Cell-Specific Roles for CASK in the Pathology of Optic Nerve Hypoplasia

Cell-specific roles for CASK in the pathology of Optic Nerve Hypoplasia

Alicia Kerr

Dissertation submitted to the faculty of the Virginia Polytechnic Institute and State

University in partial fulfillment of the requirements for the degree of

Doctor of Philosophy In Translational Biology, Medicine, and Health

Michael A. Fox (Committee Chair) Gregorio Valdez Konark Mukherjee Harald Sontheimer John Chappell

April 25th, 2019

Roanoke, Virginia

Keywords: CASK, Optic Nerve Hypoplasia

Cell-specific roles for CASK in the pathology of Optic Nerve Hypoplasia

Alicia Kerr Abstract Optic Nerve Hypoplasia (ONH) is the leading cause of childhood blindness in developed nations and its prevalence has been rising. Yet, we know little about the genetic, molecular, or cellular mechanisms underlying ONH. A previous study described ONH in a cohort of patients with mutations in CASK, an X-linked gene with established roles in neural development and synaptic function. I have demonstrated that heterozygous deletion of CASK in mice (Cask+/-) recapitulates many of the phenotypes observed in patients with CASK mutations, including

ONH. This includes reduced optic nerve size, reduced numbers of retinal ganglion cells (RGCs), reduced RGC axonal diameter, and deficits in vision-related tasks. Further analysis on a homozygous partial loss of function variant (Caskfl/fl) also displayed ONH with reduced numbers of RGCs. In order to understand the mechanisms underlying CASK-associated ONH, I explored whether RGCs, the projection neurons of the retina and the cells whose axons comprise the optic nerve, generate CASK. Indeed, mRNA analysis revealed expression of CASK by a large cohort of RGCs. In order to assess whether loss of CASK from a majority of RGCs leads to

ONH, I crossed a conditional allele of CASK (CASKfl/fl) with transgenic mice that express Cre

Recombinase (Cre) in RGCs. Deletion of CASK from RGCs did not further alter ONH size nor

RGC survival. These results demonstrate that loss of CASK signaling in this discrete neuronal populations is not sufficient to lead to further disruption in the assembly of the subcortical visual circuit, suggesting a non-cell autonomous mechanism for loss of CASK in ONH.

Cell-specific roles for CASK in the pathology of Optic Nerve Hypoplasia

Alicia Kerr General Audience Abstract

The connection between the eye and the brain is crucial for successful vision. Impairment of this connection by either loss of the retinal neurons that project axons to the brain or damage to the nerve (optic nerve) lead to blindness. This occurs in a disease called Optic Nerve Hypoplasia

(ONH), which is the leading cause of childhood blindness in developed countries. Discovering the risk factors associated with this disease and mechanisms underlying the disease can help us build tools to treat and repair the optic nerve. Previously, mutations in the CASK gene were found in patients with ONH. Here, I developed a mouse model of CASK mutations to phenocopy the human patients, and used this model to explore the development of ONH. For example, with this mouse model I described for the first time, the timeline of disease progression. Surprisingly,

I also showed that loss of CASK specifically from the neurons whose axons generate the optic nerve did not lead to ONH, suggesting that ONH may develop from a failure of a network of cells, rather than just one population of cells.

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Acknowledgements To my mentor, Dr. Michael Fox, this accomplishment is a testament to your belief in me and there are not enough words to express my gratitude for that. Thank you for all the training about the practice of using the right language to express ideas, the importance of techniques in answering scientific questions, the necessity of patience when you have to repeat an experiment for what feels like the 700th time, and most importantly, the power of a good attitude.

To my committee members, and especially Dr. Konark Mukherjee, thank you for your support and guidance throughout this process.

To my labmates, Dr. Aboozar Monavarfeshani, Dr. Jianmin Su, Ubadah Sabbagh,

Rachana Deven Somaiya, and Gabriela Carrillo, science is a team sport and you all have been the best team. I am a better scientist and a better person because of the mentorship and comradery that you have all given me in your own individual ways. I am lucky to be surrounded by colleagues who care about the human being and not just their scientific output.

To my friends who have been so patient with me throughout this process. I am surrounded by an incredible group of women who challenge me to dream bigger and embolden me every step of the way. Thank you, Ashley Alley, Alexandria Pilot, Alyssa Moore, Lexie Mellis and every other person who said “Alicia, wash your hair and let’s go get a drink.”

Finally, to my family and especially to my sister Lucinda, thank you for the never ceasing love and encouragement. I am incredibly fortunate to have been raised in a family that has always encouraged creativity and seeking knowledge and puts tremendous value in hard work, traits that my grandparents are incredible representations of. I have dedicated this work to them,

Gary and Peggy Kerr and Richard and Penny Sabau. I truly believe that if I have accomplished anything it’s because of the example the four of them set for how to work hard, ask questions, and love and care for people along the way.

Table of Contents Abstract ...... ii General audience abstract ...... iii Acknowledgments...... iv Table of Contents ...... v List of Figures ...... viii 1. General introduction ...... 1 1.1. Optic Nerve Hypoplasia ...... 1 1.1.1. Causes of ONH ...... 2 1.1.2. CASK identified as risk factor for ONH ...... 5 1.2. Retinofugal circuitry in a murine model ...... 6 1.2.1. Mice as models of human retinal diseases ...... 6 1.2.2. Retinal cell types and lamination ...... 7 1.2.3. Non-neuronal cell types in the retina ...... 13 1.2.4. Dorsal Lateral Geniculate Nucleus of the thalamus ...... 15 1.3. CASK ...... 17 1.3.1. Role of CASK at the synaptic terminal ...... 18 1.3.2. CASK mutations in human disease ...... 20 1.4. Rationale, hypothesis and experimental design ...... 22 References ...... 37 2. Optic Nerve Hypoplasia is a pervasive subcortical pathology of visual system in neonates ...... 53 2.1. Abstract ...... 54 2.2. Introduction ...... 55 2.3. Results ...... 57 2.3.1. Mutations in the CASK gene are associated with ONH ...... 57 2.3.2. Heterozygous deletion of CASK in mice produces optic nerve pathology .....58 2.3.3. Axonopathy is present in optic nerves of Cask+/- mice ...... 59 2.3.4. Retinogeniculate synaptopathy is observed in Cask+/- mice ...... 61 2.3.5. RGC numbers are reduced in Cask+/- mice ...... 62 2.3.6. ONH develops postnatally in Cask+/- mice ...... 63 2.4. Discussion ...... 64 2.5. Experimental Procedures ...... 67

2.5.1. Identification of a splice mutation in CASK ...... 67 2.5.2. Mouse breeding and genotyping ...... 68 2.5.3. Immunohistochemistry ...... 68 2.5.4. Immunoblotting ...... 69 2.5.5. Optic nerve samples preparation for toluidine blue staining and transmission electron microscopy (TEM) ...... 69 2.5.6. G-Ratio distribution spectra ...... 69 2.5.7. Serial block-face scanning electron microscopy ...... 70 2.5.8. Intraocular injections of anterograde tracers and quantification ...... 70 2.5.9. Cell counts and lamina quantification ...... 71 2.5.10. Description of coarse and fine axons ...... 72 Acknowledgements ...... 72 References ...... 91 3. Non-cell autonomous roles of CASK in optic nerve development ...... 96 3.1. Abstract ...... 97 3.2. Introduction ...... 98 3.3. Results ...... 99 3.3.1. Heterozygous loss of CASK produces behavioral but not electroretinographical deficits in vision ...... 99 3.3.2. Partial loss of function of CASK in a 3-year-old boy produces ONH ...... 101 3.3.3. Global reduction in CASK expression produces ONH ...... 104 3.3.4. CASK is expressed in retinal ganglion cells ...... 106 3.3.5. RGC-derived CASK is not required for RGC survival ...... 106 3.4. Discussion ...... 107 3.5. Experimental Procedures ...... 110 3.5.1. Visual behavior tasks ...... 110 3.5.2. Electroretinography ...... 111 3.5.3. Plasmid and point mutagenesis ...... 112 3.5.4. Cell culture and imaging ...... 112 3.5.5. Molecular dynamics simulations and analysis ...... 113 3.5.6. Neurexin mediated cell recruitment assay ...... 114 3.5.7. Solubility assay ...... 115 vi

3.5.8. Pull down assay using NXCT and GST beads ...... 116 3.5.9. Immunoblots and immunohistochemistry ...... 116 3.5.10. Optic nerve samples preparation for transmission electron microscopy..... 118 3.5.11. Axon counts ...... 118 3.5.12. Optic nerve area ...... 118 3.5.13. G ratio ...... 119 3.5.14. In situ hybridization (ISH) ...... 119 3.5.15. Mouse breeding and genotyping ...... 119 3.5.16. RGC counts ...... 120 Acknowledgements ...... 120 References ...... 138 4. Discussion and Future Perspective ...... 143 4.1. Roles for CASK in development and ONH ...... 143 4.1.1. Cell autonomous roles for CASK ...... 145 4.1.2. Non-cell autonomous roles for CASK ...... 151 4.2. Optic nerve atrophy or optic nerve hypoplasia? ...... 152 4.3. Developing therapeutics for ONH ...... 154 4.4. Other causes of ONH and models to understand them ...... 156 4.4.1. Non-genetic causes of ONH ...... 157 4.4.2. Genetic variants of ONH ...... 158 4.4.3. How other causes of ONH relate to CASK ...... 159 References…………………………………………………………………………………...162 References ...... 171

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List of Figures Chapter 1 Figure 1.1. Optic Nerve Hypoplasia ------28 Figure 1.2. Structure of human and mouse eyes ------30 Figure 1.3. Retinal cell populations and birthdates ------32 Figure 1.4. CASK binding partners and proposed functions ------34 Figure 1.5. Proposed functions of CASK at synapse------36 Chapter 2 Figure 2.1. Genetic, brain and retinal alterations associated with heterozygous CASK mutation ------76 Figure 2.2. Cask+/- ON display reduced axons and increased astrogliosis ------78 Figure 2.3. Cask+/- ON display axonal thinning ------80 Figure 2.4. Cask+/- mice display a decreased number of retinogeniculate synapses and a reduced number of active zones in the remaining synapse ------82 Figure 2.5. Decreased number of retinal ganglion cells in Cask+/- mice ------84 Figure 2.6. Cask+/- mice optic nerve display secondary reduction in size ------86 Figure 2.7. Supplemental Figure 1------88 Figure 2.8. Supplemental Figure 2 ------90 Figure 2.9. Supplemental Figure 3 ------92 Chapter 3 Figure 3.1. Heterozygous loss of CASK produces behavioral but not electroretinographical deficits in vision ------124 Figure 3.2. Partial loss of function of CASK in a 3-year-old boy produces ONH ------126 Figure 3.3. Global reduction in CASK expression produces ONH ------128 Figure 3.4. CASK is expressed in retinal ganglion cells ------130 Figure 3.5. Calb2-Cre labels retinal ganglion cells ------132 Figure 3.6. RGC-derived CASK is not required for RGC survival ------134 Figure 3.6. Supplemental Figure 1------136 Figure 3.7. Supplemental Figure 2------138

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1. General introduction

1.1. Optic Nerve Hypoplasia

Thirty-six million people worldwide are legally blind and another 217 million have moderate to severe visual impairment (Bourne et al., 2017). While the prevalence of visual impairment rises with increased age, nineteen million children are estimated to have vision impairment (World Health Organization, 2012). Children with visual impairment are at risk for developmental delays and other learning impairments.

Moreover, the long duration of the disease burden in children increases the emotional, social, and economic burden of disease in this population. On a more positive note, however, the developing nervous system (including the retina) is more plastic and exhibits a greater ability to respond to therapeutics and treatment.

The leading cause of childhood blindness and visual impairment in children in developed countries is Optic Nerve Hypoplasia (ONH), responsible for 15% of all childhood blindness in 2012, an increase of 7% from the preceding decade (Kong et al.,

2012). This increase in ONH cases in the last decade follows a six-fold increase in diagnosis from 1970 to 2000 (Patel et al., 2006). Recent studies have confirmed that the increasing incidence of ONH is showing no signs of slowing down, in one region of

Canada it has been reported that 1 in 892 live births present with ONH, where a greater than 1 fold annual increase has been reported since 1996 (Khaper et al., 2017). These numbers are especially striking, as there are likely mild cases of ONH that are undiagnosed.

1 ONH was first described in 1884 (Magnus, 1884) and then more fully characterized in 1915 (Schwarz, 1915). During a fundoscopic exam, ONH is diagnosed when an optic nerve is one-third to one-half the size of a normal optic nerve (Figure 1.1.) or when there are irregularities of the optic disc such as the “double ring sign” (Mosier et al., 1978;

Lambert et al., 1987; Kaur et al., 2013). Patients with ONH often have other anatomical brain abnormalities, including the loss of the septum pellucidum, a membrane in the midline of the brain separating the lateral ventricles of the brain. When these phenotypes occur together it is termed Septo-Optic Dysplasia (SOD) (also known as De

Morsier’s syndrome) (De Morsier, 1956; Hoyt et al., 1970). SOD may also be accompanied with hypopituitarism in a condition called sypto-optic-pituitary dysplasia

(Hoyt et al., 1970; Clark and Meyer, 1978; Kaur et al., 2013). ONH, when it is present as part of SOD or septo-optic-pituitary dysplasia, is considered a syndromic form of ONH.

ONH can occur isolated in one eye (unilateral) or in both eyes (bilateral). Children with bilateral ONH have a higher comorbidity of Autism Spectrum Disorder and intellectual disabilities as compared to unilateral ONH, suggesting a different pathogenesis for bilateral ONH that may be more developmental and cerebral

(Fahnehjelm et al., 2013). The majority of ONH presents as bilateral, with unilateral cases only accounting for 15-25% of ONH (Garcia et al., 2006). In some cases there is a partial or incomplete appearance of ONH, classified as segmental ONH.

1.1.1. Causes of ONH

Historically, ONH has been considered a sporadic disease resulting from environmental factors. As a developmental disease, prenatal risk factors have been the most studied suspects (Garcia-Filion and Borchert, 2013). Maternal recreational drug

2 use of LSD, heroin, and alcohol use have all been described as potential causes in a few cohorts of patients with ONH (Margalith et al., 1984; Michaud et al., 1982).

Prescription drug use, including anticonvulsants and antidepressants, has also been identified as a risk factor (Hoyt and Billson, 1978; Tornqvist et al., 2002). In addition, maternal age, low birth weight, primiparity and other aspects of gestational history have been implicated as risk factors for ONH (Margalith et al., 1984; Murray et al., 2005;

Tornqvist et al., 2002). Recent epidemiological studies have strengthened the hypotheses that these environmental factors are major components of ONH as well as

SOD occurrence. In one such study, the prevalence of SOD was highest in babies born of mother’s 20-24 years of age (Garne et al., 2018), another retrospective review of cases of ONH showed that the median maternal age for ONH patients was 22.5 (Khaper et al., 2017). Maternal diabetes has been strongly linked with segmental ONH, in a case study of 10 patients with segmental ONH, all 10 patients were the children of diabetic mothers (Kim et al., 1989).

While it is likely that drugs and other environmental factors have teratogenic effects on the visual system and may create sporadic cases of ONH (Garcia-Filion and

Borchert, 2013), familial cases of ONH and recent advances of molecular diagnostic technology have suggested genetic risk factors for ONH also exist. For example, we were recently contacted by a family with 3 daughters, each diagnosed with ONH. Thus far, genetic forms of ONH have been associated with the mutation or loss of a variety of transcription factors: HESX1, PAX6 and SOX2. ONH, in its syndromic form with SOD, was reported in a pair of siblings by Dattani et al (1998) where they identified a mutation in homeobox transcription factor HESX1 in these patients. HESX1 is a transcriptional

3 repressor involved in forebrain development and determination of the pituitary gland.

Thus, mutations in HESX1 seem more associated with SOD as opposed to isolated forms of ONH (Dattani et al., 1998; Dattani et al., 2004; Andoniadou et al., 2007).

Mouse models with a null mutation of HESX1 show defects in forebrain structures, with a reduction in the anterior forebrain tissue seen as early as E8.5 and, notably, these mutant mice are anophthalmic (lacking eyes). Furthermore, these mice have substantial rates of perinatal/postnatal lethality (Dattani et al., 1998; Andoniadou et al., 2007).

In addition to HESX1, PAX6, a transcription factor originally identified in eye morphogenesis (Gehring, 1996), has been linked to congenital optic nerve anomalies

(Azuma et al., 2003). Of the 8 patients identified with PAX6 mutations, three were diagnosed with ONH. Human patients with PAX6 mutations have a wide variety of ocular defects not limited to ONH, including aniridia (lack of iris), congenital cataracts, and glaucoma (Jordan et al., 1992; Glaser et al., 1994; Sale et al., 2002, Chao et al.,

2003). Mouse models for PAX6 also reflect a spectrum of ocular and retinal defects including aniridia, cataracts, craniofacial developmental defects, and in some variants

(commonly called Small eyeless or Sey) anophthalmia is present (Hill et al., 1991; Wang et al., 2017; Ton et al., 1991). Like mouse mutants of HESX1, mice lacking PAX6 have high rates of perinatal mortality (Hogan et al., 1988). Mutations in a third transcription factor, SOX2 (sex determining region Y box 2), have been identified in two patients diagnosed with bilateral ONH and nystagmus (Kelberman et al., 2006). SOX2 is a transcription factor that regulates eye morphogenesis. Heterozygous SOX2 mutations have frequently been reported in humans as a genetic cause for anophthalmia/microphthalmia (Fantes et al., 2003; Bakrania et al., 2007; Gerth-Kahlert

4 et al., 2013). However, heterozygous SOX2 mutations in a mouse model do not recapitulate the anophthalmia phenotype shown in humans, but do show abnormal pituitary development (Kelberman and Dattani, 2006, Taranova et al., 2006).

As I was interested in elucidating a mechanism of ONH these genes were poor models for a variety of reasons. First, their role as transcription factors mean they have the potential to influence and regulate a number of intracellular pathways that could directly or indirectly induce ONH. Second (but perhaps related to their role as transcription factors), their loss or mutation produces a mosaic of ophthalmic phenotypes from aniridia to anophthalmia. At best, this complicates analysis, however in some cases (like anophthalmia) it makes it not possible to study ONH. Since using the loss or mutation of these genes as animal models to study ONH is not feasible, we sought other genes linked to ONH that may provide a better model to study this devastating disease. This drove our attention to CASK, an X-linked presynaptic adaptor molecule whose mutation leads to ONH (Moog et al., 2011). Furthermore, two different mouse models of CASK mutation already existed and in both cases they do not display the same variety of ocular defects as shown in other models.

