2-immunoreactivity is present in the

cell bodies and proximal neurites of serotonergic amacrine cells. None of the somata of the SAB cells were immunoreactive for VMAT2, indicating that these cells do not utilize serotonin as a neurotransmitter. In addition, VMAT2-immunolabeling was observed in the distal and proximal strata of the IPL (Fig. 5.6b, c, e, f, h). Most of the

VMAT2-positive neurites in the IPL were concentrated in a distal stratum of the IPL

(Fig. 5.6b and e). Additionally, VMAT2 is present in neurites in the middle and

proximal strata of the IPL. Serotonin-positive processes co-localized with VMAT2 in

both distal and proximal strata of the IPL (Fig. 5.6a, c, d and f). Further, VMAT2-

positive / serotonin-negative processes were present in a distinct layer that sits directly

above the distal serotonergic processes in the IPL and in the centre of the IPL (Fig.

5.6f). The serotonin-negative, VMAT2-positive neurites in the IPL were those of TH-

positive dopaminergic amacrine cells (Fig. 5.6g-i). Consistent with the findings of

immunolabeling studies, VMAT2 mRNA was shown to be expressed in the cell bodies

of amacrine cells but not bipolar cells in the INL (Fig. 5.6j and k).

159

Figure 5.6: Immunoreactivity and in situ hybridization indicate expression for vesicular monoamine transporter 2 (VMAT2) in amacrine cells in the retina. An intraocular injection of serotonin was made at P14, and the retina was harvested 2 hours after the injection. Vertical sections of the retina were labeled with antibodies to serotonin (green; a, c, d and f), TH (green; g and h) and VMAT2 (red; b, c, e, f, h and i). The box in panel c has been enlarged in panels d-f, and the channels have been separated to show to different labels used. In panels a-f, arrows indicate serotonergic amacrine neurons that accumulate serotonin (a, c, d and f) and express VMAT2 (b, c, e and f), and arrowheads indicate SAB cells that are not immunoreactive for VMAT2 (a, c, d and f). In panels g-i, arrows indicate VMAT2-positive amacrine cells that express TH and arrowheads indicate VMAT2-positive amacrine cells that are not immunoreactive for TH. Vertical sections of P14 retina were hybridized with riboprobe to VMAT2 (j and k). Panel k is a threefold enlargement of the box in panel j. The calibration bar (50µm) in panel c applies to panels a-c, the one in panel f applies to panels d-f, that in panel i applies to panels g-i and the one in panel j applies to panel j alone. Abbreviations : OPL – outer plexiform layer, INL – inner nuclear layer, IPL – inner plexiform layer

160

Figure 5.6

161 Discussion

We report here that bipolar cells in the retina are capable of accumulating serotonin within neurites and cell bodies, even though they do not synthesize this neurotransmitter. Serotonin is likely to be synthesized by amacrine cells in the inner nuclear layer. Studies have shown that tryptophan hydroxylase 1 (TPH1), the serotonin biosynthetic enzyme, is expressed in the chicken retina (Iuvone et al. 1999). TPH1 is expressed predominantly in the photoreceptors of the outer nuclear layer and at low levels in presumptive amacrine cells of the inner nuclear and ganglion cell layers

(Liang et al. 2004). Further, a population of amacrine cells, but not bipolar cells, has been reported to be immunoreactive for anti- phenylalanine hydroxylase using an antibody that also recognizes tryptophan hydroxylase (Cotton et al. 1988). Thus, serotonin has been proposed to be a neurotransmitter in amacrine cells.

Our findings are consistent with the notion that some types of bipolar cells accumulate serotonin in different vertebrate retinas (Schlemermeyer and Chappell

1996). However, the role of serotonin accumulation in the SAB cells remains unclear.

It is possible that one function of the SAB neurons is to degrade transmitter that is released from another type of neuron - a function that is normally ascribed to glial cells. Our data indicate that the SAB cells utilize SERT to take-up serotonin that is normally released from a distinct type of amacrine cell. We found that an MAO(A) inhibitor increased levels of serotonin in both amacrine and SAB cells during daylight conditions. This finding indicates that normally some of the serotonin that is transported into the SAB cells is degraded via MAO(A) and that the serotonin

162 originates from amacrine cells. However, we cannot exclude the possibility that the

MAO(A) inhibitor blocks serotonin degradation in amacrine cells or Müller glial cells

thereby enhancing serotonin levels in bipolar cells. Although we can detect the mRNA

from whole retinal extracts, it is unclear which cell types in the retina express MAO(A).

Subsequent to uptake, the SAB cells degrade some of the serotonin, performing

a function that is expected of glia (such as astrocytes or Müller glia) but not of

neurons. It is widely believed that monoamine transporters are expressed selectively on

presynaptic cells that produce the specific monoamine (Gainetdinov and Caron 2003).

The purpose of SERT expression on the pre-synaptic terminal is to remove the transmitter that escapes the cleft and allow for re-cycling of the transmitter for subsequent synaptic transmission. However, our data suggest that serotonin transporters can be expressed by neurons that do not re-use the transmitter for synaptic release. To the best of our knowledge, there are currently no reports of neurons in the mature nervous system that actively transport and degrade serotonin that is released from a nearby orthologous neuronal cell type.

A small population of amacrine cells is the only type of neuron in the retina that uses serotonin as a synaptic transmitter. Although, serotonin can be detected in both amacrine and bipolar neurons in the retina (current findings and references), VMAT2 is only detected in the serotonergic and dopaminergic amacrine cells. Although the bipolar cells accumulate serotonin, these cells do not express the transporter to load synaptic vesicles. These findings exclude the possibility that SAB cells utilize serotonin as a ‘borrowed’ neurotransmitter.

163 It is unlikely that the bipolar cells use VMAT1 to load synaptic vesicles with

serotonin. VMAT1 has not been identified in Gallus gallus and has only been shown to

be present in non-neuronal cells in other species including rat pinealocytes (Hayashi et

al. 1999), adrenal medulla and intestinal endocrine cells (Hansson et al. 1998; Weihe et

al. 1994), human neuroendocrine tissue (Erickson et al. 1996), whereas VMAT2 is

known to be present in neurons in the brain (Schutz et al. 1998) including the retina

(Ostergaard et al. 2007; Upton et al. 1999). Differential studies on the expression of

VMAT1 and VMAT2 have shown that VMAT2 is predominantly expressed in the

central and peripheral nervous system whereas VMAT1 is only expressed outside the

nervous system (Peter et al. 1995). Given the evidence in the literature, it can be assumed that (a) there may be no orthologue for VMAT1 in chicken and / or (b)

VMAT1 is not present in the neurons of the chicken retina.

It is unlikely that the SAB cells convert serotonin to melatonin using the enzyme N-acetyltransferase (NAT). NAT is known to be expressed in photoreceptors

(Niki et al. 1998; Tosini et al. 2006). However its presence has not been documented in

bipolar neurons. If the SAB cells do not express NAT, then it may be possible that

some of the serotonin that the SAB cells scavenge from the amacrine cells is

transported to the outer retina and is, subsequently, somehow transferred to photoreceptors that convert the serotonin into melatonin. The sources and the

mechanisms by which serotonin reaches photoreceptors remain uncertain. It is well

established that retinal melatonin is synthesized in a circadian manner by

photoreceptors (Iuvone et al. 2005; Lundmark et al. 2007; Tosini et al. 2008). Further,

164 reports have suggested that the expression and activity of tryptophan hydroxylase in the

retina are regulated by circadian rhythms (Chong et al. 1998; Green and Besharse 1994;

Liang et al. 2004; Thomas and Iuvone 1991) indicating that diurnal changes in metabolism influence serotonin synthesis. We assessed whether the accumulation of serotonin in the SAB cells was also dependent upon photoperiod but found no differences, indicating that the accumulation of serotonin in bipolar neurons may not be modulated by ambient light levels (data not shown). It remains possible that there are subtle diurnal changes in serotonin accumulation that were not detected by our methods.

The SAB cells have a unique morphology among previously described bipolar

cells. Ramon y Cajal described seven morphologically distinct types of bipolar cells in

the chicken retina (Cajal 1972). The SAB cells that we currently describe appear to be

different from those described by Cajal with regard to the pattern and distribution of

terminal arbors in the OPL and IPL. Further, the SAB cells do not form a Landolt's

club, unlike 6 of the 7 types of bipolar cells described by Cajal. It is possible there may

be bias against labeling the SAB cells via silver chromate impregnation. For example,

rare cell types like bullwhip cells have gone unnoticed by this technique and were

described only recently (Fischer et al. 2005; Fischer et al. 2006). Likewise, here we provide the first detailed description of the morphology and immunohistochemical profile of the SAB cells; loading with exogenous serotonin has allowed this. However, we cannot exclude the possibility that serotonin is excluded from the Landolt's club,

165 which might otherwise indicate that the SAB could, indeed, be included among the bipolar types that were described by Cajal.