1.1.2. CASK identified as risk factor for ONH

In 2011, mutations in the CASK gene were identified in a population of 25 females with a variety of phenotypes including postnatal microcephaly, developmental delays, and ONH (Moog et al., 2011). CASK, an X-linked gene, had previously been implicated in X-linked brain malformation and microcephaly and cerebellar hypoplasia, as well as in other ophthalmic defects including nystagmus and optic nerve atrophy (Najm et al.,

5 2008; Hackett et al., 2010). As opposed to the other genes discussed above, CASK is a more ideal target for studying mechanisms of ONH. As the other genes discussed are all transcription factors, CASK’s role as a synaptic scaffolding protein with known binding partners makes elucidating CASK’s mechanism of action more sustainable.

1.2. Retinofugal circuitry in a murine model

To understand mechanisms underlying the development of ONH, we must understand the retina. Located in the posterior chamber of the eye, the retina is a specialized outcropping of brain tissue that is responsible for detecting, processing and transmitting light-derived information to the brain. Much of this light-derived information is important for forming perceptual constructs of the visual world in sighted animals and must include information about several visual features including captures color, size, shape, motion of other animals and the environment. In addition to this “image-forming” visual information, light-derived signals are also important for non-image forming processes such as pupillary reflexes and photo-entrainment of circadian rhythms.

1.2.1. Mice as models of human retinal diseases

Before I discuss the structure and function of the murine retina and retinal targets, it is important to consider whether using this model will accurately reflect the human condition of ONH. Apart from our ability to genetically modify them, mice have been a valuable resource to enhance our understanding of human disease development and the pathogenesis of these diseases for more than 90 years (Keeler, 1924; Dalke

6 and Graw, 2005). Structurally, mouse eyes develop similarly to human eyes with three notable differences (Figure 1.2.). First, the mouse lens is spherical in shape and much larger than its human counterpart when compared to the size of the eye is housed in, reducing the intravitreal space of the mouse eye (Sundin, 2005). Second, the mouse lacks a macula, the region of the retina with the highest density of photoreceptors

(Volland et al., 2015). Third, as a nocturnal animal, the mouse has fewer cone photoreceptors so it is more primed to see movement in the dark (Jeon et al., 1998).

Despite these differences the diversity, density and morphology of other cells in the mouse retina resemble those in the human retina. Not only are mouse retinas structurally similar human retinas (Seabrook et al., 2017), recent work has revealed similar transcriptome profiles of human and macaque retina compared with mice

(Hoshino et al., 2017; Peng et al., 2019). For example, there is high expression of Math5 in the development of both murine and human eyes (Brown et al., 2002, Hoshino et al.,

2017). Thus, with so many shared molecular, cellular and morphological features with the human retina, the mouse retina (and eye) serves as an appropriate model to understand human ONH.

1.2.2. Retinal cell types and lamination

The retina is responsible for both detection of light and transmitting that information to retino-recipient locations in the brain. To perform both of these functions requires an intricate network of different retinal neurons that are anatomically segregated into distinct layers of the neuro-retina. The three nuclear layers of the retina contain five

7 major types of neurons: photoreceptors, bipolar cells, horizontal cells, amacrine cells, and retinal ganglion cells (RGCs) (Figure 1.3.A).

Light enters the eye and is absorbed by photoreceptors in the outermost cell layer, aptly named the Outer Nuclear Layer (ONL). Photoreceptors contain photopigments that have a light absorbing component (11-cis retinal) and a protein component (opsin).

There are two main types of opsins in retinal photoreceptors, rhodopsins and cone opsins, which detect different elements of light information. These two types of opsins differentiate the two types of photoreceptors: rods and cones. Rods, which house rhodopsins, are important for detecting contrast and are especially important for scotopic (night) vision, as rhodopsins are sensitive to dim light stimulation and can amplify light signals more than cones can. Cones, which house cone opsins, function to detect color information from stimuli. In fact, there are three specialized types of cone opsins which respond to different parts of the light spectrum. In both types of photoreceptors, photoactivation of these opsins causes a biochemical cascade in these cells which results in a hyperpolarization of the cell. Rather than generating an action potential in response to stimulus, they create graded potentials that relay information about changes in light intensity as detectable changes in membrane potential. This allows photoreceptors to transmit information about not just whether light is present or not, but any changes in that light information. As nocturnal animals, mice rely heavily on rods for detection of light in scotopic conditions. In fact, rods vastly outnumber cones in the murine retina (97.2% of cells in the ONL are rods and 2.8% are cones) (Jeon et al.,

1998, Carter-Dawson and LaVail, 1979a). While the high ratio of rods in mouse retina

8 differs from the human macula (and fovea) it is remarkably similar to the proportion of rod photoreceptors in the human peripheral retina (Jonas et al., 1992).

Photoreceptors directly innervate glutamatergic interneurons called bipolar cells that reside in the Inner Nuclear Layer (INL). Bipolar cells account for ~41% of the total cell population in the INL (Jeon et al., 1998), and can be divided into two types, either they are depolarized by light increments (ON Bipolar) or hyperpolarized by light increments

(OFF Bipolar). Bipolar cells synapse on RGCs, which reside in the ganglion cell layer

(GCL). Most ganglion cells receive their main excitatory input from bipolar cells in a direct pathway, and thus ganglion cells also have on or off responses.

Apart from on and off responses, RGCs also respond to spatial and temporal contrasts of light as most of the useful information to build a visual scene is contained in patterns of contrasts. This is done by establishing receptive fields for both bipolar cells and RGCs. Receptive fields are areas of neighboring photoreceptors or bipolar cells that converge onto one cell, forming an area which bipolar cells or RGCs can “monitor” for different types of stimulation patterns. These receptive fields are circular and can be divided in to two parts: a circular zone at the center of the receptive field, called the center, and the remaining area of the receptive field, called the surround.

To relay information about the surround receptive field, the retina has formed lateral pathways using interneurons called horizontal cells. Photoreceptors in the surround of the receptive field will innervate this horizontal cell rather than the bipolar cell. The horizontal cell then feedbacks onto photoreceptors in the center of the receptive field.

These horizontal cells transmit signals passively and, using gap junctions, can relay information from even more distant photoreceptors. Other cells in the lateral pathways

9 that mediate complex visual patterns are interneurons called amacrine cells that are primarily inhibitory. There are 20 distinct types of amacrine cells responsible for mediating different aspects of visual stimulation, from starburst amacrine cells that are necessary for direction-selective computation (Vaney et al., 2012) to AII and A17 amacrine cells that have roles in night vision (Grimes et al., 2010). Both amacrine and horizontal cells reside in the INL (Figure 1.3.A) where horizontal cells account for 3% of the population of cells and amacrine cells account for 39% (Jeon et al., 1998).

Interestingly, the Ganglion Cell Layer (GCL) also contains displaced starburst amacrine cells, with some estimates that these cells account for ~50% of the neurons in the GCL

(Jeon et al., 1998).

Ultimately both direct and lateral pathways in the retina end at the RGCs, which are the soleprojection neurons responsible for sending axons into the brain via the optic nerve. With rare exceptions, all RGC somas are located in the GCL and their dendrites arborizein the inner plexiform layer. All RGCs are glutamatergic, however they can divided into many different subtypes using at least four criteria (Sanes and Masland,

2015) including: 1) Morphology. A classification that dates back to Ramon y Cajal,

RGCs have been labeled and grouped based on properties such as soma size, dendritic morphology, dendritic branching patterns, and dendritic stratification within the IPL

(Cajal, 1892; Doi et al 1995; Weng et al, 2005, Volgyi et al 2005). 2) Molecular Markers.

There is a genetic diversity amongst RGCs, as single cell RNA sequencing has been applied to segregate RGCs into 40 distinct clusters based on transcriptome profiles

(Rheaume et al., 2018). Though this single cell data provides a new thorough perspective on gene profiling in RGCs, previous work has identified some major players

10 in different classes of RGCs. As we have developed transgenic lines and other genetic markers, a variety of genes have been identified that are generated in subtype-specific patterns in RGCs, such as cadherins (Osterhout et al., 2011), Islet2 (Kay and Triplett,

2017), and SMI32 (Coombs et al., 2006). As the physiology of RGCs is identified, some of these genes have been grouped together within distinct sub-classes. For example,

Opn4, Eomes, and Igf1are all genes indicative of melanopsin-expressing intrinsically photosensitive RGCs (ipRGCs) (Mao et al., 2014; Hattar et al., 2002; Berson et al.,

2010). 3.) Spacing. RGCs tile within the retina, where sampling from one area of the retina reveals a variety of subtypes of RGCs as RGCs from one class will avoid cells within their class during migration, creating a functional diversity of RGCs (Rockhill et al., 2000; Reese, 2008). 4) Physiological properties. This is indicative of the retinal area that an RGC “monitors” and whether an increase in light delivered to the center of their receptive field will excite the RGC (ON-center) or inhibit the RGC (OFF-center), and similarly for stimulation of the edges “surround” of the receptive field. Further classification includes ON-OFF RGCs, which are excited by both increase and decrease in light intensity. This category has been further developed as RGCs also respond to movement of light throughout their receptive field, indicated by direction sensitivity (DS)

(up versus down, left versus right, etc.) (Barlow and Levick, 1965; Nelson, 1978, 1983;

Huberman, 2009; Baden et al., 2016). As we develop better techniques to assess each category, we can combine these techniques and identify subtypes that possess specified aspects of each category. For example, ON directionally sensitive RGCs

(category 4) all display a distinct dendritic morphology (category 1). Grouping or

“lumping” these categories reduces the number of subtypes of RGCs and by these definitions there are around 25 different subtypes of RGCs in mammals (Sanes and

11 Masland, 2015). Other groups take a “splitting” approach where there is a narrower definition of each subtype. In these classifications as many as 40 RGC subtypes have been identified in mammals (Baden et al., 2016; Rheaume et al., 2018). 80 RGC subtypes have been proposed in zebrafish (Robles et al, 2013).

All six neuron classes in the retinas, as well as Muller glial cells, are derived from retinal progenitor cells. This process of cell-differentiation is tightly controlled by environmental signals and transcription factors (Reh, 1991; Reichenbach et al., 1998) and occurs in such a way that the generation of each cell type occurs in an order which is conserved amongst all species studied (Wetts and Fraser, 1988; LaVail et al., 1991;

Stiemke and Hollyfield, 1995; Belliveau and Cepko; 1999; Hu and Easter, 1999). RGCs are among the first type of neuron born in the retina, with peak differentiation happening around E10.5, though some subtypes are born considerably later (Figure 1.3.B)

(Osterhout et al., 2015). RGCs quickly extend axons out of the retina (through the optic disc), and by E12 these axons can be seen crossing the midline of the brain at the optic chiasm. At this point in the retina other cell types are becoming postmitotic, including horizontal cells, cones, and amacrine cells (Carter-Dawson and LaVail, 1979b). By E15 some RGC axons have reached the most distant retinorecipient area of the brain, the superior colliculus of the midbrain (Drager, 1985; Godement et al.,1984) and in the retina, both horizontal and amacrine cells have migrated to their target area and begun developing dendrites (Reese, 2011). By P3, as RGCs are forming sublamina-specific dendritic arbors in the IPL of the retina (where they receive input from bipolar cells) and forming axonal arbors and synapses on target neurons in retinorecipient areas of the brain, in the retina, rods, bipolar cells, and Muller glial cells are becoming postmitotic

12 (Carter-Dawson and LaVail, 1979b; Ilia and Jeffery, 2000). By eye opening, around P12 in the mouse, RGC synapses have already gone through activity dependent refinement which segregates retinal arbors in retino-recipient nuclei into both topographic maps and eye-specific domains (Campbell et al., 1992; Lo et al., 2002; Hong and Chen, 2011).

Likewise, by eye-opening the rest of the retinal neurons have established their canonical dendritic morphologies to complete the retinal circuits. By the end of development, ~6.4 million rods and ~50,000 RGCs are born in mice (Jeon et al, 1998).

1.2.3. Non-Neuronal Cell Types in Retina

In addition to neuronal cell types, three different types of glial cells reside in the murine retina: Müller glial cells, microglia, and astrocytes. These cells are important for a variety of roles involved in retinal homeostasis including structural support, formation of blood-retina barrier, regulation of neuronal activity, synaptic pruning, and metabolism

(Hollander et al, 1991; Newman, 1996). In different states of retinal damage and disease such as increased intraocular pressure, as in glaucoma, ageing, and diabetic

retinopathy, glial functions become impaired (Mizutani et al., 1998; Puro et al., 2002;

Williams et al., 2017). Astrogliosis within the retina has been described in conditions

ranging from retinal detachment (Erickson et al., 1997, Okada et al., 1990), Alzheimer’s

disease (Blanks et al., 1996), transient retinal ischemia (Osborne et al., 1991), and

mechanical injury to the eye (Wen et al., 1995) suggesting that glial cells are sensitive to

a variety of non-homeostatic conditions in the retina.

13 Müller glial cells are the most predominant glial subtype in the retina and are the last cells derived from retinal neural progenitor cells to become post mitotic (Young, 1985;

Cepko et al., 1993) (Figure 1.3.B). They account for 16% of the total cells in the inner nuclear layer (Jeon et al, 1998) and 90% of all retinal glia (Vecino et al, 2016). As they share a progenitor with RGCs, amacrine cells, bipolar cells, and cones (Sidman et al.,

1961; Vecino and Acera, 2015) and because their bodies span from the ONL to the GCL

it has been suggested that Müller cells are necessary to form columnar arrays in the

retina, contacting all neuronal types in the retina and promote formation of functional

synapses onto bipolar cells and RGCs, thereby providing both structural and functional

support in the retina (Pfreiger and Barres, 1996; Newman and Reichenbach, 1996).

Müller cells are also known to ensheath blood capillaries and thus are thought to be

involved in the exchange of molecules like glucose from retinal blood vessels to neurons

(Mantych et al, 1993; Reichenbach and Robinson, 1995). Like astrocytes in the brain,

Muller glia are responsible for the uptake and recycling of neurotransmitters (e.g.

GABA and glutamate) and for synthesis and storage of glycogen (Newman and

Reichenbach, 1996; Reichenbach et al., 1993).

Retinal neural progenitor cells are the only progenitor pool in the retina. However,

due to the opening at the optic nerve head, other cells are able to migrate into the

retina. This includes astrocytes which migrate along blood vessels to enter the

developing retina (Stone and Dreher, 1987; Turner and Cepko, 1987; Watanabe and

Raff, 1988). The presence and distribution of these astrocytes in the retina are

concurrent with the distribution of the vasculature of the retina (Vecino et al., 2016).

Astrocytes are known to produce vascular endothelial growth factor (VEGF) and thus

14 are necessary for vascularization of the retina (Stone et al, 1995). Astrocytes also

increase neurotrophic support as well as maintain the blood-retina barrier. Though

astrocytes are necessary for normal functioning of the retina, reactive astrocytes are

known to promote RGC death and degrade the extracellular matrix within the retina

(Zhang et al, 2004). Astrocytes are different from other glial populations in the retina as

they are limited to the vitreal surface and thus play a pivotal role in maintaining

endothelial barrier properties (Ridet et al., 1997).

Microglia are also present in the retina and control immune responses in the eye, secreting cytokines and neurotrophic factors, as well as eliminating cellular debris

(Carson et al, 2006). Microglia are also important for remodeling of neuronal circuits

(Kettenmann et al, 2013; Prinz et al 2011).

1.2.4. Dorsal Lateral Geniculate Nucleus of the Thalamus

RGC axons exit the retina at the optic disc and extend through the optic nerve, chiasm, and tract to target more than 40 distinct subcortical retinorecipient areas in the brain (Mastersteck et al., 2017; Morin and Studholme, 2014). These retinorecipient areas include brain regions that process image forming visual information and nonimage forming information (such as those that regulate pupillary reflexes and eye movements).

In mice, the largest target of retinal axons are superior collicular and the dorsal Lateral

Geniculate Nucleus (dLGN) of the thalamus. The dLGN is a thalamic relay station that transmits visual information to primary visual cortex by retinorecipient thalamocortical

(TC) relay cells. These relay cells were historically believed to be, as their name

15 suggests, simple “relays” of visual information to primary visual cortex (Hubel and

Wiesel, 1961; Lee et al., 1983). Recently, evidence for higher rates of convergence of different types of RGC onto TCs suggests that there is a level of processing that is occurring at the dLGN (Hammer et al., 2015; Morgan et al., 2016; Rompani et al., 2017).

In the mouse, the dLGN is organized into three different types of maps

(Kerschensteiner and Guido, 2017). 1) Eye-specific maps. Anterograde tracers (such as fluorescently conjugated Cholera toxin subunit B) have revealed projections from contra- and ipsi-lateral projecting RGCs (Jaubert-Miazza et al., 2005; Hammer et al.,

2014). In mouse, with laterally displaced eyes, 95% of projections to dLGN originate from the contra-lateral eye and the remaining 5% of projections originating from the ipsilateral eye (Godement et al., 1984; Reese, 1998). 2.) Cell-type specific maps. The majority of RGC classes have been identified to project to dLGN in mice (Ellis et al.,

2016) and, using labeling strategies, patterns have emerged showing different subclasses preferentially targeting specific regions of the dLGN. For example, many types of direction sensitive RGCs target the outer “shell” of the dLGN, as opposed to

ON RGCs which target the inner “core” (Cruz-Martin et al., 2014; Ellis et al., 2016).

There is also a diversity of TC morphology, originally identified in the cat (Friedlander et al., 1981; Stanford et al., 1981; Krahe et al., 2011), TCs in the mouse dLGN also bear resemblance to these three morphological classes: X (bi-conical), Y (symmetrical) and

W (hemispheric). These three classes of TCs help to further distinguish the “shell” and

“core” of the dLGN as W cells reside exclusively in the shell (Bickford et al., 2015) and X and Y reside in sub sections of the core. 3) Topographic map. Axons of neighboring

16 RGCs project to neighboring TCs, forming a spatially preserved map of visual information (Cang and Feldheim, 2013).

Relay cells in the dLGN project their axons to only two ipsilateral regions in nocturnal rodents: primary visual cortex (V1) which processes visual information for action and the thalamic reticular nucleus (TRN) which modulates activity of the dLGN (McAlonan et al.,

2006; Crabtree and Killackey, 1989; Reese and Cowey, 1983; Warton et al., 1988). As our understanding of classes of RGCs and TCs has developed, our understanding of the presence of specialized, parallel retino-geniculo-cortical pathways in the mouse visual system has grown, such as a distinct pathway for DSGCs through the shell of the dLGN (W cells) to layer I of V1 (Cruz-Martin et al., 2014) and non-DSGCs through X and

Y cells to layer IV of V1 (Bickford et al., 2015). We have yet to identify class-specific projections to the TRN.

1.3. CASK A known genetic risk factor for ONH (Najm et al., 2008; Hackett et al., 2010), CASK

(Calcium/calmodulin-dependent Serine Protein Kinase) is an X-linked gene that encodes for a MAGUK (Membrane-Associated Guanylate Kinase) presynaptic scaffolding protein (Mukherjee et al., 2008). MAGUK proteins are defined by their PDZ

(Post-synaptic density 95, Disc large, Zonula occludens-1), SH3 (Src homolog 3), and

GK (Guanylate kinase) domains. While CASK is included in the p55 subfamily of

MAGUK proteins, defined by the inclusion of one or two L27 domains, it differs from other members of this subfamily due to the presence of a CaMK-like domain in the N- terminal portion of CASK (Figure 1.4A; Hata et al., 1996; Mukherjee et al., 2008).

1.3.1. Role of CASK at the synaptic terminal

CASK was discovered as being a critical gene in development and neuronal synaptic function in three different species at nearly the same time. Genetic screens in

Drosophila and C. elegans identified animals with CASK mutations that exhibited abnormal mobility or vulva development, respectively (Martin and Ollo, 1996; Hoskins et al., 1996). At the same time, biochemical approaches in vertebrates identified CASK as a binding partner of the C-terminal domain of neurexin, a presynaptic membrane protein important for synapse formation and function (Hata et al., 1996; Ushkaryov et al., 1992;

Ullrich et al., 1995). Neurexins and their post-synaptic binding partners, neuroligins, were the first transsynaptic cell adhesion molecules implicated in regulating synaptogenesis in the brain (Scheiffele et al., 2000; Graf et al., 2004; Chih et al., 2005).

Different isoforms of neurexins and neuroligins are important for different classes of synapses (Graf et al. 2004). It has also been shown in vivo that neurexins are essential for coupling voltage gated calcium channels to the presynaptic release machinery

(Missler et al., 2003), a process necessary for calcium triggered neurotransmitter release in the neuron. Binding of CASK with neurexin has been shown to be integral for the targeting of neurexin to presynaptic sites and is therefore vital for proper function of neurexin (Fairless et al., 2008). These findings led to the hypothesis that CASK is essential for synapse formation and function. However, initial reports showed that deletion of CASK does not alter the ability of a neuron to form a structurally normal synapse in vitro with only limited functional defects (Atasoy et al., 2007).

18 Aside from its ability to bind neurexin, analysis of CASK protein initially showed no autophosphorylation or kinase activity, but that it does act in a Ca2+-dependent manner

(Hata et al., 1996). The CaMK-like domain, though a protein kinase domain, was not expected to have catalytic activity due to residues of the Mg2+-binding motif being mutated and thus CASK is often called a “pseudokinase” (Boudeau et al., 2006). Recent work showing the structure of CASK challenged this, as CASK forms a constitutively active conformation. Further work shows that CASK functions as an Mg2+-independent neurexin kinase (Mukherjee et al., 2008).

Because of the presumed inactivity of CASK, most studies have focused on its role as a scaffolding protein. As a multidomain protein, each domain of CASK has been demonstrated to interact with a number of different proteins, implicating CASK in many cellular processes even beyond the presynapse. A summary of these potential roles for

CASK beyond the synapse is shown in Figure 1.4B. However, CASK is primarily a synaptic protein and has been implicated in many different functions at the synapse

(Figure 1.5). CASK’s ability to bind to many different proteins is integral to its multitudinous roles at the synapse. The CaMK-like domain of CASK binds to cellular proteins such as Mint1, Caskin1, and Liprin suggesting a role in organization of proteins at the synapse. In drosophila, the CaMK-like domain has been shown to bind with

CaMKII and localize it to the synapse in a calcium dependent manner (Lu et al., 2003).

The regulation of CaMKII by CASK further suggests it plays a critical role in neurotransmission as CaMKII is critical for vesicle mobilization in the active zone (Wong et al., 2008). The PDZ domain of CASK forms a hydrophobic pocket which allows a free

C-terminal residue to fit in and is the site at which CASK is known to bind with

19 cytoplasmic domains of transmembrane proteins such as neurexins, syndecans, and synCAMS, all of which are cell-adhesion molecules (Biederer et al., 2002; Cohen et al.,

1998; Hata et al., 1996). The SH3 domain of CASK interacts with N-type Ca2+ channels, mutations in this domain impair synaptic distribution of these channels (Maximov and

Bezprozvanny, 2002), as well as interacting with the GK domain of other MAGUK proteins forming large scaffolding complexes (Nix et al., 2000). The GK domain of

CASK not only binds to the SH3 domain in CASK creating a homodimer (Nix et al.,

2000), it also interacts with nuclear proteins such as T-box transcription factor 1 (TBR1)

(Hseuh et al., 2000) and CASK Interacting Nucleosome Assembly Protein (CINAP) (Lin et al., 2006). The two L27 domains of CASK interact with L27 domains of other proteins,

L27A interacts with the L27 domain of SAP97, a PSD-95 protein (Lee, 2002). L27B interacts with the L27 domain of Veli/mLIN-7/Mals (Setou et al., 2000). These domains of CASK and their interactions are summarized in Figure 1.4.

Protein interactions suggest three potential roles for CASK: 1.) Synaptic adhesion

(PDZ domain). 2.) Protein trafficking and synaptic targeting (CaMK, L27B, SH3 domains) and 3.) Regulating gene expression (GK domain).

1.3.2. CASK mutations in human disease

CASK was initially thought to be an essential gene and homozygous knockout of

CASK in mice was lethal (Atasoy et al., 2007). However, in lower organisms, such as C. elegans and Drosophila, CASK deletion is compatible with life (Slawson et al., 2011;

LaConte et al., 2013), suggesting that fundamentally CASK may not be an essential

20 gene. The majority of CASK mutations reported in human diseases have been heterozygous mutations (Najm et al., 2008; Piluso et al., 2009; Moog et al., 2011), though in some cases homozygous deletions have been reported (Saitsu et al., 2012).

Though CASK is a part of the presynaptic machinery, deletion of CASK does not alter the ability of a neuron to form a structurally normal synapse in vitro with only limited functional defects (Atasoy et al., 2007).

Patients with CASK mutations were identified to have a spectrum of brain malformation phenotypes such as microcephaly with pontine and cerebellar hypoplasia

(MICPCH) (Najm et al., 2008). Mutations in this X-linked gene have since been shown in patients with X-Linked intellectual disability (XLID) (Burglen et al., 2012; Hayashi et al.,

2017). A catalog of the phenotypic spectrum of CASK mutations in patients showed many ophthalmological conditions such as nystagmus, strabismus, optic nerve atrophy, and optic nerve hypoplasia (Hackett et al., 2010; Moog et al., 2011). Defects in brain development in patients with mutations in CASK have been hypothesized to arise due to its interaction with the transcription factor TBR1 and its downstream targets Reelin and

GRIN2B, genes critical for neuronal migration and cortical development (Hseuh et al.,

2000; Najm et al., 2008).

As CASK mutations have been linked to XLID and other neurodevelopmental disorders known to have impairment in cognitive function, it is important to note the difference in visual perception versus integration. It has historically been thought that the subcortical pathway was exclusively involved in the perception of visual information and had no role in the processing or interpretation of these signals. While recent work has challenged this concept of the thalamus as a “relay center” to the cortex (Hammer et

21 al., 2015; Morgan et al., 2016), it has been shown that retinal phenotypes and visual impairments can arise independent of higher processing abnormalities in certain neurodevelopmental disorders (Rossignol et al., 2014). Understanding the mechanisms of ONH, a disease of the subcortical visual system, is thus critical not only for the many children affected by ONH, but also may further our understanding of how healthy and abnormal subcortical visual systems develop. This information may prove to be valuable to understanding other neurodevelopmental disorders such as Fragile X Syndrome which results in a sensory impairment that has both central integration defects and peripheral perception defects (Rossignol et al., 2014). Analysis of our CASK model of

ONH may provide insight in how the peripheral perception defects occur separate from the central integration defects.

1.4. Rationale, hypothesis and experimental design

Of the genes that have been linked to ONH, CASK is the most ideal target for developing an animal model to understand mechanisms of ONH. Though ONH may present with other cerebral defects in a syndromic fashion, to understand how ONH develops, I was interested in developing a model to study ONH in its isolated form.

Understanding the molecular mechanism of ONH and how development and maintenance of the subcortical visual system is altered in this disease will be valuable to our ability to develop therapeutics to treat them.

Other genes identified as potential links to ONH, such as HESX1 and PAX6, are transcription factors that cause a variety of devastating defects including craniofacial

22 abnormalities and anophthalmia in both animal models and human patients when mutated (Hill et al.,1991; Wang et al., 2017; Ton et al.,1991), showing that these genes play larger roles in eye and brain morphogenesis. As such, these transcription factors do not represent the isolated ONH phenotype. Furthermore, as transcription factors, elucidating a mechanism amongst potential downstream targets is difficult.

Heterozygous deletion of PAX6 from mouse models results in 559 genes differentially expressed in just the lens as compared to wild-types at P1 (Wolf et al., 2009).

Therefore, CASK is a more ideal target since 1. It is not a transcription factor, and elucidating a mechanism of ONH pathogenesis is more feasible and 2. Though complete knockout of CASK is lethal in mice (Atasoy et al., 2007), these mice are born with no macroscopic defects in the brain, suggesting that though CASK is important for neuronal function, it isn’t playing an essential role in eye or brain morphogenesis. Thus,

I was interested in the molecular mechanism with which CASK mutation leads to ONH.

As an X-linked gene, the majority of patients with CASK mutations are heterozygous females, as homozygous and hemizygous mutations likely lead to miscarriage (Hiyashi et al., 2017). Using a mouse model that replicates mutations seen in humans is essential for our ability to translate the results and develop functional therapeutics. Therefore, one of my aims was to test the hypothesis that heterozygous loss of CASK (Cask+/-) would produce ONH in mouse models. To test this hypothesis, I analyzed not only optic nerve area, but different aspects of subcortical visual system including RGCs and their ability to form synapses in the dLGN in the absence of CASK.

My data presented in Chapter 2 of this dissertation (following chapter) showed that, using this Cask+/- mouse model, I was able to replicate many aspects of ONH that are

23 observed in human patients with ONH such as a reduction in the number of RGCs and a decrease in RGC axon diameter as well as the reduction in optic nerve area. In humans this combination of phenotypes lead to a visual impairment, so I hypothesized that due to the alterations in the subcortical visual system in these mice I would also observe a visual deficit. To test this hypothesis I performed a behavioral analysis with a two-alternative forced swim assay to measure visual acuity and contrast sensitivity in these mice (Prusky et al., 2000; Monavarfeshani et al., 2018). Furthermore, to validate that the reduction in RGCs and their axonal diameter in these mice are responsible for any change in visual behavior I observe, it is important to consider that though retinal lamination remains unadulterated, there may be a functional defect in photoreceptors or bipolar cells. Thus, I performed electroretinography alongside the visual behavior assay to test functional output of photoreceptors and bipolar cells.

As I have validated Cask+/- as a sufficient model to study ONH, it can then be used to answer outstanding questions regarding both ONH and CASK. One such question in the field is whether ONH is a true hypoplasia (disease of development) or if it is mislabeled and is actually ONA (a disease of degeneration). Furthermore, does the reduction of optic nerve predate the loss of RGCs or does the loss of RGCs cause the reduction in the optic nerve? Using this model I analyzed the developmental timeline of both the optic nerve area as well as RGC counts and observed that reduction of the optic nerve occurs postnatally but before eye-opening and that RGC reduction is a secondary event to optic nerve area reduction.

As a multidomain protein CASK has been identified to interact with a large number of proteins involved in different cellular process, from transcriptional regulation

24 to synaptic adhesion. In vivo CASK knockdown in Drosophila has shown seemingly contradictory results: one group showed deletion of CASK leading to an increase in spontaneous neurotransmitter release at the neuromuscular junction (NMJ) with no alteration in vesicle size (Zordan et al., 2005) and another showed loss of CASK leading to less evoked synaptic transmission and fewer spontaneous synaptic events in

Drosophila NMJ (Chen and Featherstone, 2011). In vitro analysis of CASK knockout neurons showed only minor alterations in synaptic function and these synapses are ultrastructurally normal (Atasoy et al., 2007). Though the null mutation in CASK is lethal perinatally in mice, it has been proposed that because there are no macroscopic defects of the brain this lethality is not due to a failure of synapse formation (Atasoy et al.,

2007). The two different CASK mouse models described (Cask+/- and Caskfl/fl) provides us an opportunity to evaluate the role of CASK in synapse formation in vivo beyond perinatal ages. Importantly, I was interested in whether CASK produces ONH and synaptic deficits in a cell autonomous way, as CASK has been shown to recruit presynaptic machinery in a cell autonomous way (Chen and Featherstone, 2011).

Though we have known that CASK is widely expressed in the brain, to analyze

CASK’s role in the development of the subcortical visual system, it was important to first establish CASK expression in the retina, and particularly in RGCs. To accomplish this, I developed riboprobes against CASK mRNA and was able to visualize expression of

CASK in RGCs. Based on this result, I hypothesized that loss of RGC-derived CASK would lead to ONH through a cell-autonomous mechanism. Using a conditional allele of

CASK (Caskfl/fl) I deleted CASK from a majority of RGCs using a Cre-recombinase

(Calb2-Cre). My data, presented in chapter 3, showed that surprisingly, in this mutant

25 model (where there is already a global reduction in CASK levels) complete deletion of

CASK from a majority of RGCs did not exacerbate the ONH phenotype as compared to global reduction of CASK. This suggests that RGC-derived CASK is not necessary for

RGC survival, an unexpected discovery.

26 Figure 1.1. Optic Nerve Hypoplasia (A) Representative ophthalmoscopy (fundoscopy) image from a healthy patient.

Adapted from www.optometrystudents.com (B) Fundoscopy image from a patient with

ONH. Note the arrows defining the edge of the optic disc. Adapted from Spandau et al.,

2002.

A B

28 Figure 1.2. Structure of the human and mouse eye (A) Schematic cross-section of human and mouse eye. While they are overall very similar, human eyes contain a fovea and a macula which the mouse lacks. The lens of the mouse eye is much larger in comparison to total globe size and has a more circular structure. (B) Mouse retinal section with H&E staining for nuclear cell layers. Adapted from Veleri et al., 2015.

Figure 1.3. Retinal cell populations and birthdates. (A) Schematic of retinal cell types and their corresponding position in three nuclear layers.

(Black: Retinal Ganglion Cells, Green: Amacrine Cells, Gray: Bipolar Cells, Pink: Muller Glial

Cells, Blue: Horizontal Cells, Orange: Cone Photoreceptors, Red: Rod Photoreceptors) (ONL-

Outer Nuclear Layer, INL- Inner Nuclear Layer, GCL- Ganglion Cell Layer). (B) Map of developmental timeline for cells born from Retinal Neural Progenitor Cells.

32 Figure 1.4. CASK binding partners and proposed functions (A) Schematic domain organization of CASK. Blue proteins listed are known or proposed binding partners of each domain. (B) Schematic of a tripartite synapse. Red – Presynaptic Neuron.

Orange – Postsynaptic Neuron. Green – Astrocyte. Lines indicate different regions where CASK is known or proposed to have some function.

34 Figure 1.5. Proposed functions of CASK at synapse Due to CASK’s ability to interact with many different proteins, CASK has been hypothesized to have many different functions at the synapse. This schematic of a synapse represents these varying roles that occur at many different regions.

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52 2. Optic Nerve Hypoplasia is a pervasive subcortical pathology of visual system in neonates

Authors: Chen Liang1.2,5, Alicia Kerr1,3,5, Yangfengzhong Qui1, Francesca Cristofoli4, Hilde Van Esch4, Michael Andrew Fox1,2, and Konark Mukherjee1,2

1Developmental and Translational Neurobiology Center, Virginia Tech Carilion Research Institute, Roanoke, Virginia, United States 2Department of Biological Sciences, Virginia Tech, Blacksburg, Virginia, United States 3Graduate Program in Translational Biology, Medicine, and Health, Blacksburg, Virginia, United States 4Center for Human Genetics, University Hospitals Leuven, Leuven, Belgium 5Co-first author *Correspondence: [email protected]

This chapter was published October 2017 in Investigative Ophthalmology and Visual Science. https://iovs.arvojournals.org/article.aspx?articleid=2659950

53 2.1. Abstract Optic nerve hypoplasia (ONH) is the most common cause of childhood congenital blindness in developed nations. Despite the high prevalence and increasing incidence of

ONH, the mechanisms underlying its pathogenesis, timing of onset and exact nature of pathology remain unclear. Mutations in the X-linked gene CASK have been associated with ONH clinically. In our current study we confirm the genetic association by documenting that heterozygous loss of function mutation in CASK gene in females may associate with ONH. In order to investigate the effect of CASK gene haploinsuffciency on optic nerve (ON), we examined the effect of heterozygous deletion of CASK gene (Cask+/-

) using a rodent model. We demonstrate that CASK haploinsufficiency is sufficient to produce ONH with complete penetrance. Optic nerves in Cask+/- mice are thin, comprised of atrophic axons and display reactive astrogliosis. Using 3-D reconstructed ultrastructural analysis we demonstrate that the ON axons form aberrant synaptic terminals.

Furthermore we observe a significant loss of retinal ganglion cells also. Importantly, the onset of pathology is early in postnatal life of mice, which corresponds to third trimester of human fetal development. A common molecular underpinning and a similar pathology suggest that Cask+/- mice is a valid model of ONH. Our data therefore indicate that disruption of CASK mediated signaling produces ONH, a subcortical pathology of the visual system.

2.2. Introduction

The prevalence of childhood blindness (~0.7/1000) is nearly a tenth of that of adult blindness, however due to early affliction it poses a huge socioeconomic challenge (Rahi et al, 1999). The loss of vision in childhood may be due to toxins, trauma, infection or genetic factors. Many cases of congenital blindness are associated with defects in the optic nerve. Optic nerve hypoplasia (ONH) is the most common form of optic nerve dysfunction and may account for as much as 15-25% of childhood blindness in developed nations (Kaur et al., 2013). Surprisingly, the degree of ONH may not be directly correlated with visual acuity (Kaur et al., 2013) and ONH may be present with near normal vision.

ONH may be associated with other ophthalmological manifestations like microphthalmia, aniridia, nystagmus and strabismus (Zeki et al., 1991). Typically, ONH is a stable condition with no age dependent deterioration and is diagnosed clinically by fundoscopy, which reveals a small pale optic disc. In most cases, the ratio of disc diameter to macula/disc diameter is also increased in ONH (Zeki et al., 1991). In some cases, the optic disc may also appear in funduscopic images to be surrounded by double rings (Frisen and

Holmegaard, 1978). However, mild cases of ONH are often more difficult to diagnose.