Conclusions

The serotonin-accumulating bipolar (SAB) cells represent a distinct type neuron that has not been well characterized previously. Under normal conditions, the SAB cells utilize SERT to accumulate serotonin that is synthesized and released from a minor population of amacrine cells in the retina. Some of the serotonin that is transported into the SAB cells is degraded via MAO(A). We conclude that the SAB cells perform functions normally ascribed to glial cells.

166

CHAPTER 6

Summary and Future Directions

Summary of findings

Developmental and functional studies of the nervous system are often challenging due to the complex structure and organization in the brain and spinal cord.

The retina is an excellent model system for studying the mechanisms of development, regeneration and cellular function within the central nervous system, mainly for 3 reasons – (1) its easy accessibility, (2) its relatively simple organization (compared to the brain), and (3) because this tissue is not essential for viability, which permits the use of multiple approaches that would otherwise compromise the existence of the organism.

Furthermore, given that several diseases of the eye involve cellular and functional abnormalities of the retina, it becomes relevant to investigate mechanisms in this tissue that jeopardize vision.

This dissertation addresses aspects of retinal development, regeneration, structure as well as cellular responses to damage and growth factors. In Chapter 2, we see that blocking Notch-signaling in embryonic cell cultures enhances premature neuronal differentiation. Interestingly, cell-fate specification varies depending on the mode of inhibition and the specific bHLH effector being blocked. Our findings indicate that bHLH factors downstream of Notch-signaling do not exhibit similar functions; cHes1 inhibits neurogenesis whereas cHes5 enhances neurogenesis. However the

167 overall effect of blocking Notch-signaling is to enhance neurogenesis as we see when

DAPT is used in embryonic cell cultures.

In Chapter 2, we also find that Notch regulates aspects of Müller-glial derived

de-differentiation, proliferation and regeneration. We report that components of the

Notch-signaling pathway are increased within 2 days after retinal damage, and that

blocking the Notch-pathway with DAPT enhances proliferation of Müller-glial derived

cells as well as neurogenesis in central regions of the retina and gliogenesis in temporal

regions. The stage at which the Notch-pathway is blocked after damage is important;

from our observations, we have determined that the earliest stage at which the Notch-

pathway affects regeneration is when glial-derived cells are in the process of de-

differentiating and proliferating. At this stage, blocking Notch enhances neurogenesis as well as gliogenesis. However, as Hayes et al have reported, blocking Notch when glial-derived progenitor cells have already de-differentiated enhances neurogenesis to a much greater extent (Hayes et al. 2007).

In Chapter 3, we determine the role of Notch in regulating Müller glial properties in an undamaged postnatal retina. Our most striking finding is that low levels of Notch-signaling diminish the neuroprotective capacity of Müller glia. Furthermore, low levels of Notch-signaling are required to maintain the ability of Müller glia to become progenitor-like cells. Interestingly, we see that there is cross-talk between

Notch and MAPK pathways. FGF2 induces the expression of Notch-pathway genes and active Notch is required for the FGF2-mediated accumulation of p38 MAPK and pCREB in Müller glia. Further, Notch-signaling is essential for the proliferation- stimulating effects of FGF/MAPK-signaling in Müller glia in damaged retinas.

168 In all our studies, we observe that glia are more responsive in peripheral regions of the retina compared to those in central regions. These results are in agreement with findings that a centro-peripheral gradient of maturity exists across the retina(Hitchcock et al. 2004; Peng and Lam 1991; Prada et al. 1991; Vecino 1998). In Chapter 4, we demonstrate that neurons in far peripheral regions of the retina remain immature and differentiate far more slowly compared to the neurons in central regions of the retina.

These neurons are adjacent to the CMZ, the zone of stem cells at the outer edge of the retina in adult chickens. Our findings suggest that the microenvironment at the periphery of the retina that promotes the persistence of a zone of retinal progenitors may also keep some types of cells in that region immature for extended periods of time.