Although the first case of ONH was described in 1884 then there has recently been an increase in awareness and understanding of this disorder and at present it is classified into three distinct forms 1) optic nerve hypoplasia simplex, 2) septo-optic dysplasia (SOD) and 3) septo-optic-pituitary dysplasia (Kaur et al., 2013).

55 The precise etiopathogenesis of ONH and its differences from perinatal optic nerve atrophy (ONA) remains unclear (Hoyt and Good, 1992). Maternal age and primiparity are the most consistent risk factors associated with ONH (Garcia-Filion and Borchert, 2013).

Although ONH has been observed in a murine model of fetal alcohol syndrome (Parson et al., 1995), large scale studies have failed to uncover evidence for association of ONH with maternal consumption of alcohol and other recreational drugs (Garcia Filion and

Borchert, 2013). Most cases of ONH are sporadic in nature, with only few genes that have been directly implicated. For instance, mutations in HESX1 may be associated with SOD in limited number of cases (Dattani et al., 1998; Tajima et al., 2003). However, for the majority of cases of ONH no genetic cause has been defined and there is both a paucity of genotype-phenotype correlation of patients with ONH (Garcia-Filion and Borchert,

2013) and a lack of appropriate animal models with which to study ONH to gain mechanistic insight into the pathogenesis of this debilitating disease. Therefore, it is essential to develop novel models of ONH by using known genetic associations from

GWAS studies. Recently mutations in the X-linked gene CASK have been strongly associated with ONH (Moog et al., 2011) and mutations in CASK therefore represent a potential avenue to generate a novel model of ONH for mechanistic evaluation.

CASK was initially described as a candidate gene for X-linked ONA (Dimitratos et al., 1998) and mutations in CASK are associated with ONH (Moog et al., 2011; Moog et al., 2015; Burglen et al., 2012), however, roles for CASK in retinal and optic nerve formation have not been investigated. Here, we report a new subject with CASK mutation and ONH, further confirming the association in humans, and we show that heterozygous deletion of CASK recapitulates many of the phenotypes observed in humans indicating

56 that these phenotypes represent CASK loss of function. Furthermore, our results demonstrate that fewer and thinner retinal axons are present in the optic nerve of Cask+/- mice, and that there is increased interaxonal space and astrogliosis. The numbers of retinogeniculate synapses are reduced with specific loss of smaller boutons; furthermore the large boutons exhibit an overall decrease in number of active zones. In retina, we observe a decrease in number of retinal ganglion cells in Cask+/- mice. Finally, we demonstrate that the optic nerve pathology in Cask+/- mice occurs very early in postnatal rodent developmental, a stage considered equivalent to the late third trimester and neonatal period of human fetal development (Dobbing and Sands, 1979).

Haploinsufficiency of CASK therefore produces a combination of retinopathy, optic nerve pathology as well as synaptopathy. Thus a genetically validated mouse model provides crucial information on the pathogenesis and timing of ONH.

2.3. Results 2.3.1. Mutations in the CASK gene are associated with ONH Mutations in CASK gene have been previously associated with ONH, ONA and glaucoma (Moog et al., 2011; Burglen et al., 2012). Here we describe a 3-year-old girl with a heterozygous splice site mutation in CASK gene

(NM_001126054:exon24:c.2233+1G>A) leading to skipping of exon 24 that exhibits microcephaly, global developmental delays, and spastic quadriparesis (Figure 2.1.A-C).

Magnetic resonance imaging did not reveal any specific midline structural defect as seen in septo-optic dysplasia or any white matter lesion (Figure 2.1.D-E). Although, truncated proteins may be generated in the subject from the mutated CASK allele, they are unlikely

57 to be functional in absence of the Hook motif and the guanylate kinase domain, especially since the interdomain interaction with the guanylate kinase domain is critical for proper folding and function of MAGUK proteins (Tavares et al., 2001; Li et al., 2014).

Furthermore, the transcripts with premature stop codon are likely to get degraded due to non-sense mediated decay. Thus the pathogenic mutation described here is likely to produce haploinsufficiency. The girl displays ONH and other ophthalmic conditions including hyperopia, nystagmus, and infantile esotropia. Fundoscopy revealed a bilaterally normal anterior segment, however both optic discs were pale, small and hypoplastic (Figure 2.1.F). Hypoplastic optic nerve has also been observed in MRI scans of another girl with heterozygous nonsense mutation in the N-terminal kinase domain of

CASK (personal communication). These observations indicate that heterozygous loss of function mutations of CASK gene commonly associates with ONH.

2.3.2. Heterozygous deletion of CASK in mice produces optic nerve pathology

To test whether heterozygous mutation of CASK is sufficient to alter the ON and subcortical visual system in mice with complete penetrance, we analyzed female mouse mutants with a single allele of CASK deleted (Cask+/- mice). Previously, we documented that Cask+/- mice exhibit secondary microcephaly, disproportionate cerebellar hypoplasia as well as thinning and hypoplasia of the optic nerve indicating that it recapitulates the human phenotypes associated with CASK mutation or loss (Srivastava et al., 2016). We examined optic nerves following toluidine blue staining of semithin cross-sections – a method that labels myelin and allows the quantification of myelinated retinal axons in the

58 ON. Surprisingly besides being small (Fig. 2.2.A-B), the optic nerve displayed an overall decrease in axonal density (Fig. 2.2.C-D). The number of axons in a given area was reduced by ~25% and the interaxonal space was expanded (Fig. 2.2E). Histopathology on a single optic nerve hypoplasia case suggested that there may be an increase in number of astrocytes in ONH (Saadati et al., 1998). We therefore examined the optic nerve of Cask+/- mice for levels of glial fibrillary acidic protein (GFAP), a marker of astrocytes. We found a large increase in the GFAP staining in the optic nerve of Cask+/- mice compared to sex-matched littermate controls suggesting that haploinsufficiency of

CASK gene may be associated with increased astrocytes in optic nerve (Fig. 2.2F).

2.3.3. Axonopathy is present in optic nerves of Cask+/- mice A reduced axonal density in the Cask+/- mice optic nerve may reflect not only a loss of RGC axons, but also a thinning of individual axons of RGCs. We therefore performed transmission electron microscopy on cross sections of ON and quantified axonal area in

Cask+/- mutants (Fig. 2.3.A-B). Our data indicate that the cross-sectioned area of individual axons was significantly decreased in Cask+/- mice (Fig. 2.3.C). Since optic nerves contain populations of coarse and fine retinal axons (Guillery et al., 1982; Williams et al., 1983), we measured the area of coarse and fine axons separately. Our data demonstrate that the area of both the types of axons are decreased in Cask+/- mice (Fig.

2.3.D-E).

We next turned our attention to myelin content in mutant ON, to assess whether reduction in ON diameter reflects myelin loss. G-ratio distribution spectra, which assess the ratio of axonal diameter to the diameter of the myelinated axon (Guy et al., 1989) were

59 applied to compare myelin of wild type littermate mice to that of the Cask+/- mice. The probability density distribution of G-ratios in mutant mice appears shifted to the left, indicating a higher myelin-to-axon ration compared to controls (Fig. 2.3.F). We interpret these results to indicate that while CASK haploinsufficiency influences axon diameter it does not appear to limit the development of compact myelin on these axons.

Because CASK is an X-linked gene and is therefore subject to X-linked random inactivation, two types of RGCs exist in Cask+/- mutant mice, those which produce CASK and those which did not. Differences in ON diameter in CASK+ and CASK- RGCs may produce an increased variability in axon diameter in these studies. We therefore measured the mean of deviation of the axonal areas. Surprisingly, the mean deviation for the axonal area in Cask+/- optic nerve is smaller than the wildtype (data not shown).

Overall, the distribution of the area of individual axons was smaller but roughly parallel in

Cask+/- mice and their sex-matched wild type littermate indicating random inactivation of deleted CASK allele had no bearing on the development or health of individual axons (Fig.

2.7.). We have previously documented that CASK promotes postnatal brain growth in a non-cell-autonomous manner (Srivastava et al., 2016), and consistent with these findings the distribution of axonopathy in Cask+/- mice did not correlate with expression of CASK in individual cells. One possible explanation for this observation may be that there is specific loss of only CASK negative axons in Cask+/- mice. However, specific loss of axons from cells which are CASK negative will lead to an artificial skewing of RGCs towards being CASK positive. To determine if such is the case, we blotted optic nerve homogenates for CASK and an axonal marker, neurofilament associated antigen (NFAA)

(Fig. 2.3.G). Our data suggest that there is a slight reduction in NFAA which is consistent

60 with a decrease in axonal density; however the CASK content within optic nerve was

~47% indicating survival of CASK negative axons (Fig. 2.3.H). Our data thus suggest that

ONH in Cask+/- mice may not be due to specific loss of CASK negative axons.

2.3.4. Retinogeniculate synaptopathy is observed in Cask+/- mice Axonopathy may alter axonal transport and produce defects in synapse formation

(Sterman and Sposito, 1984). CASK has also been considered to be a trafficking molecule

(Wei et al., 2011) and a synaptic scaffolding protein and it is known to interact and phosphorylate presynaptic adhesion molecule neurexin in an activity dependent manner

(Mukherjee et al., 2008, Mukherjee et al., 2010). Neurexins themselves are thought to be synaptogenic (Scheiffele et al., 2000). We therefore examined retinogeniculate synapses.

It is pertinent to note in previous animal model studies involving worms, flies and mouse, deletion of CASK did not produce defects in synapse formation (Atasoy et al., 2007;

Slawson et al., 2011; Hoskins et al., 1996). We initially examined RGC terminals in thalamus by anterogradely labeling RGCs with intraocular injection of fluorescently conjugated cholera toxin subunit B. We found that there is a significant reduction in small diameter RGC terminals in Cask+/- mice compared to wildtype littermate (Fig. 2.4.A).

Because of this change in terminal populations we examined the ultrastructure of RGC terminals using serial block face scanning electron microscopy (Wei et al., 2011). With this technique, retinal terminals can be unequivocally identified based on their ultrastructural morphology (Zhen et al., 1999). Synapse volume in Cask+/- mice remained largely unchanged (Fig. 2.4.B-D), however smaller terminals have significant reduction in volume (Fig. 2.7.; Fig. 2.8.). In order to test if the defect associated with CASK haploinsufficiency is restricted to smaller synapses only or not, we carefully

61 analyzed the ultrastructure of all reconstructed RGC terminals. Our data demonstrate that the number of active zones (release sites) per unit volume of RGC terminals are reduced in Cask+/- (Fig. 2.4.E-F). Thus, besides the thinning of axons produced by CASK haploinsufficiency we also observed defects in retinogeniculate synapse morphology and ultrastructure.

2.3.5. RGC numbers are reduced in Cask+/- mice Since we detected both a thinner optic nerve and a lowered axonal density in optic nerve of Cask+/- mice, we next examined if there was also a loss of retinal integrity or

RGCs in Cask+/- mice. Immunolabeling mutant and control retinal cross-sections with antibodies against Calretinin (Calr) (Fig. 2.5.A), revealed that the general cytoarchitecture and laminated structure of the retina was preserved in the absence of CASK. To assess whether there was a specific loss of retinal ganglion cells in Cask+/- mice we counted the total number of cells within the ganglion cell layer of the retina (by labeling nuclei with

DAPI) and the total number of cells expressing RBPMS, an RNA binding protein selectively expressed by RGCs in the rodent retina (Anjum et al., 2014) (Fig. 2.5.A).

Together these data reveal a significant loss of RGCs in Cask+/- mutants (Fig. 2.5.A-B).

Taken together, we find that ONH in Cask+/- mice is a complex developmental disorder involving a decrease in the number of RGCs, thinning of individual axons, reactive astrogliosis within the ON, and alterations in retinogeniculate synapses. Thus, mutation of CASK appears to phenocopy many of the features observed in human patients with

ONH and therefore represents an appropriate mouse model to elucidate the mechanisms underlying ONH.

2.3.6. ONH develops postnatally in Cask+/- mice The first mechanistic question regarding ONH pathogenesis that has eluded our understanding is when exactly the disease is initiated. The decrease in RGCs and their axons could occur pre- or postnatally. While it is difficult to parse these differences out in humans, in this mouse model the difference can be studied by observing ON diameter throughout development. We therefore examined toluidine blue stained semi-thin section of Cask+/- and control optic nerves during the first three weeks of postnatal mouse development (Fig. 2.6.A). It is important to point out that the first two postnatal weeks of rodent development correspond to developmental milestones that occur in the third trimester of human fetal development (Dobbing and Sands, 1979). Moreover, neonatal mice do not open their eyes until the end of the second postnatal week of development.

Surprisingly, the ON of P1 Cask+/- mice was of similar size to that of the wildtype controls

(although, a slight trend towards being smaller was observed). A more clear and significant difference between the size optic nerves became apparent by the end of the first week of postnatal development and the difference from the wildtype reaches the difference observed in adults by P22 (Fig. 2.6.B). Although myelinogenesis in mouse start around P5 it is most intense between the second and fourth week after birth (Dangata and Kaufman, 1997) at P22 myelinated axons are easily quantifiable. To confirm that the decrease in axonal density also occurs at an early age and not as a part of later degeneration we examined the number of axons in P22 optic nerve per unit area. Our data indicate that axonal density is already reduced (at par as noted in adults Fig. 2.2.C) by this stage suggesting that CASK-linked ONH can clearly be classified as a developmental and not a degenerative process. Thus, our data demonstrate that ONH in

63 Cask+/- mice occurs early in development, prior to the full maturation of the subcortical visual system.

2.4. Discussion ONH is the commonest cause of childhood blindness in developed nations, and its prevalence is growing at an alarming rate (Hackett et al., 2010). A validated genetic animal model of ONH recapitulating the clinical condition with high fidelity will serve not only as a necessary tool to understand the mechanism of disease pathogenesis, but also serve to test therapeutic interventions in the future. ONH may occur with midline defects as observed in SOD, with other brain malformations like atrophic brain or even as isolated condition (Garcia-Filion and Borchert, 2013). The co-incidence of ONH with other brain malformation is not surprising given that the retina and optic nerves are a part of our central nervous system and subject to the same developmental processes as the brain.

Therefore, a clear understanding of ONH may also help unlock the pathophysiology of other neurodevelopmental disorders. In fact, ONH has been considered as a type of pervasive disorder and subjects with ONH display a high incidence of autistic phenotype such as stereotypy (Ek et al., 2005; Parr et al., 2010). In this regard mutations in CASK gene as a causal factor for ONH is particularly important since CASK mutations are also known to produce autistic traits (Hackett et al., 2010). CASK directly binds and phosphorylates neuronal adhesion molecule neurexin (Mukherjee et al., 2008, Mukherjee et al., 2010). It is pertinent to note that mutations in both neurexins and their trans-synaptic interacting partners also are associated with autism (Cao and Tabuchi, 2017). Mutations in the CASK gene are also associated with postnatal microcephaly (Srivastava et al.,

64 2016). It is possible to argue that the ONH may simply be a part of microcephaly. However other monogenic syndromes associated with postnatal microcephaly such as Rett syndrome or Angelman syndrome does not present with ONH, indicating that ONH

(Michieletto et al., 2011, Saunders et al., 1995) is an independent manifestation of CASK mutation.

CASK is a membrane associated guanylate kinase protein which interacts with a large number of other molecules. Specifically, CASK has been considered to be a synaptic scaffolding molecule both at the pre and post-synaptic compartments (Hu et al.,

2016; Samuels et al., 2007). We have recently made two independent observations, 1)

CASK stabilizes neuronal adhesion molecule neurexin and links it to the signaling molecule liprin-α (LaConte et al., 2016). Liprin-αs are critical for photoreceptor axonal targeting in drosophila (Hofmeyer et al., 2006) as well as for forming active zones (Zhen and Jin, 1999). Neurexins themselves may play a critical role in intercellular adhesion and signaling (Graf et al., 2004). A reduction in neurexin signaling and defect in axonal targeting may produce ONH, secondary to defect at the retinogeniculate synapses due to retrograde degeneration (Mosier et al., 1978). 2) We also found that CASK may regulate cellular metabolism including mitochondrial respiration (Srivastava et al., 2016). Retina and optic nerve are highly susceptible to metabolic defects, and mitochondrial damage, and mutations in mitochondrial genes are frequently associated with optic neuropathies

(Carelli et al., 2004). In fact as many as 12% cases of non-syndromic mitochondrial cytopathies may display ONH (Taban et al., 2006). Thus a change in metabolic status of retinal cells may also lead to ONH in Cask+/- mice and CASK haploinsufficient girls.

65 Mutations in CASK have been associated with diagnosis of both ONH and ONA.

The difference between these two conditions is often blurred and it has been speculated that better imaging techniques may differentiate these conditions. Many authorities even suggest that both ONH and ONA may be similar pathology with only difference being the timing (Hoyt and Good, 1992). What does our study reveal regarding the pathology of optic nerve? First of all, we present a case report of CASK haploinsufficiency displaying

ONH, confirming the genetic associations. Secondly, using a rodent model we are able to causally link ONH with CASK haploinsufficiency. Previous rodent model of ONH using alcohol produced thinning of optic nerve only during adulthood (after 9 weeks) (Parson et al., 1995). In contrast, significant decrease in optic nerve area in in Cask+/- mice was observed as early as P6 which corresponds to late third trimester in human fetal development (Dobbing and Sands, 1979). The time period of optic nerve pathology in

Cask+/- mice is indeed developmental in nature and is therefore a superior rodent model of ONH. Several histopathological hallmarks of atrophy however are present in ONH such as astrogliosis and thinning of axons. A detailed examination of optic nerve, retina and dLGN in Cask+/- mice undermine the idea that simply better imaging tool may be able to distinguish between ONH and ONA in human patients, these optic neuropathies may not be amenable to straightforward binary classification simply by morphology. We suggest that mutations in CASK is typically associated with ONH, the diagnosis of ONA may simply stem from the timing when originally uncovered. In fact, many infants that are diagnosed with ONA may actually have ONH which was not diagnosed earlier.

Overall, the evidence presented in this study argues that haploinsufficiency of

CASK is causative of ONH and therefore Cask+/- mice serve as a good model for ONH.

66 In the future careful biochemical and physiological analysis of Cask+/- mice will shed light on the precise mechanism of the highly prevalent optic nerve disorder, ONH. Finally, although CASK haploinsufficiency causes ONH, depending on nature (mis-sense) and timing (embryonic/somatic) of CASK mutation it is possible that the optic neuropathy may develop later in life presenting as ONA or even glaucoma. Careful experimentation with ablation of CASK from different cell populations in a time dependent manner will further clarify the role of CASK in subcortical visual pathway formation and maintenance.