Most of the studies described in this dissertation focus on the phenotype and function of Müller glia in the retina. In Chapter 5, we describe and discuss our interesting findings about some glia-like characteristics of a non-glial cell type in the retina – the bipolar cell. Bipolar cells are interneurons with cell bodies located in the

INL, much like Müller glia. In our studies, we have identified a unique sub-type of these bipolar cells that transiently accumulates serotonin in their nuclei and cell bodies and degrades this neurotransmitter. To our knowledge, this is the first example of a neuron assuming a glial function in the retina.

169 Future directions for studies on Müller glia

Introduction

For postmitotic Müller glia to be harnessed as a local source of stem cells within

the eye, it is essential that we understand the factors that drive the ability of these cells to take on an immature phenotype. In response to retinal damage or growth factors,

Müller glia can proliferate, de-differentiate into progenitor-like cells and produce

neurons (Fischer and Reh 2001). On the other hand, the same factors can contribute to

an alternate Müller cell phenotype – reactive gliosis, which can lead to scar formation and compromise neuronal survival (Bringmann et al. 2006). Müller glia have been implicated in the pathogenesis of several sight-threatening retinal diseases such as X- linked juvenile retinoschisis (Harris and Yeung 1976), cystoid macular edema (Loeffler et al. 1992), Müller cell sheen dystrophy (Fisher 1994; Kellner et al. 1998) and retinal detachment (Erickson et al. 1987; Fisher 1994; Lewis et al. 1994). Further, the ability of Müller glia to re-enter the cell cycle makes them attractive candidates in mediating the pathology of disorders involving cell division, such as retinal gliosis (Nork et al.

1986), retinal tumors (Craft et al. 1985) and proliferative diabetic retinopathy (Nork et al. 1987). Researching the processes that modulate glial cell plasticity would yield important insights into controlling the detrimental effects of gliosis while at the same time, promoting the regenerative ability of glia to replace lost neurons.

170 Notch and the effects of other signaling pathways

The mechanisms that contribute to the plastic nature of the Müller glial phenotype are just beginning to be understood. From our studies, it is apparent that the

Notch-signaling pathway regulates the ability of glia in a mature retina to be protective, proliferative or regenerative. However, there are clearly more factors that control or restrict the potential of these cells to assume stem cell-like characteristics. For example, the Shh pathway is known to be an important modulator of retina regeneration (Spence et al. 2004). Interactions between the FGF/MAPK pathway and the Sonic Hedgehog pathway have been reported in the process of stimulating retinal regeneration (Spence et al. 2007). Ongoing studies from our lab indicate that active Shh signaling make the glia less neuroprotective via activation of Patched and Glis (Scott and Fischer 2007). It would be interesting to see whether Notch and Shh pathways also interact with each other reciprocally to regulate Müller glial de-differentiation and proliferation. Wnt- signaling may also regulate glial-derived regeneration. Osakada et al have shown that activation of the Wnt pathway in rodent retinas stimulates proliferation of Müller glia and promotes their de-differentiation into retinal progenitors (Osakada et al. 2007).

Interactions between Wnt and the downstream effector of Notch, Hairy1, have been shown to be present in the chick retina (Kubo and Nakagawa 2009). Thus, future studies focusing on which developmental pathways get activated in Müller glia in the presence of secreted factors (such as IGF, FGF and CNTF) and researching these interactions would yield useful information on how to manipulate Müller glial plasticity.

171 From our studies, it appears that Notch and MAPK pathways have independent effects on Müller glial properties as well as combined effects: i.e., MAPK-induced

Müller glial proliferation requires active Notch-signaling. However, at low levels in the postnatal retina, Notch-signaling maintains a pool of Müller glia with progenitor-like properties. This low level of Notch inhibits neuroprotective properties of Müller glia and promotes neuronal cell death under certain conditions (stress or damage to the neurons).

Whether FGF/MAPK acts only through Notch to enhance proliferation remains unanswered. It is possible that active Notch-signaling acts as a gateway to allow MAPK effectors to stimulate proliferation or relieve inhibition on these factors. To test this possibility, we would have to upregulate Notch in Müller glia with exogenous insulin/IGF1 and FGF2 (or by overexpressing Notch ICD) and then block the downstream effects of insulin/FGF2 using MAPK inhibitors. If our hypothesis is correct, we do not expect to see any proliferation of glial-derived progenitors when

MAPK-signaling is blocked, even though Notch-signaling is active. However, if we do see proliferation in the retina, we would conclude that Notch-signaling drives the proliferation of de-differentiated Müller glia, independent of MAPK.