2.5. Experimental Procedures 2.5.1. Identification of a splice mutation in CASK We performed whole exome-sequencing (WES) on genomic DNA of the patient.

Using the Illumina HiSeq2000 platform (Illumina, San Diego, CA, USA) and the acquired reads were aligned to the reference human genome (UCSC Genome Browser hg19). Data processing was performed with the Genome Analysis ToolKit (GATK) and variants were annotated using Annovar and an in-house developed web interface called Annotateit. In a first stage, we analyzed all genes that were associated with microcephaly before. The heterozygous splice variant in exon 24 (NM_001126054:exon24:c.2233+1G>A) of

CASK, was predicted to lead to exon skipping, and was confirmed by Sanger sequencing.

The variant arose de novo and was absent in both biological parents. To further test the effect of the mutation, we performed a PCR with primers in exons 25 and 23 on cDNA made from RNA extracted from EBV transformed lymphocytes of the patient. The RTPCR on the patient clearly showed two bands and sequencing of the smallest band confirmed exon 24 skipping.

2.5.2. Mouse breeding and genotyping Cask+/- female mice, generated at VTCRI (Srivastava et al., 2016), were crossed with C57BL6 male mice to propagate Cask+/- female pups and their Cask+/+ female litter mates. Genotyping was done using polymerase chain reaction (PCR) based method with following primer pair (forward: tttggggactagatgggtgtggtg, reverse: cttggtcgcagcttgggagta).

2.5.3. Immunohistochemistry Mice were anesthetized and perfused with PBS and then 4% paraformaldehyde

(PFA). Brain, retina or optic nerve were dissected and postfixed in 4% PFA. Tissues were cryopreserved in 30% sucrose before embedding in Neg50 matrix and cryosectioned using Leica Cryostat to generate 25 μm-thick tissue sections. Sections were permeabilized and blocked with 0.25% TritonX-100 and 5% BSA in PBS. Immunostaining were performed using primary antibodies at an appropriate concentration followed by labeling with fluorophore conjugated secondary antibodies. Primary antibodies include:

Calretinin (Rabbit polyclonal Ab, Millipore, 1:2000), RBPMS (Rabbit polyclonal Ab,

PhosphoSolution, 1:500), GFAP (Neuromab, 1:1000). Secondary antibodies include:

Alexa Flour 488 (Goat anti-rabbit, Invitrogen, 1:1000) and Dylight 633 (anti-mouse IgG

Thermo Scientific). Sections were mounted with Vectashield containing DAPI (Vector labs) and were imaged using a Zeiss 700 or 710 laser scanning confocal microscope.

68 2.5.4. Immunoblotting

Tissue samples were boiled in sample buffer and proteins separated by 8%

SDSPAGE. Proteins were then transferred onto nitrocellulose membrane and blocked in

5% skimmed milk for 2 hours and incubated using appropriate primary and secondary antibodies. For quantitative immunoblots, secondary antibodies conjugated with fluorophore were used and images were captured using appropriate filter in ChemidocTM

(Biorad). Antibodies used are CASK (1:1000, NeuroMab), Neurofilament-associated antigen (NFAA/3A10, 1:1000, DSHB) and tubulin (DSHB, 1:1000).

2.5.5. Optic nerve samples preparation for toluidine blue staining and transmission electron microscopy (TEM) Mice were anesthetized and perfused with PBS and then sodium cacodylate

(NaCa) buffer (2% glutaraldehyde/2% PFA (2G/2PF) in 0.1M sodium cacodylate in PBS).

After perfusion, optic nerves were dissected and immersed in NaCa buffer overnight and then put in 2% OsO4 for 2 hours for post fixation. Dehydration was achieved by sequentially adding 15%-100% ethanol. Samples were embedded in 100% Epoxy resin for 24 hours and then placed in oven at 58°C for 2 days. Sectioning of samples and successive toluidine blue stain or TEM was performed at Virginia-Maryland College of

Veterinary Medicine at Virginia Tech.

2.5.6. G-Ratio distribution spectra

G-ratio, defined as the ratio of inner axonal diameter to the outer axonal diameter

(including myelin sheath), of 200 axons in the ON were measured on TEM images

69 (×4000) using ImageJ software (http://imagej.nih.gov/ij/; provided in the public domain by the National Institutes of Health, Bethesda, MD, USA). The measurements of inner and outer axonal diameters (d) were calculated from inner and outer axonal areas (a), respectively, to reduce error because the sections were not perfectly round in shape, where d ≈ 2 √ (a/π).

2.5.7. Serial block-face scanning electron microscopy Control and CASK+/- mice were transcardially perfused with PBS and 4% paraformaldehyde / 2% glutaradehyde in 0.1M cacodylate buffer. Brains were removed,

300 µm coronal sections were obtained using vibratome and dLGN were dissected.

Tissues were stained, embedded, sectioned and imaged by Renovo Neural Inc.

(Cleveland, OH). Images were acquired at a resolution of 6 nm/pixel and image sets included > 200 serial sections (with each section representing 65 nm in the z axis). Data sets were analyzed in TrakEM2 (Cardona et al. 2012) and retinal terminals were unequivocally identified by the presence of synaptic vesicles and pale mitochondria

(Mukherjee et al., 2016; Hammer et al., 2015).

2.5.8. Intraocular injections of anterograde tracers and quantification Intraocular injection of cholera toxin subunit B (CTB) conjugated to AlexaFluor 555

(Invitrogen) was performed as described previously (Su et al., 2011). After 2 days, mice were euthanized, perfused with 4% PFA and brains were dissected and postfixed in 4%

PFA overnight. 80–100µm coronal slices were sectioned on a vibratome and mounted

70 using VectaShield (Vector Laboratories, Burlingame, CA, USA). Confocal Z-stack images from coronal dLGN sections were acquired on a Zeiss LSM 700 confocal microscope

(Oberkochen, Germany). Z-stacks were obtained with a Zeiss 20x Plan-Apochromat objective and contained 14-20 optical sections (in 3µm steps). To quantify CTB-labeled retinal terminals in dLGN images were obtained from 4 adult Cask+/- mutants and 4 littermate controls. The size and quantity of synapses in each dLGN (from the center of its rostral-caudal extent), was performed in ImageJ, blind to the genotype of the sample.

Five images of each dLGN Z-stacks were randomly selected, images were manually thresholded, and the quantity and surface area of the isolated puncta was measured.

Data was exported into Microsoft excel, binned by size, and plotted into a histogram.

2.5.9. Cell counts Cell counts were performed on 16 μm cryosectioned 4% PFA-fixed retinal tissue as described previously (Fox et al., 2007, Su et al., 2012). Slides were stained with

DAPI

(diluted 1:5000 in PBS) for 60 seconds and then mounted with VectaShield (Vector

Laboratories, Burlingame, CA, USA). Images of central retina were acquired on a Zeiss

LSM 700 confocal microscope (Oberkochen, Germany) or a Zeiss Axio Imager A2 fluorescent microscope. All DAPI and RBPMS-positive cells in the ganglion cell layer were counted manually under 20x objective. Cell counts were analyzed from at least 4 animals for each age and genotype. Five images per retina per mouse were analyzed.

71 2.5.10. Description of coarse and fine axons Since the axons in Cask+/- mice were overall small, it is difficult to apply the same threshold for fine and coarse axons on these mice as with those of wildtype. On examining the distribution of the axonal area we found they are distributed in a bimodal manner (two different slopes) (Figure 2.7.). Since the Cask+/- axons were smaller we applied a different threshold for each genotype, the value of the threshold was the point at which the two slopes intersected.

Acknowledgements The current study is supported by R01EY024712 from NEI to KM. The Fox laboratory is supported by RO1AI124677 and RO1EY021222 to MAF. Jeffery Willis and

Helen Clark provided technical help. We gratefully acknowledge the participation and cooperation of families of children with CASK mutation in our studies.

Figure 2.1. Genetic, brain and retinal alterations associated with heterozygous

CASK mutation

A) Sanger sequences showing the de novo heterozygous splice variant in exon 24

(NM_001126054:exon24:c.2233+1G>A), B) The schematic of exon skipping C) CASK.

RT- PCR with primers (arrows) in exons 25 and 23 on cDNA of the patient and subsequent sequencing confirmed exon 24 skipping. D, E) MRI images of brain indicate no specific midline structural defect or white matter lesion. F) A fundoscopic image from this haploinsufficient 3-year-old girl who was diagnosed with ONH. Note the pale, hypoplastic optic disc.

73 A NC_000023.10:g.41393958C>T NM_001126054:c.2233+1G>A NP0036792:pGly741His768delinsAsp

Proband Mother Father

B C * Ex1 Ex23 Ex24 Ex25 Ex27

exon s kipped

D E F

Figure 2.2. Cask+/- ON display reduced axons and increased astrogliosis

A and B) representative images of semithin cross section of optic nerves from indicated genotype stained with toluidine blue. Scale bar = 100 μm. C and D) higher magnification of toluidine blue stained semithin optic nerve sections of indicated genotype. Note that

Cask+/- optic nerve has lesser axonal density and increased interaxonal space. E)

Axonal density from optic nerve of 4 different mice is quantified, data is plotted as mean±SEM. (*: p<0.05). F and H) Panels showing fluorescence microscopy of cryosections of optic nerve of indicated genotypes. Green panel represents fluorophore tagged cholera toxin B subunit (CTB-alexa488), red panel represents immunostaining with GFAP. Scale bar = 200 μm. G and I are higher magnification image from the same optic nerve indicating increased GFAP staining in Cask+/- optic nerve.

76 Figure 2.3. Cask+/- ON display axonal thinning

A and B) Representative transmission electron micrograph of optic nerve thin sections from indicated genotype. C) Quantitation of axonal area from anonymized images obtained from 3 mice in each group (+/+ and +/-), data is plotted as mean±SD; N=60. D and E) Comparisons of fine and coarse axons area changes from anonymized images obtained from 3 mice in each group (+/+ and +/-), data is plotted as mean±SD; N=60. F)

G ratio of myelinated axons, inner and outer axonal diameter were calculated from inner and outer axonal area in order to reduce error since the sections were not perfectly round in shape, data is plotted as mean±SD; N=60. G) Immunoblot of optic nerves from six randomly selected animals of indicated genotype. The antigens are indicated, NFAA is neurofilament associated antigen which is axonal marker. H) Quantitation of the blots against tubulin, the wild-type amount is considered to be 1.0. Data is plotted as mean

±SEM, n=3. (*: p<0.05; **: p<0.01).

78 Figure 2.4. Cask+/- mice display a decreased number of retinogeniculate synapses and a reduced number of active zones in the remaining synapses.

A) Quantification of the area of terminals in the shell of the dLGN by anterograde labeling of CTB with showing of representative images of dLGN of mice of indicated genotype injected with an anterograde fluorescent tracer (CTB-Alexa) to label the retinal nerve endings. Data represents mean±SEM, n=4. B and C) Representative image of SBFSEM ultramicrograph with a retinogeniculate presynapse (identified by presence of pale mitochondria) labeled red and a postsynapse labeled green. Examples of reconstructed retinal terminals from a series of SBFSEM micrographs are shown on the right. D, E and

F) Volumetric quantitation (D), number of active zones (E) and active zone density (F) of the presynaptic bouton, 60 reconstructed presynapses from SBFSEM analysis of three different mice were used. Data represent mean±SEM, n=3. (*: p<0.05; **: p<0.01).

80 Figure 2.5. Decreased number of retinal ganglion cells in Cask+/- mice.

A) Immunostaining of retina with antibodies against calretinin reveals normal lamination in Cask+/- mutants at P26. Representative images of retinas stained with DAPI and

RNABinding Protein with Multiple Splicing (RBPMS) reveal a significant loss of RGCs in

Cask+/- mutants compared to controls. Scale bar = 50 μm. B) Quantition of cells in the retinal ganglion layer in indicated staining. (*: p<0.05).

82 Figure 2.6. Cask+/- mice optic nerve display secondary reduction in size A)

Representative images of toluidine blue stained semithin-sections derived from optic nerve of indicated age and genotypes (P=postnatal). Scale bars = 100 μm. B) Quantitation of optic nerve areas from mice of indicated age and genotype, data is plotted as mean±SEM, n=3. C) Quantitation of axonal density, data is plotted as mean ±SD. (*: p<0.05; **: p<0.01)

84 Figure 2.7. Supplemental Figure 1

Ophthalmoscopy (fundoscopy) image from a control 3.5 year old girl. Note the bright, well defined optic disc.

86 Figure 2.8. Supplemental Figure 2

Mean deviation of optic nerve area is plotted for both genotypes. Mean deviation is a measure of variation and calculated with following formula: mean deviation = Σ(x-µ)/N, where x represents each value, µ is the mean and N=number of values.

88 Figure 2.9. Supplemental Figure 3 Retinal lamination remains the same among genotypes. (A) Representative images of retinas taken with confocal microscope. These images were imported to Image J, lines were drawn for the Inner Plexiform Layer (IPL), shown with red lines, as well as the

Inner Nuclear Layer (INL), shown with yellow lines. Lengths of red and yellow lines were quantified for each image (several locations per image, n = 5 mice per genotype). (B)

Quantification of width for IPL and INL. Data are shown as mean ± SEM.

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95 3. Non-cell autonomous roles for CASK in optic nerve development.

Alicia Kerr1,2, Paras Patel1,2, Leslie LaConte1,3, Chen Liang1, Ching-Kang Chen4, Veeral Shah4,5, Michael Andrew Fox1,6,7*, Konark Mukherjee1,8*

1Developmental Translational Neurobiology Center, Fralin Biomedical Research Institute at Virginia Tech Carilion, Roanoke, Virginia, United States 2Graduate Program in Translational Biology, Medicine, and Health, Virginia Tech, Blacksburg, United States 3Department of Basic Science Education, Virginia Tech Carilion School of Medicine, Roanoke, Virginia, United States 4Department of Ophthalmology, Baylor College of Medicine, Houston, Texas, United States 5Texas Children’s Hospital, Houston, Texas, United States 6Department of Biological Sciences, Virginia Tech, Blacksburg, Virginia, United States 7Department of Pediatrics, Virginia Tech Carilion School of Medicine, Roanoke, Virginia, United States 8Department of Psychiatry and Behavioral Medicine, Virginia Tech Carilion School of Medicine, Roanoke, Virginia, United States

*Correspondence to: [email protected] and [email protected]

This chapter was submitted to Investigative Ophthalmology and Visual Science, March 2019.

96 3.1. Abstract Heterozygous mutations in the essential X-linked gene CASK are associated with

Optic Nerve Hypoplasia (ONH) and other retinal disorders in girls. Cask+/- heterozygous mice exhibit ONH with a loss of retinal ganglion cells (RGCs) without any changes in retinal lamination or morphology. It remains unclear if reduction in CASK expression selectively affects RGCs or also involves other retinal cells. Furthermore, it is not known if CASK expression in RGCs is critical for optic nerve (ON) development and maintenance. The visual behavior of CASK+/- mice was assessed and electroretinography

(ERG) was performed. We also examined the ophthalmological condition of a boy with a hemizygous CASK partial loss-of-function (LOF) mutation. Using a mouse line with a floxed CASK gene that expresses ~40% CASK (hypomorph) under hemizygous and homozygous conditions, we investigated effects of CASK reduction on the retina and ON.

Finally CASK was completely deleted from RGCs to examine the CASK’s role specifically in RGCs. CASK+/- heterozygous mutant mice display abnormal performance in visual behavior tasks, but ERG is indistinguishable from wild type. The boy with a CASK sequence variation (p.Pro673Leu) exhibits isolated ONH with unremarkable retina. CASK hypomorph mice exhibit ONH, but deletion of CASK from RGCs in this background does not further exacerbate the condition. Our results demonstrate that mosaic or global reduction in CASK expression and/or function disproportionately affects RGCs. CASK expression in RGCs does not appear critical for cell survival, indicating a non-cell autonomous role for CASK in the development of ON.

97 3.2. Introduction Optic Nerve Hypoplasia (ONH) is the most common cause of childhood blindness in developed nations, and its incidence is on the rise (Khaper et al., 2017; Kong et al.,

2012). ONH involves thinning of the optic nerve (ON) and is typically associated with loss of retinal ganglion cells (RGCs) and their axons (Katagiri et al., 2017). Most cases of ONH are non-genetic in nature, and the etiogenesis of ONH has therefore remained obscure.

Genetically identified forms of ONH are typically associated with transcription factors that are directly involved in the development of RGCs (Chen et al., 2017). However, in many instances of ONH, RGCs initially begin to develop and connect with the brain, but development stalls or RGCs undergo atrophy before complete ON development (Frisen et al., 1978; Hoyt and Good, 1992). Therefore, to better understand the etiopathogenesis of ONH, face-validated animal models of this disease need to be investigated.

We have demonstrated that haploinsufficiency of the X-linked gene CASK

(Calcium/calmodulin Activated Serine Kinase) produces ONH in both humans and mice

(Liang et al., 2017). The protein product of this gene (also CASK) is a peripheral scaffolding protein (Butz et al., 1998) and does not directly regulate RGC development, thus providing a model to examine the etiogenesis of ONH. In addition to ONH, mutations in CASK are also associated with other retinopathies, including retinal dystrophy, indicating that retinal cells distal to RGCs may also be affected (Burglen et al., 2012;

LaConte et al., 2019; Moog et al., 1993). We did not observe any change in lamination or loss of non-RGC retinal cells in previous work (Liang et al., 2017). Here, we investigate the effect of CASK haploinsufficiency on the function of retinal cells and determine if

CASK deficiency produces an RGC-specific pathology.

98 Our results demonstrate that CASK haploinsufficiency produces isolated RGC pathology. Furthermore, using a human case report and a CASK-hypomorph mouse line, we demonstrate that any reduction in CASK expression/function is likely to damage RGCs and produce ONH. Surprisingly, we find that despite this disproportionate sensitivity of

RGCs to CASK deficiency, CASK expression in an RGC is not important for its survival or maintenance. Taken together these results suggest that the RGC pathology observed with CASK loss is likely to be non-cell autonomous in nature.