Notch ligands – where are they expressed?

An important issue that needs to be addressed in our studies is determining which cells express Notch ligands and effectors in the retina. Using in situ hybridization, we clearly show that cNotch1 is expressed by most, if not all, Müller glia in chicken retina. However, we have used a range of antibodies and RNA probes to

172 cDelta1, cHes1 and cHes5 and met with limited success. PCR on mRNA extracted from

whole retinas reveal that these ligands are definitely present in the retina and their

expression can be regulated by growth factors, damage and Notch inhibitors. As

discussed in Chapter 3, we speculate that Notch ligands could be expressed by microglia and/or the NIRG cells. Some studies have reported the expression of Notch

ligands in microglia in rat brains (Cao et al. 2008) and that Notch-signaling controls

microglial activation and inflammation in the CNS (Grandbarbe et al. 2007). Further,

Notch-Delta signaling in microglia was shown to promote microglial release of

activators of neuronal proliferation in the brain (Morgan et al. 2004). Recent

experiments in our lab have shown that we can successfully deplete microglia and

NIRG cells from the retina using a combination of IL-6 and clodronate liposomes.

Clodronate-containing liposomes have been used to deplete macrophages (Hawkes and

McLaurin 2009; Horn et al. 2008) and microglia (Checchin et al. 2006; Li et al. 2009)

in the CNS. An indirect way to test whether Notch ligands are expressed on these cells

is to deplete them from the retina and determine the effects on Notch-signaling. We

hypothesize that depleting microglia and NIRGs from the retina would attenuate the

Notch-mediated effects on the proliferation of Müller-glial derived progenitors.

However, Müller glia would be more neuroprotective as Notch-signaling would be

decreased in the absence of the Notch ligands.

Implications of Notch-mediated neuronal survival

In the studies discussed in Chapter 3, we observe that suppressing the baseline

levels of Notch in the retina prior to NMDA or colchicine-induced damage enhances

173 neuronal survival. The survival effects were especially pronounced for the colchicine

studies, with a twofold increase in the number of surviving ganglion cells in nasal

regions of the retina and a threefold increase in the number of surviving cells in temporal regions. Although we have not tested the effects of Notch on photoreceptor survival, we speculate that blocking Notch activity in Müller glia may also protect photoreceptors from degeneration, similar to the effects seen on neurons in the inner

nuclear layer as well as the projection neurons in response to injury.

It would be interesting to see whether this cell survival in the retina equates to

preserved vision. The optokinetic drum is an appropriate tool to measure visual

responses in several species including mice (Puk et al. 2008), zebrafish (Huang and

Neuhauss 2008), chicks (Montiani-Ferreira et al. 2003), frog (Yucel et al. 1990), turtle

(Ariel 1997) and monkey (Buttner et al. 1983). We have recently constructed such tool

in the lab, which consists of a black-and-white striped cylindrical drum that is rotated

around the chicken. During the optokinetic response (OKR), involuntary eye

movements are evoked through coherent whole-field motion on the retina (Huang and

Neuhauss 2008). The resulting slow eye movements following the drum rotation (slow

phase) that are interrupted by fast resets in opposite direction (fast phase) are referred to

as optokinetic nystagmus (OKN) (Huang and Neuhauss 2008). Thus, the OKR is a

qualitative measure of visual acuity and is a good tool for behavioral retinal studies. We

speculate that chickens that received DAPT prior to colchicine will show better

optokinetic responses than those that did not receive DAPT.

174 Epigenetic mechanisms may control glial plasticity

Epigenetic mechanisms such as DNA methylation (mediated by the enzyme

DNA methyltransferases or DNMT), histone deacetylation (mediated by histone deacetylases or HDACs and histone acetyltransferases or HATs) and microRNA regulation may also directly affect the ability of retinal Müller glia to become gliotic, proliferative and/or progenitor-like cells. HDACs and HATs mediate the reversible acetylation of lysine residues in histone tails (Feng et al. 2007; Suzuki and Miyata

2006). Acetylation of histone tails is generally thought to relax the DNA backbone and facilitate transcription, while histone deacetylation silences the gene through chromosomal compaction (Chen and Cepko 2007; Feng et al. 2007), although the opposite may also be true, depending on the specific chromatin architecture (Feng et al.