3.3. Results 3.3.1. Heterozygous loss of CASK produces behavioral but not electroretinographical deficits in vision We previously described that heterozygous loss of Cask (Cask+/-) in mice results in ONH (Liang et al., 2017). To assess visual function in Cask+/- mice, we used a twoalternative forced-swim task. In this task, mice learn to associate a visual cue of a sinegrating with a hidden platform used to escape the water (Fig. 3.1.A) (Huberman and

Niell, 2011; Prusky et al., 2000). Mice genetically devoid of RGCs are unable to perform this task (Monavarfeshani et al., 2018). We first assessed the ability of Cask+/- mice (and female wild-type littermates) to learn this task. Cask+/- mice and controls were trained for

8 days on a vertical grating (0.17 cycles per degree [cpd]) on the S+ monitor above a hidden platform, (compared to a gray screen on the S- monitor). When their ability to locate the hidden platform in ten trials (per day) exceeded 70%, they were considered to have successfully discriminated between the visual cues (Prusky et al., 2000). Cask+/- mice and female littermate controls performed at similar rates through the 8 days of

99 training, and by the end of this training both genotypes were performing at near 100% accuracy (Fig. 3.1.B).

By changing the spatial frequency of the S+ sine-grating, we were able to test visual acuity of mutant and control mice. Although Cask+/- mice performed slightly worse than littermate controls, there was no significant difference in performance regardless of the spatial frequency of the gratings (Fig. 3.1.C). We then altered the contrast of the gratings. Cask+/- hypomorphs performed statistically worse than littermate controls in these tasks, and their performance consistently worsened (compared with littermates) as the contrast of the grating decreased (Fig. 3.1.D). Thus in addition to ONH and reduced

RGCs, Cask+/- hypomorphs exhibit readily noticeable decreased visual performance.

To assess whether this behavioral deficit was due to ONH (and RGC loss) or from defects within the retina, electoretinograms (ERGs) were recorded on Cask+/- mice and controls. ERG waveforms are well-documented to reflect neural activity from the outer retina, with the a-wave reflecting photoreceptor activity (Brown, 1968) and the b-wave reflecting activity of the depolarizing bipolar cells (Gurevich et al., 1993; Stockton et al.,

1989). Under photopic conditions, ERG b-wave responses appeared similar between

CASK+/- and wild-type littermates (Fig. 3.1.E). Moreover, the average photopic b-wave amplitudes elicited by light flashes of 25 Cd/sec/m2 in intensity showed no difference between genotypes (Fig. 3.1.F). Likewise, scotopic a-wave amplitudes were also comparable between Cask+/- and wild-type littermates (Fig. 3.1G-H). By fitting the ensemble scotopic b-wave amplitudes vs. retinal illuminance to a modified Naka-Rushton function for maximum rod- and cone-driven responses and their half-saturating light intensities (Li et al., 2005), we further found their ERG responses to be very similar (Fig.

100 3.1.I). Taken together, these results indicate that retinal function is normal in Cask+/- mice and thus suggest that the observed deficit in visual behavior is likely due to the loss of

RGCs and their axonal thinning.

3.3.2. Partial loss of function of CASK in a 3-year-old boy produces ONH CASK loss in males usually leads to lethality or neurodevastation with epileptic encephalopathies (Burglen et al., 2012; Saitsu et al., 2012). We report a 3-year-old boy with a milder condition exhibiting microcephaly and severe pontocerebellar hypoplasia

(MICPCH) similar to haploinsufficient females (Fig. 3.2.A) and ON thinning (Fig. 3.2.B).

This subject presented with decreased visual acuity (20/130; Teller acuity test) and horizontal nystagmus (with a pendular jerk of small amplitude and frequency) that did not dampen with convergence, indicating that it may be central in origin. These phenotypes

(thin ON, non-retinal defects) led to the diagnosis of ONH. Exome sequencing uncovered a CASK variant c.2018C>T (p.Pro673Leu) that was absent from the parents and had not been identified or reported in any other database.

We expressed recombinant CASK containing the P673L variation (CASKP673L), fused with GFP protein for biochemical analysis. Wild-type CASK (CASKWT) expressed in

HEK293 cells diffusely fills the cytosol (Fig. 3.2.C), whereas expression of CASKP673L results in an aggregation of protein (Fig. 3.2.D), indicating that this protein may be partially misfolded or prone to aggregation compared to wild-type. CASKP673L also exhibits higher insolubility, supporting the notion that this protein variant may have a propensity to misfold

(Fig. 3.2.E).

101 We next employed molecular dynamics (MD) simulations to characterize the structure of CASKP673L. Three 100 ns MD simulation trajectories were run using a model of the CASK wild-type and CASKP673L PDZ-SH3-GK (PSG) supradomain (LaConte et al.,

2018). Analysis of the simulations reveals that CASKP673L’s radius of gyration (2.51 nm) over the course of the three trajectories is slightly, significantly larger than CASKWT (2.48 nm) and explores a larger range of radii throughout the simulations (Supplemental Figure

1). A comparison (Fig. 3.2.F) of representative CASKWT and CASKP673L structures (from a cluster analysis of the three MD trajectories) reveals that the PDZ domain of CASKP673L exhibits a larger RMSD difference (3.1nm) from the wild-type structure than seen with other variations within the PSG supradomain (e.g., CASKM519T RMSD difference is 2.49 nm; CASKG659D RMSD difference is 2.11 nm), an important observation given CASK’s known interactions with proteins containing PDZ-binding domains (Fairless et al., 2008;

Hata et al., 1996; Mukherjee et al., 2008). When comparing secondary structure propensity and mobility at each residue (b-factors) throughout the MD trajectories, of particular interest is the notable absence in helical propensity and mobility in the α-C helix of the CASKP673L PDZ domain (Zeng et al., 2018), a region purported to be responsible for the coupling between the PDZ and SH3 domains necessary for ligand binding. The

P673L variation in CASK’s SH3 domain disrupts packing between all three of the domains, causing a reorientation of the domains with respect to each other. The GK domain is rotated approximately 45 degrees with respect to the PDZ-SH3 face that it interacts with

(Fig. 3.2.F). This disruption in packing may indicate that CASKP673L is more likely to adopt a conformation that oligomerizes. Several differences in the PDZ domain are induced by the P673L variant (note the loop between the first two beta strands and the orientation of

102 the alpha helix between beta strands 5 and 6), but the most substantial difference is in the α-C “helix” of the PDZ domain (which retains no helical form in CASKP673L) and a consequent shift in the interface between the SH3 domain and the PDZ domain. In sum, these structural differences point to a CASKP673L structure which is slightly less compact than its wild-type counterpart and has alterations in the region predicted to interact with known binding partners such as neurexin (Mukherjee et al.,

2008; Biederer et al., 2001; LaConte et al., 2016; Pak et al., 2015; Biederer et al., 2002;

Cohen et al., 1998). We therefore tested whether the structural changes predicted by MD in CASKP673L indeed impairs CASK-neurexin binding by using a previously described recruitment assay where distribution of GFP-CASK can be altered from cytoplasmic to membrane-bound upon co-expression of neurexin-1β (Fig. 3.2.C; Fig. 3.2.G). For this recruitment assay, analysis was done only on cells where GFP-CASKP673L was not aggregating. Upon co-expressing neurexin-1β, we did not observe co-localization of

P673L CASK and neurexin-1β on the membrane (Fig. 3.2.H), revealing that neurexin is unable to efficiently recruit the remaining soluble fraction of CASKP673L from the cytosol.

In order to quantitatively measure the change in CASKP673L affinity towards neurexin, we performed GST pulldown assays using the cytosolic tail of neurexin-1 fused to the GST protein. Our results indicate that the affinity of CASKP673L for neurexins is reduced by

~80% compared to CASKWT (Fig. 3.2.I-J).

Overall, these results suggest that the P673L sequence variation affects the structure of CASK sufficiently to increase misfolding and impair its interaction with

103 neurexins. Such a reduction in functional CASK is sufficient to produce ONH manifestation, indicating RGCs may be highly sensitive to CASK deficiency.

3.3.3. Global reduction in CASK expression produces ONH Due to random X-inactivation, CASK heterozygous mutations result in a mosaic condition, with ~50% of brain cells generating 100% of CASK and ~50% of cells being

CASK-null. This mosaicism is likely to be a confounding variable in our experiments designed to understand the cellular role of CASK in RGCs and ON development. A global

CASK-LOF mutation, either hemizygous (male) or homozygous (female), is a useful model to help disentangle this confound. Cask-/- mice do not survive, however a conditional allele of CASK was previously generated (Caskfl/fl) which was shown to cause a global reduction in CASK expression due to selection cassette interference (Atasoy et al., 2007). This Caskfl/fl mouse has also been reported to have microcephaly and cerebellar hypoplasia (Srivastava et al., 2016; Najm et al., 2008).

Here, we show similarly reduced levels of CASK in the ON of Caskfl/fl mice (Fig.

3.3.A) and therefore used these mice to explore if uniform reduction in CASK impacts the developing visual system. Toluidine blue sections show that Caskfl/fl ONs are thin and hypoplastic compared to littermate CASK+/+ controls (Fig 3.3.B). Thus, global reduction of

CASK in all cells results in ONH. We also found that mice heterozygous for the floxed allele (Caskfl/+) have a slight but significant decrease in ON size compared to controls (Fig.

3.3.C). Next, we evaluated RGC numbers in Caskfl/fl, Caskfl/+, and controls using a marker for RGCs (RBPMS) (Rodriguez et al., 2014). We saw a dose-dependent decrease in

RGCs in the presence of the floxed allele of CASK (Fig. 3.3.D): one floxed allele (Caskfl/+)

104 significantly reduced RGC numbers compared to controls (Cask+/+), and homozygous floxed alleles (Caskfl/fl) significantly reduced RGC numbers compared to both heterozygotes (Caskfl/+) and controls (Cask+/+) (Fig. 3.3.E). Finally, we explored potential axonopathy in Caskfl/fl mice. RGC axon number and diameter was evaluated using TEM of ON cross sections from Caskfl/fl and control mice (Fig. 3.3.F). No reduction in the density of axons in Caskfl/fl samples was observed compared to Cask+/+ ONs (Fig. 3.3.G),

However we observed a significant decrease in the average cross-sectional area of axons

fl/fl 2 in Cask ON (WT 1.54 ± 0.70 µm , CASK 1.20 ± 0.52 µm2, p < 0.05). RGC axons in the

ON can be classified as fine or coarse, based on diameter (Guillery et al., 1982; Williams and Chalupa, 1983). Previous analysis in Cask+/- hypomorph mice revealed a loss of both fine and coarse RGC axons. We plotted RGC axon diameter and could identify the

fl/fl 2 inflection point between axon types in Cask and controls (Fig 3.3.H; 2.208 µm in controls). By separating the fine and coarse axons and averaging axon size for each type, we observed that both fine (Fig. 3.3.G) and coarse axons (Fig. 3.3.H) were significantly reduced in Caskfl/fl ON. Myelination patterns were evaluated in TEM images, and Caskfl/fl

ON myelination is apparently normal in adulthood. However, analysis of the

ON at P12 showed a difference in total myelinated axon numbers compared to Cask+/+

(Fig. 3.7.). This difference was no longer apparent by P18, suggesting that global reduction in CASK delays the onset of myelination. Overall, our data suggest RGCs are extremely sensitive to level of CASK expression

105 3.3.4. CASK is expressed in retinal ganglion cells To analyze the role of CASK in RGC development, we first asked if RGCs express detectable levels of CASK using in situ hybridization (Fig. 3.4.A). Riboprobes against

Cask mRNA were developed and revealed widespread expression in all three nuclear layers (Fig. 3.4.B); no appreciable reactivity in these regions (versus synaptic regions) was observed with a sense riboprobe (Fig. 3.4.C). Importantly, we observed significant reactivity of the CASK antisense riboprobe in the ganglion cell layer (GCL), suggesting

CASK is generated by RGCs. Results were validated by showing CASK immunoreactivity in the GCL using IHC (Fig. 3.4.D).

3.3.5. RGC-derived CASK is not required for RGC survival To specifically remove CASK from RGCs we sought a driver line that generated

Cre-recombinase in RGCs to excise floxed-Cask allele. One option was (Calretinin-Cre

(Calb2-Cre)). The percentage of RGCs in which Calb2-Cre exhibited recombination activity was quantified by crossing it to a reporter line (Rosa-stop-tDT Ai9). As expected, widespread expression of tdT was observed in the retinas of Calb2-Cre::Rosa-stop-tdT mice (Fig. 3.5.A, A’). Cross-section analysis showed that tdT expression is present in both the inner nuclear layer (INL) and GCL (Fig. 3.5.B) (77 ± 2.5% of DAPI-labeled cells in the

GCL were tdT+; n=4 mice). Since misplaced amacrine cells also reside in the GCL and may also express Cre in this driver line (Fig. 3.5.C), we used immunostaining for RBPMS to assess the percent of RGCs expressing Cre in this line (Fig. 3.5.C’). We found 91.1 ±

2.4% (n=4 mice) of RBPMS cells were tdT+ in Calb2-Cre::Rosa-stop-tdT. Furthermore

+ 87.6 ± 5.1% of the tdT cells were immunoreactive for RBPMS.

106 To assess the requirement for CASK in RGCs, Caskfl/fl::Calb2-Cre+/+ mice were generated. These mutants have a global reduction in CASK expression (as described above) as well as a complete loss of CASK in ~91% of RGCs. We compared the ON of

Caskfl/fl::Calb2-Cre+/+ mice with Caskfl/fl mice using toluidine blue staining as described earlier (Figure 6A). Surprisingly, there was no further reduction in the size of ON when

CASK was nearly abolished from RGCs (Fig. 3.6.B). Furthermore, when RGC’s were labeled with RBPMS, no significant loss of RGCs in the Caskfl/fl::Calb2-Cre+/+ mice was observed as compared to Caskfl/fl, suggesting that loss of CASK expression from RGCs does not affect RGC survival (Fig. 3.6.D). Finally, axonal morphology in the ON of these mutants (Fig. 3.6.E) was observed; there were no differences in RGC number or morphology of Caskfl/fl::Calb2-Cre+/+ or Caskfl/fl mice (Fig. 3.6.F-I).

3.4. Discussion Although the prevalence of ONH is on the rise, the mechanism that underlies its onset and progression remains unknown. ONH is most likely a heterogeneous disorder with differing etiologies that may share similar downstream pathogenesis. Many cases of

ONH arise much later in perinatal and postnatal development and are associated with environmental factors (Hoyt and Billson, 1986; Lambert et al., 1987). The mechanism of this later-onset ONH is not clear. Mouse models which target genes involved in RGC development, however, often lead to complete non-development of eyes (anophthalmy)

(Taranova et al., 2006) or of the ON itself (Brown et al., 2001) and thus do not recapitulate

ONH. Heterozygous deletion of CASK produces ONH and microcephaly in girls and female mice (Liang et al., 2017; Burglen et al., 2012; Srivastava et al., 2016; Moog et al.,

107 2011). We have shown that microcephaly linked to CASK mutation may stem from loss of PDZ domain-mediated interactions with proteins like neurexin (LaConte et al., 2016).

Analysis of a CASK variant (CASKP673L) in a male subject with microcephaly and ONH suggests both microcephaly and ONH are likely to result from disruption of PDZ domainmediated interactions of CASK.

Intriguingly, CASK mutations have also been associated with retinopathies

(Burglen et al., 2012; LaConte et al., 2019). These observations pose an interesting question: do CASK mutations produce isolated ONH or are the ON pathologies reflective of a broader retinal disorder? This question is not only critical for a better classification of the ocular phenotypes seen with CASK mutations but is also crucial for the validity of

CASK mutant mice as tools to investigate mechanisms underlying ONH.

Cask+/- mice display a postnatal decrease in the number of RGCs, thinning of the

ON, and atrophy of RGC axons (Liang et al., 2017). Despite deficits in RGCs, no defects were observed in other cells or lamination in the Cask+/- retina (Liang et al., 2017). Cask+/- mice also exhibit significantly decreased contrast sensitivity. Based on retinal morphology and ERG data, we suggest that this visual deficit is due to RGC pathology and not due to defects in retinal circuitry. Thus, CASK haploinsufficiency specifically affects RGCs to produce ONH. Furthermore, we demonstrate that global reduction of CASK expression by ~60% is also sufficient to induce ONH, indicating that RGCs are extremely sensitive to

CASK loss. Why are RGCs disproportionately affected by CASK loss? The anatomy and physiology of RGCs are unique; they are the predominant action potential-generating neurons within the retina and have extremely long, myelinated axons unlike other retinal

108 cells. In other brain regions, Cask loss or mutation appears to disproportionately affect projection neurons, such as those in pontocerebellar circuits (Burglen et al., 2012).

Since the CASK hypomorph line is based on a floxed CASK allele, we also used this line to delete all CASK expression from RGCs. Surprisingly, total loss of CASK from

RGCs did not exacerbate the ONH phenotype, indicating RGC-derived CASK is not critical for RGC survival. How then does CASK loss produce ONH? There are at least two possible explanations. First, although the discernible pathology of ONH appears late in the course of development, it is possible that the biological insult required for ONH occurs earlier in development before the Cre-mediated deletion of CASK gene. A second, more plausible explanation may be that the effect seen on RGCs is non-cell-autonomous in origin. A strong argument for this possibility is that in Cask+/- mice, we did not observe secondary selection (apoptosis) of neurons resulting from random X-linked inactivation, suggesting that neurons lacking CASK do not display reduced survival (Liang et al., 2017;

Srivastava et al., 2016).

CASK is present not only in neurons but also in glial and endothelial cells (Horng et al., 2017; Weigand et al., 2012). Aberrant functioning of these cell populations is likely to affect survival and health of RGCs by producing changes in retinal metabolism and vascularization. In fact altered vasculature, such as vascular tortuosity, has been suggested to be one of the hallmarks of ONH (Lambert et al., 1987; Cheng et al., 2015;

Kaur et al., 2013), strongly suggesting a common underlying mechanism for the development of ONH that can be investigated by in-depth study of CASK mutations.

109 3.5. Experimental Procedures 3.5.1. Visual behavior tasks

Two alternative forced swim tasks were performed in a trapezoid shaped pool

(sides a = 25 cm, b = 80 cm, c and d = 143 cm) with two side-by-side monitors (19 inches,

V196L, Acer) placed at the wide end (b) of the tank and separated by a black divider (42 cm). Detailed instructions for the apparatus were described previously (Prusky et al.,

2000). A rescue platform (37 cm ×13 cm × 14 cm) was hidden under water below the monitor with the positive visual cue (termed the S+ side). Visual cues (i.e. different grating pattern) were generated in the Gabor-patch generator

(https://www.cogsci.nl/gaborgenerator). The visual cue and hidden platform were moved to the right or left screens in a pseudorandom manner with the following orders:

LRLLRLRLRR, RLRRLRLRLL, RRLRLLRLRL and LLRLRRLRLR. During the behavioral tasks the room was dark, but a 60 W bulb was positioned above the holding cages. During the visual tasks, mice were held in separate cages placed on to heating pads and lined with paper towels. A day before starting experiments, mice were acclimated to the experimenter and the pool through handling, a 1–2 min period of direct contact with the hidden platform at either arms, and submersion into the water at gradually-increasing distances from the hidden platform. Behavioral tasks included a training phase (8 days) and a testing phase (10–12 days). For training phases, mice were placed at the center of the narrow end and given one minute to find the platform for 8–10 trials per day. A trial was recorded as a correct choice if a mouse passed the choice line on the S+ side, while passing the choice line on the S- side was recorded as an incorrect choice. After arriving at the rescue platform, mice were placed back into their individual cages. When a mouse made an incorrect choice, it was placed back at the release chute to perform another trial

110 immediately before going back to its home cage. After 8 days of training, mice learned to find the positive visual cue (i.e. vertical gratings) with a >80% accuracy. To test visual acuity and contrast sensitivity, we increased the spatial frequency and decreased the contrast of vertical gratings (i.e. the S+ cue), respectively. In the testing phase of the detection tasks, 10 trials of a given task (e.g. detection of vertical gratings with spatial frequency of 0.32 cpd versus a gray screen) were performed in 10 consecutive days (one per day). No more than nine animals were tested in a given session. Each mouse performed no more than 10 trials per day. A total of 10 CASK+/+ and 11 CASK+/- were tested. Mice were 2-6 months old at the start of training.

3.5.2. Electroretinography Three female Cask+/- mice and three wild type female mice were dark-adapted overnight and anesthetized under infrared illumination by ketamine/xylazine/acepromazine through intraperitoneal injection (94/5/1 mg/kg), and pupils were dilated with eye drops containing 1% tropicamide and 2.5% phenylephrine

(Bausch & Lomb, Tampa, FL). Body temperature was maintained at 35~37 °C by 43.5 °C circulating water through a plastic heating coil wrapped around the animals.

Stimulusdependent transcorneal potential changes from both eyes were simultaneously recorded using the UTAS BigShot system (LKC Technologies, Gaithersburg, MD). A series of calibrated white flashes with intensities ranging from 0.0000025 to 250 Candela sec m−2 were used as stimuli (Li et al., 2005). The interstimulus interval was 3 sec for dimmer flashes and increased to 120 sec for brighter flashes to ensure proper dark adaptation.

111 Responses from 5 to 15 independent measurements were averaged for analysis.

Photopic recordings ensued immediately after scotopic recordings by exposing the animals to a white background light of 30 Candela m−2 intensity for 10 min followed by flashes of 25 Candela sec m−2 in intensity and presented at 1 Hz in frequncy for 90 sec.

Ensemble scotopic ERG b-wave amplitude vs. retinal illuminance (Fig. 1I) was fitted to a modified Naka-Rushton function: A(x)=[R*x/(x+A)] + [C*x/(x+B)], where R and C are rod- and cone-driven maximum response amplitudes (in µV), respectively, and A and B are half-saturating light intensities for rod- and cone-driven responses, respectively.

3.5.3. Plasmid and point mutagenesis

pEGFP-C3-CASK (rat) plasmid has been previously described (Mukherjee et al.,

2008). Using Phusion polymerase (NEB), point mutation P673L was generated in the pEGFP-C3-CASK plasmid by Site-directed mutagenesis and sequenced in the Core

Laboratory Facility in the Virginia Bioinformatics Institute at Virginia Tech.

3.5.4. Cell culture and imaging

Human embryonic kidney (HEK-293) cells (ATCC) were plated on 50 µg/ml polyL-lysine (Sigma Aldrich Inc.)-coated coverslips (Fisherbrand, Inc.) in 24-well plates

(JetBiofil) and maintained in DMEM (Hyclone) containing 10% fetal bovine serum

(Hyclone) supplemented with 5 mg/ml penicillin-streptomycin (Hyclone). Cells at 80% confluency were transfected with 0.5 µg of GFP-CASKWT and GFP-CASKP673L DNA per well using calcium phosphate method. Twenty hours post-transfection, cells were washed

112 twice with phosphate buffered saline (Sigma Inc.) and fixed for 15 minutes at room temperature using a 4% paraformaldehyde solution. Coverslips were mounted on microscope slides (Premiere) using Vectashield (Vector Laboratories Inc.) and visualized using confocal laser scanning microscopy (ZEISS Axio Examiner.Z1 LSM 710).

3.5.5. Molecular dynamics simulations and analysis

The UCSF Chimera software package (Pettersen et al., 2004) was used for molecular visualization and editing. Molecular dynamics simulations were performed using the program GROMACS 5.1.3 (Abraham et al., 2015) on a previously constructed homology model of CASK’s PDZ-SH3-GuK (PSG) supradomain (LaConte et al., 2018) and the corresponding P673L mutant, which was constructed using with Chimera’s

Rotamers tool to replace the native proline with leucine at the position analogous to 673 with a leucine rotamer from the Dunbrack backbone-dependent rotamer library with the highest probability. The AMBER99SB-ILDN force field (Lindorff-Larsen et al., 2010) was used for all simulations. Structures were solvated with a three-point water model (SPC/E) in an explicit rhombic dodecahedron water box (solute box distance of 1.0 nm) under periodic boundary conditions, with charges neutralized by chloride ions. Structures were then minimized and 100 ps NVT and NPT equilibration simulations were performed as previously described (LaConte et al., 2018). An unrestrained 100 ns NPT molecular dynamics simulation was run after equilibration. Three trajectories initiated with different random seeds were generated for both the wildtype and CASKP673L structures.

113 After post-processing to correct for periodicity, trajectories were used to calculate root mean square deviation (RMSD) and radius of gyration (Rg) of the protein backbone from the starting structure at each time point using the GROMACS rms and gyrate command, respectively, and a Welch’s two-sample t-test (chosen due to unequal variance as evaluated by Levene’s test) was run in R to evaluate differences in Rg between

CASKWT and CASKP673L. The root mean square fluctuation (RMSF) of all alpha carbons from each of the three trajectories was also calculated (GROMACS rmsf command).

RMSF values were used to calculate B-factors for each residue, which were then

2 2 averaged, using the equation: B-factor = (8∏ /3) x (RMSF) . Average representative structures were selected based on a cluster analysis of the three concatenated molecular dynamics trajectories (excluding the first 25 ns of each trajectory) for the wildtype and

P673L PSG structures, respectively, that was performed using the GROMACS cluster command with a 3Å cutoff employing the gromacs algorithm (Daura et al., 1999).

Secondary structure prediction to evaluate helical propensity was performed by using

GROMACS to call the dssp program (Touw et al., 2015); DSSP output was then analyzed using a Python program.

3.5.6. Neurexin mediated cell recruitment assay

Human embryonic kidney (HEK-293) cells (ATCC) were plated on 50 µg/ml polyL-lysine (Sigma Aldrich Inc.)-coated coverslips (Fisherbrand, Inc.) in 24-well plates

(JetBiofil) and maintained in DMEM (Hyclone) containing 10% fetal bovine serum

(Hyclone) supplemented with 5 mg/ml penicillin-streptomycin (Hyclone). Cells at 80%

114 confluency were co-transfected with 0.5 µg of Flag tagged Neurexin1-β and GFPCASKWT or GFP-CASKP673L DNA per well using calcium phosphate method. Twenty hours post- transfection, cells were washed twice with phosphate buffered saline (Sigma Inc.) and fixed for 15 minutes at room temperature using a 4% paraformaldehyde solution. Cells were permeabilized using PBS with 0.01% Triton-X 100 solution and blocked using 5% fetal bovine serum. Cells were immunostained using monoclonal antiFLAG antibody

(1:100) followed by Alexa-633(1:250) anti-mouse antibody. Similar to the mounting and imaging procedure described above.

3.5.7. Solubility assay

Human embryonic kidney (HEK-293) cells (ATCC) were plated in 6-well plates

(JetBiofil) and maintained in DMEM (Hyclone) containing 10% fetal bovine serum

(Hyclone) supplemented with 5 mg/ml penicillin-streptomycin (Hyclone). Cells at 80% confluency were transfected with 10 µg of EGFPN1, GFP-CASKWT or GFPCASKP673L

DNA per well using calcium phosphate method. Two days later cells were harvested and solubilized in RIPA buffer containing PBS, 1%Triton X-100, 0.5% sodium deoxycholate,

2mM EDTA, 1mM EGTA and protease inhibitors (0.1 mg/ml aprotinin, 0.1 mg/ml leupeptin, 0.1 mg/ml pepstatin, and 0.01 mg/ml PMSF; all purchased from Fisher

Scientific). Pellets and supernatant were obtained after centrifuging lysates for 30 min. at

15,000 rpm at 4°C. Pellets and supernatants were boiled with SDS sample buffer, proteins were separated using SDS PAGE and blotted with CASK antibody.

115 3.5.8. Pull down assay using NXCT and GST beads

Human embryonic kidney (HEK-293) cells (ATCC) were plated in 6-well plates

(JetBiofil) and maintained in DMEM (Hyclone) containing 10% fetal bovine serum

(Hyclone) supplemented with 5 mg/ml penicillin-streptomycin (Hyclone). Cells at 80% confluency were transfected with 10 µg of EGFPN1, GFP-CASKWT or GFPCASKP673L

DNA per well using calcium phosphate method. Two days later cells were harvested and solubilized in RIPA buffer containing PBS, 1%Triton X-100, 0.5% sodium deoxycholate,

2mM EDTA, 1mM EGTA and protease inhibitors (0.1 mg/ml aprotinin, 0.1 mg/ml leupeptin, 0.1 mg/ml pepstatin, and 0.01 mg/ml PMSF; all purchased from Fisher

Scientific). The lysates were incubated by rocking at 4°C for 30min and then centrifuged for 30 min. at 15,000 rpm at 4°C to collect the supernatants. The precleared supernatants were incubated with 30ul of GST (glutathione-s-transferase) and NxCT GST sepharose beads and incubated on rocker for 2hr 4°C. The beads were washed three times with PBS containing 1% Triton X 100 buffer. 30 μl of 2x SDS sample buffer was added to the beads and boiled 100C for 10min. Protein samples were separated using SDS-Page and immunoblotted for CASK.

3.5.9. Immunoblots and immunohistochemistry

Proteins were resolved using SDS-PAGE (sample volumes loaded were based on estimates of appropriate protein concentration), transferred to a nitrocellulose membrane

(Fisher), and immunoblotted for the proteins of interest. HRP-conjugated secondary antibody were used. The blots were developed using Amersham ECL western blotting detection reagents (GE Healthcare Life Sciences) for HRP secondaries. All blots were

116 imaged using a ChemiDoc™ MP System (BioRad). Antibodies used are CASK (mouse monoclonal Ab, Cat. No. 75000, 1:1000; NeuroMab), Synaptophysin (polyclonal, Cat. No.

336A-76, 1:1000, Sigma), β-Actin (mouse monoclobal Ab, Cat. No. MABT825 clone 4C2 lot# Q2653600, 1:1000, Sigma).

Mice were anesthetized and perfused with PBS and then 4% paraformaldehyde

(PFA). Brain and retina were dissected and postfixed in 4% PFA. Tissues were

cryopreserved in 30% sucrose before embedding in tissue freezing medium (Electron

Microscopy Studies, Hatfield, PA) and cryosectioned using Leica CM1850 Cryostat

(Leica Biosystems, Wetzlar, Germany) to generate 16-μm thick tissue sections for brain.

Sections or tissue were permeabilized and blocked with 0.25% Triton X-100 and 5%

BSA in PBS. Immunostaining was performed using primary antibodies at an appropriate concentration followed by labeling with fluorophore conjugated secondary antibodies.

Primary antibodies include: RNA-binding protein with multiple splicing (RBPMS; rabbit polyclonal Ab, Cat. No. 1830, 1:500; PhosphoSolution, Aurora, CO, USA), CASK

(Mouse IgG1, 1:100, Clone K56A/50, UC Davis/NIH NeuroMab Facility, Davis, CA). For

IHC with anti-CASK antibodies tissues were fixed in 100% Methanol. Secondary antibodies include: Alexa Fluor 488 (goat anti-rabbit, Cat. No. A-11008, 1:1000;

Invitrogen) and

Alexa Fluor 555 (goat anti-mouse IgG, Cat. No. A32727, 1:1000; Thermo Scientific,

Waltham, MA, USA). Sections were mounted with VectaShield containing 4′,6- diamidino2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA, USA) and were imaged using a Zeiss 700 or 710 laser scanning confocal microscope (Oberkochen,

Germany).

117

3.5.10. Optic nerve samples preparation for transmission electron microscopy Mice were anesthetized and perfused with PBS and then 0.1 sodium cacodylate

(NaCa) buffer containing 2% glutaraldehyde/2% PFA [2G/2PF] pH 7.4. Optic nerves were dissected and immersed in NaCa buffer overnight and then put in 2% OSO4 for 2 hours for postfixation. Dehydration was achieved by sequentially adding 15% to 100% ethanol.

Samples were embedded in 100% epoxy resin and cured at 58°C. Sectioning of samples for TEM was performed at Virginia-Maryland College of Veterinary Medicine at Virginia

Tech.

3.5.11. Axon counts

Axons were counted on 4000X and 8000X TEM images using syGlass (Pidhorsky et al., 2018) virtual reality software and Oculus rift hardware (Best Buy). Images were uploaded into syGlass as TIFFs with voxel dimension of 1x1x1 nm. In spyglass, each image was optimized for quantification by reducing the stride and optimizing image intensity. The counting feature of syGlass was used to mark every axon in the field of view.

3.5.12. Optic nerve area Toluidine blue images were imaged on a Zeiss AxioImager A2 fluorescent microscope at 20X objective. Images were uploaded to ImageJ and the freehand tool was used to circle the optic nerve.

118 3.5.13. G Ratio

G-ratio, defined as the ratio of inner axonal diameter to the outer axonal diameter

(including myelin sheath), of 200 axons in the ON were measured on TEM images (×4000) using ImageJ software (http://imagej.nih.gov/ij/; provided in the public domain by the

National Institutes of Health, Bethesda, MD, USA). The measurements of inner and outer axonal diameters (d) were calculated from inner and outer axonal areas (a), respectively, to reduce error because the sections were not perfectly round in shape, where d ≈ 2 √

(a/π).

3.5.14. In situ hybridization (ISH)

ISH was performed on 16 μm sections as previously described (Su et al., 2016;

Su et al., 2010). Sense and antisense riboprobes were generated against full-length

CASK IMAGE Clone (catalog number MMM1013-202761641, Dharmacon, Lafayette,

CO). Riboprobes were synthesized using digoxigenin (DIG) (Roche, Mannheim,

Germany) and the MAXI-Script In Vitro Transcription Kit (Ambion, Austin, TX). Probes were hydrolyzed to ≈500 base pairs. Images were obtained on a Zeiss LSM700 confocal microscope.

3.5.15. Mouse breeding and genotyping CASK(+/−) female mice generated in-house (Srivastava et al., 2016) were crossed with C57BL6 male mice to generate CASK(+/−) female pups and their CASK(+/+) female littermates. Calb2-Cre (stock #010774), Rosa-stop-tdT (Ai9) (stock #007909), CASKfl/fl

(stock #006382) were all obtained from Jackson Laboratories. Genotyping was done

119 using a PCR-based method with following primer pairs ([CASK] forward:

TTTGGGGACTAGATGGGTGTGGTG, reverse: CTTGGTCGCAGCTTGGGAGTA;

[CASK floxed] forward: GTCGCAGCTTGGGAGTAGAG, reverse:

ACTAACCCTCCTCCCTTTCG; [Cre] forward: CGTACTGACGGTGGGAGAAT, reverse:

TGCATGATCTCCGGTATTGA); [tdT] forward ACC TGG TGG AGT TCA AGA CCA TCT, reverse: TTG ATG ACG GCC ATG TTG TTG TCC).

3.5.16. RGC Counts

Cell counts were performed on 16-μm cryosectioned 4% PFA- fixed retinal tissue as described previously (Liang et al., 2017). Slides were stained with DAPI (diluted 1:5000 in PBS) for 60 seconds and then mounted with VectaShield (Vector Laboratories). Images of central retina were acquired on a Zeiss LSM 700 confocal microscope or a Zeiss Axio

Imager A2 fluorescent microscope. All RBPMS-positive cells in the ganglion cell layer were counted manually under 320 objective. Cell counts were analyzed from at least four animals for each age and genotype. Five images per retina per mouse were analyzed.

Acknowledgements The authors thank the participation and cooperation of families and children with

CASK mutation in our studies. We thank Vrushali Chavan for technical help. Ching-Kang

Chen is the Alice McPherson Retina Research Foundation Endowed Chair at Baylor

College of Medicine and is supported by grants from the National Institute of Health

(EY026930 and EY002520) R01EY024712 (KM). Work in the Fox and Mukherjee

120 laboratories is supported by the National Institutes of Health (EY029874 [KM], AI124677

[MAF] and EY021222 [MAF]).

121

Figure 3.1. Heterozygous loss of CASK produces behavioral but not electrophysiological deficits in vision.

(A) Schematic of two-alternative forced-swim test with a choice between a sine- positive grating (S+) and a sine-negative grating (S-). (B) Cask+/- and wild-type

littermates learn the task at a similar rate. (C) Increasing the spatial frequency of sine-

grating leads to similar decrease in hypomorph and control performance. (D)

Hypomorphs perform significantly worse than controls when contrast (%luminence) was

decreased. (STATS) (E-H) Electroretinography in both photopic (E) and scotopic

conditions (G). N=3 mice per genotype. There were no statistical differences in photopic

b-wave amplitudes (F) or scotopic a-wave amplitudes (H). A Naka-Rushton fit of scotopic b-wave amplitudes vs. retinal illuminance for maximum rod- and cone-driven responses reveal comparable fits between Cask+/+ and Cask+/- mice (I). For all panels, data is plotted as mean±SEM. * indicates p<0.05, ** indicates P<0.01 by two-way ANOVA.

122

123 Figure 3.2. Partial loss-of function of CASK in 3-year-old boy produces ONH

(A-B) CASK variant in a 3-year-old boy with microcephaly and severe pontocerebellar hypoplasia with CASK variant c.2018C>T (p.Pro673Leu) mutation. T2-weighted magnetic brain resonance images show small cerebellum (black arrow) and thinning of optic nerve

(red arrow). (C) Expression of WT-CASK-GFP fusion protein in HEK293 cells shows localization of CASK in cytosol. (D) CASKP673L-GFP fusion protein shows cotton-wool aggregates (white arrows). (E) Blot for CASK in soluble and soluble fractions of cell lysate show that CASKP673L is predominantly present in the insoluble fraction. (F) Structural models of the PSG supradomain of CASKWT (left) and CASKP673L (right). Site of variation shown in orange, PDZ domain in purple, a-C helix in green, SH3 domain in yellow, hook/hinge in salmon, and GK domain in gray. Arrows indicate structural differences between wild-type and variant structures. (G) Recruitment assay using a known

WT CASKsubstrate neurexin-1β shows that CASK colocalizes with neurexin in cytosol. (H)

CASKP673L is mislocalized, with some colocalization with neurexin. (I) CASKWT is able to capture GST-NRX while CASKP673L does not. (J) Effectiveness of capture for CASKWT and CASKP673L is plotted after normalization to wild-type. Data is plotted as mean±SEM;

* indicates P<0.05 by two-way ANOVA. n=3. Scale bar for all = 5µm.