2007; Nusinzon and Horvath 2005). HDAC inhibitors have been shown to induce neuronal differentiation in adult hippocampal progenitor cells. A report by Hsieh et al demonstrates that modification of chromatin by using the HDAC inhibitors valproic acid (VPA), Trichostatin A (TSA) or sodium butyrate (NaB) promotes neuronal differentiation, while suppressing oligodendrocyte and astrocyte differentiation in progenitors (Hsieh et al. 2004). In the developing mammalian retina, however, Cepko et al have shown that HDAC inhibition in retinal explant cultures resulted in the loss of photoreceptors, suggesting that HDACs play an important role in cell-fate decisions

(Chen and Cepko 2007), although the effects may vary depending on the cellular environment. The role of HDACs is yet to be determined during chicken retinal development and regeneration. Very little is known about the role of microRNAs in the retinal transcriptome (Karali et al. 2007). Walker et al have reported that microRNAs

175 play a role during retinal development by repressing apoptosis in Xenopus retina

(Walker and Harland 2009). Removal of Dicer, an RNase III endonuclease that is

essential for the production and function of mature miRNAs leads to retinal

degeneration (Damiani et al. 2008).

In previous studies and current studies discussed in Chapters 2, 3 and 4 , we

have seen that glia in peripheral regions of the retina are more responsive to damage

and/or growth factors injected into the retina, than glia in the central regions (Fischer

2005; Fischer et al. 2002; Fischer et al. 2004). Indeed, glia located in central regions of the retina are generated earlier than those located in peripheral regions (Prada et al.

1991), and the latter may hence be able to maintain a greater degree of plasticity in their phenotype. Additionally, the glia in lower vertebrates such as fish and frogs retain a greater capacity to regenerate than those in higher vertebrates (Hitchcock et al. 2004;

Raymond and Hitchcock 1997; Yurco and Cameron 2005), suggesting that fewer

‘epigenetic blocks’ are present in the glia of lower vertebrates. Thus, a goal for future studies could be to elucidate the epigenetic differences that may account for the varying degrees of phenotypic plasticity of Müller glia in response to injury or growth factors, depending on their location within the retina as well as investigating the effects of

DNMT/HDAC inhibitors as well as microRNAs on the phenotypic plasticity of Müller glia-derived cells in the chick retina.

176 Future directions for studies on serotonin-accumulating bipolar cells

Our exciting discovery of serotonin-accumulating bipolar cells in the retina has

led to several interesting questions. The only cell type known to synthesize serotonin in

the retina is the amacrine cell, and the primary cell type known to synthesize the

hormone melatonin in the retina is the photoreceptor (Chong et al. 1998). Serotonin is

the precursor of melatonin, which is an important regulator of retinal circadian rhythms.

Our results indicate that serotonin does not merely diffuse from amacrine cells into this

subtype of bipolar cells; it is specifically transported into SAB cells via the serotonin

transporter (SERT). The presence of serotonin in bipolar cells could indicate two possibilities : (1) the imported serotonin is degraded in SAB cells, as is revealed in our studies and/or (2) SAB cells serve as a temporary reserve of serotonin for melatonin synthesis in photoreceptors. Curiously, the mechanisms by which serotonin reaches photoreceptors remain uncertain, nor has serotonin ever been detected in them to the best of our knowledge. We hypothesize that SAB cells pass this neurotransmitter onto photoreceptors when the latter require it to produce melatonin. To test this hypothesis,

we would have to deplete SAB cells from the retina and measure melatonin levels at

various times of the day, using techniques such as High Performance Liquid

Chromatography (HPLC). A decrease in melatonin levels would indicate that serotonin from bipolar cells is required for melatonin synthesis and generating circadian rhythms within the retina. These findings could have important implications for understanding retinal circadian rhythms, dysfunctions of which could lead to abnormal photoreceptor function and degeneration.

177 Concluding remarks

The data presented in this dissertation adds to the growing body of knowledge about the

functions and properties of Müller glia in the retina. It is important to understand the

factors and signaling pathways (such as the Notch-pathway) that contribute to the

‘plastic nature’ of these cells. This information could be applied towards using these

glia as an intrinsic source of stem cells within the retina, to replace dying neurons,

preventing neuronal cell death and preventing gliosis, thereby preserving vision.

178

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