124

125 Figure 3.3. Global reduction in CASK expression produces ONH.

(A) Immunoblot shows reduced levels of CASK in Caskfl/fl ON compared to wild-type controls. (n=3 mice per genotype). (B) Toluidine blue staining shows reduced ON size in

Caskfl/fl mice compared to wild-type controls. Scale bar = 50µm. (C) Quantification of reduced cross-sectional area in Caskfl/fl compared with Caskfl/+ and Cask+/+ (n=4 mice per genotype). (D) Reduction in number of cells in GCL layer using RBPMS (RNA-Binding

Protein with Multiple Splicing). Scale bar = 50 µm. (E) Quantification of reduced number of cells in GCL of Caskfl/fl compared with Caskfl/+ and Cask+/+, and Caskfl/+ compared to

Cask+/+ (n=10 mice per genotype). (F) TEM of RGC axons in the ON of Caskfl/+ and wildtype controls. Scale bar = 5 µm. (G) Quantification of density of axons per image in

Caskfl/fl mice (n=4 mice per genotype). (H-J) Analysis of axon area in Caskfl/fl ON and wild- type controls. Quantification of the average size of fine axons (I) and course axons (J).

For all panels, data is plotted as mean±SEM; *indicates p<0.05, **indicates p<0.01 by two-way ANOVA.

126

127 Figure 3.4. CASK is expressed in Retinal Ganglion Cells.

(A) Schematic of mouse retina. ONL – outer nuclear layer, INL- inner nuclear layer, GCL- ganglion cell layer. (B) In Situ hybridization shows Cask mRNA is expressed in the INL and GCL of the P14 retina. Scale bar = 50µm. B’ shows higher magnification of expression in GCL. Scale bar = 10µm. (C) No appreciable signal was detected in the GCL with the sense riboprobe. C’ shows higher magnification of GCL in C. (D) CASK immunoreactivity in the GCL of P21 retina. Scale bar = 50µm.

128

129

Figure 3.5. Calb2-Cre labels Retinal Ganglion Cells.

(A) Retinal whole mount of the GCL of Calb2-Cre::Rosa-stop-tdT (Calb2-Cre::tdT). Note the large number of cells labeled with tdT. (B) Retinal cross section of Calb2-Cre::tdT shows tdT+ cells in the INL and GCL. (C) tdT+ cells in the GCL of Calb2-Cre::tdT co-label with RBPMS. For all panels, scale bar = 50µm.

130

131

Figure 3.6. RGC-derived CASK is not required for RGC survival.

(A) Toluidine blue staining shows no reduction in ON size in Caskfl/fl::Calb2-Cre+/+ compared to Caskfl/fl. Scale bar = 50µm. (B) Quantification of cross-sectional area in

Caskfl/fl::Calb2-Cre+/+ compared to Caskfl/fl. (n=4 mice per genotype). (C) No reduction in number of cells in GCL layer using RBPMS (RNA-Binding Protein with Multiple Splicing).

Scale bar = 50 µm. (D) Quantification of number of cells in GCL in Caskfl/fl::Calb2-Cre+/+ compared to Caskfl/fl. (n=10 mice per genotype). (E) TEM of RGC axons in the ON in

Caskfl/fl::Calb2-Cre+/+ compared to Caskfl/fl. Scale bar = 5 µm. (F) Quantification of density of axons per image in Caskfl/fl::Calb2-Cre+/+ compared to Caskfl/fl. (n=4 mice per genotype). (G-I). Analysis of axon area in Caskfl/fl::Calb2-Cre+/+ ON compared to Caskfl/fl.

Quantification of the average size of fine axons (I) and course axons (J). For all panels data is plotted as mean±SEM.

132

133

Supplemental Figure 1: CASKWT PSG model adopts, on average, a more compact conformation than CASKP673L.

Box plots of the minimum, median (line), mean (x), and maximum radius of gyration (Rg) of the protein backbone calculated at each timepoint of the concatenated MD trajectories

WT P673L WT for the PSG structures of CASK and CASK . The mean Rg’s (CASK , 2.48 nm;

CASKP673L, 2.51 nm) are statistically significantly different from each other based on a

Welch’s t-test, chosen due to the evident unequal variance, as confirmed using

Levene’s test.

134

135

Supplemental Figure 2: Delay of myelination in Caskfl/fl RGC axons

(A) TEM of RGC axons in the ON in Caskfl/fl and wild-type controls at P12 and P18. Scale bar = 5 µm. (B) Quantification axon of density (per µm2). (C) G-ratio measurements (i.e. myelin thickness divided by axon diameter) in age-matched mutants and controls. n=3 mice per genotype. * denotes p < 0.05 by two-way ANOVA. Data is plotted as mean±SEM.

136

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4. Discussion and Future Perspectives ONH is a devastating disease of development, generally diagnosed in children between birth and three years of age (Mohney et al., 2013). Importantly, onset of this disease occurs at an age when the visual system is still plastic and can be targeted.

Thus, by understanding the etiopathogenesis of ONH and developing models to understand mechanisms underlying its pathology, we can target potential therapeutics to treat ONH. ONH has been described in humans as thinning of the optic nerve and a loss of retinal ganglion cells (Lambert et al., 1987) and, here I describe two different mouse models (i.e. CASK mutant mice) that develop ONH and exhibit RGC loss.

Results I present here confirm my hypothesis that disruption in CASK signaling generates appropriate models for studying the development and pathogenesis of ONH and demonstrate for the first time the developmental progression of ONH in mice.

Moreover, heterozygous loss of CASK, as well as a global reduction of CASK, leads to a reduction in optic nerve size as well as a reduction in the number of RGCs. Surprisingly,

RGC-derived CASK was not necessary for survival of RGCs. While the former is not surprising given the pathogenesis of CASK mutations in humans, what was surprising was the RGC-derived CASK is not necessary for the survival of RGCs or for ON size.

This suggests a non-cell autonomous role for CASK, discussed in more detail below.

4.1. Roles for CASK in development and ONH CASK has well-established roles as a presynaptic scaffolding protein. Based on the different domains of CASK and as a MAGUK protein, CASK has been implicated to

143 have potential roles in synaptic formation, protein trafficking, and transcriptional regulation. Initial reports showed that deletion of CASK does not alter the ability of a neuron to form a structurally normal synapse in vitro with only limited functional defects including reduction in dendritic spine density (Atasoy et al., 2007; Chao et al., 2008), however my in vivo analysis indicates that there is a reduction in total number of synapses due to a heterozygous loss of CASK (Fig. 2.4). This may be due to a pruning of weak synapses, since loss of CASK has been shown to lead to less evoked synaptic transmission and fewer spontaneous synaptic events in Drosophila neuromuscular junctions (NMJ) (Chen and Featherstone, 2011). Interestingly, this data contradicts previous reports that argues deletion of CASK leads to an increase in spontaneous neurotransmitter release at NMJ with no alteration in vesicle size (Zordan et al., 2005).

Homozygous deletion of CASK in vivo in mice is lethal, however, they exhibit no major macroscopic deformities apart from a partially penetrant cleft palate syndrome (Atasoy et al., 2007). Here, I have shown that heterozygous loss of CASK in vivo leads to ONH with reduced number of RGCs, reduced axons in the ON, and reduced axon diameter in the ON.

As CASK is believed to have many roles in vivo, I was interested in the mechanism with which loss of CASK leads to ONH, as well as CASK’s role in development. Are cells lacking CASK still able to make synapses? Are they functional?

Does loss of CASK result in systemic disruption of retinal processes that then causes loss of RGCs, or is it the reverse order? My initial hypothesis, based in vitro and in vivo analysis of CASK function, was a cell-autonomous role for CASK. However, analysis of my data shows that CASK may act through both cell-autonomous and non-

144 cellautonomous pathways in different tissues, different developmental periods, and different diseases.

4.1.1. Cell-Autonomous roles for CASK

Previous literature has shown cell-autonomous roles for CASK in the recruitment of presynaptic machinery (Chen and Featherstone, 2011). I therefore hypothesized that

CASK functioned in a cell-autonomous role to produce ONH. As such, I expected that loss of CASK from targeted cells would lead to specific deficits through the inability to form functional synapses in the cells without CASK, death of those cells, or some other loss of function. I focused my attention in a main retinorecipient region innervated by retinal axons, the dorsal LGN of the thalamus. If loss of CASK causes ONH in a cellautonomous way, I expected to see that loss of CASK in RGCs would alter their ability to form synaptic inputs onto thalamocortical relay cells. Using a fluorescently conjugated anterograde tracer, Cholera Toxin B (CTB), I labeled RGCs, their axons, and retinal nerve terminals in the dLGN in Cask+/- mice and their littermates. With this technique I estimated the total number of retinogeniculate synapses in dLGN as well as the size of these synapses. Loss of CASK led to a total reduction in the number of synapses in adulthood. As there is both a loss of axons and synapses, loss of CASK may either 1.) Fail to successfully target the retinorecipient nuclei, and thus no synaptogenesis occurs or 2.) Successfully target LGN but not create a functional synapse and is then pruned. In regards to the first option, many mechanisms have been identified as important in the process of RGC axonal targeting and the formation of retinotopic maps in the dLGN (Diao et al., 2018). Dozens of molecular guidance cues,

145 temporally controlled in specific spatial patterns throughout the retina, optic tract, and in target nuclei, have been identified as necessary for pathfinding of RGC axons (Diao et al., 2018). Some genes identified have been cell surface molecules like ephrins

(Nakagawa et al., 2000; Triplett and Feldheim, 2010) and extracellular matrix proteins like aggrecan (Bandtlow and Zimmerman, 2000) and reelin (Su et al., 2011). However, in all of the genes listed above, disruption in their functions leads to a mistargeting or disorganization of these RGC axons, rather than the lack of targeting observed in

Cask+/- dLGN. Interestingly, a cell adhesion molecule, Contactin-4, has recently been identified as necessary for targeting in a manner which leads to a reduced innervation in target nuclei, as opposed to mistargeting (Osterhout et al., 2015). In this case, however,

Contactin-4 is a homophilic adhesion protein expressed in both RGCs and their target nuclei, notably different than CASK’s role as a synaptic adhesion protein. Alternatively, it is possible that RGCs lacking CASK are still able to successfully target these nuclei but do not create functional synapses and may then be more susceptible to use-dependent pruning. It has been shown that loss of neurexins leads to non-functional synapses

(Missler et al., 2003). As binding with CASK is necessary for proper function of neurexins (Fairless et al., 2008), this may suggest that the loss of smaller RGC terminals in the dLGN of Cask+/- mice could be due to loss of non-functional synapses.

However, though loss of neurexins in vivo impaired neurotransmitter release, they could not demonstrate noticeable changes structurally to suggest activity-dependent elimination (Fairless et al., 2008), suggesting that potential loss of CASK deficient synapses is through another mechanism. Many neurological diseases such as autism and schizophrenia are diseases of the synaptome (Grant 2012; Vonhoff and Keshishian,

146 2017). Interestingly, in schizophrenia, decreased spine density (Glantz and Lewis, 2000) and a decrease in presynaptic proteins (Faludi and Mirnics, 2011) has suggested that there is a loss of synapses due to errant pruning (Boksa, 2012).

Alternatively, previous work has shown that the inability of RGCs to form synapses on target areas has an effect first on development of dendritic morphology in

RGCs and then eventually cell death (Hughes and LaVelle, 1975). This degeneration and death of RGCs results from the loss of retrograde transport of neurotrophic factors from target neurons in the retinorecipient region. There is also a possibility that, as

CASK is expressed in relay cells of dLGN as early as P3 (unpublished data, Fox Lab), loss of CASK in relay cells may be fatal to these cells and therefore RGCs would have a reduced number of targets in the LGN.

As CASK is widely expressed in the central nervous system and mutations in

CASK often result in cerebellar hypoplasia (Najm et al., 2008), I was also interested in the role of CASK outside of the retinogeniculate pathway, and the potential for a cellautonomous role of CASK in motor function. Disruption of CASK in Drosophila melanogaster has shown a variety of behavioral phenotypes including defective courtship (Lu et al., 2003) and impaired ability to fly (Zordan et al., 2005). Analysis of different aspects of motor behavior when CASK expression was reduced in Drosophila

(with a CASK-ß null mutation) showed changes in motor initiation, maintenance, speed, and acceleration (Slawson et al., 2011). There is also evidence that these motor deficits result from a defect in the pre-motor circuit, and not from motor neurons as expression of CASK in glutamatergic cells with a CASK-ß null background does not rescue locomotion phenotypes in Drosophila (Slawson et al., 2011). Though I did not observe

147 any locomotor phenotypes in Cask+/- mice (and their ability to swim in the twoalternative forced swim task was indistinguishable from controls), I unexpectedly discovered that the cell-specific deletion of CASK from calretinin expressing cells (which represent interneurons in many telencephalic brain regions and a subpopulation of granule cells in the cerebellum) led to ataxia (Schiffmann et al., 1999; Bearzatto et al., 2005). In

Caskfl/y::Calb2-Cre+/+ I have observed an unsteady gait and dyskinesia (n=4) though I have not yet observed the cerebellum for any structural or molecular defects.

Interestingly, a previous report of deletion of CASK from two different classes of neurons in the cerebellum: Purkinje cells (Caskfl/y::PCP2-Cre+/+) and granule cells

(Caskfl/y::Math5-Cre+/+) showed no changes in cerebellar morphology or other structural changes, nor locomotor deficits (Srivastava et al., 2016). These disparate results could reflect the unique expression patterns of Cre recombinase or it could reflect different developmental patterns of Cre expression in these driver lines. Though as a transcription factor, I would expect that Cre expression would be upregulated very early in the cerebellum of Math5-Cre. More likely, as the unsteady gait does not present until around P40 this phenotype is not exclusively due to a cerebellar hypoplasia or some other defect in the cerebellum as this would lead to presentation of dyskinesia earlier, as is seen in the reelin mutants (Falconer et al., 1951; Alter et al., 1968).

The cerebellum receives input from the motor cortex via the pontine nuclei.

Defects in gait could also then be due to alterations in neurons in motor cortex or pons.

For example, spontaneous reeler knockout mice (Relnrl/rl) which were ininitially identified by their severe ataxic gait, have a very pronounced cortical mislayering in addition to the cerebellar hypoplasia (D’Arcangelo and Curran, 1998). As Calb2-Cre is expressed by

148 interneurons, loss of CASK from interneurons may have a cell-autonomous effect on synapses in cerebellum at a later age. This suggests, considering the loss of RGC axons in Cask mutants, that CASK loss may have a targeted effect on projection neurons.

As we have worked to establish CASK’s role in development and synaptic function, it has been argued that the GK-domain of CASK is able to interact with T-box brain region 1 (TBR1) and upon interaction, CASK is translocated in to the nucleus and increases the transcriptional activity of TBR1 (Hsueh et al., 2000; Wang et al., 2004). As a transcription factor TBR1 has many targets, including NMDAR subunit 2b, Cadherin 9, and Reelin, among others (D’Arcangelo and Curran, 1998; Wang et al., 2004; Bedogni et al., 2010). This suggests that if loss of CASK decreases expression of TBR1 it then alters expression of all of those genes listed. As discussed above, reeler mutant mice

(Relnrl/rl) have cerebellar hypoplasia, cortical mislayering (D’Arcangelo and Curran,

1998), and a distinct unsteady gait, indistinguishable from the one seen in

Caskfl/y::Calb2-Cre+/+. Thus, it would be interesting if alteration in CASK-TBR1-RELN pathway could explain the dyskinesia seen in Caskfl/y::Calb2-Cre+/+. However, Reln mutant mice show dyskinesia at a much earlier developmental age than

Caskfl/y::Calb2Cre+/+, potentially suggesting a different mechanism of action. Moreover, there has been no in vivo data to support a CASK-TBR1 interaction and RNAseq of

Cask+/- showed no alterations in Reelin expression (unpublished data, Mukherjee Lab).

To create a more thorough approach to deleting CASK from RGCs as Calb2-Cre only labels ~91% of RGCs, I crossed the CASK floxed allele mouse with a VGluT2-Cre driver line (Caskfl/fl::VGluT2-Cre+/+). This line, driven by vesicular glutamate transporter 2 expression, has reported expression in 100% of RGCs (Land et al., 2004; Engelund et

149 al., 2004), but is also known to have widespread expression in excitatory neurons throughout the brain, including the target cells of RGCs (Borgius et al., 2010). Though full knockout of CASK (Cask-/-) is lethal perinatally, there have been reported cases in which a neuronal knockout, using a Synapsin-Cre line, is viable (Srivastava et al., 2016).

As such, I did not expect to see that knockout of CASK from VGluT2+ cells would be lethal. However, in a breeding scheme where 12.5% of pups should have been

Caskfl/y::VGluT2-Cre+/-, 1 out of 122 were found alive by the age of P5-7. I changed the breeding strategy to increase the odds, where 18.75% of pups should have been

Caskfl/y::VGluT2-Cre+/-, and another 6.2% should be Caskfl/fl::VGluT2-Cre+/-, altogether nearly 25% of a given litter should be a mutant. In this condition, out of 63 mice, I found

0 viable at the age of P5-P7. This was surprising, as other neuronal knockouts did not produce such effects. Ultimately, VGluT2-Cre as a knock-in may be a more thorough and faithful driver line for neurons, as opposed to Synapsin-Cre which is a transgenic driver line (Rempe et al., 2006). Additionally, though VGlut2 expression is restricted to certain brain regions in adult mice (e.g. it is robustly expressed in dorsal thalamus and largely absent from forebrain), developmentally there is a robust expression of VGlut2 in a much broader population of cortical and subcortical neurons (Borgius et al., 2010).

Analysis of whether these mice are dying prenatally or perinatally would be informative to the necessity of CASK in excitatory neurons, or if this death is due to widespread expression of VGlut2-Cre developmentally. If they are indeed dying perinatally, similar to the Cask-/- mice, do they also display no macroscopic defects in the brain?

150 4.1.2. Non-cell Autonomous roles for CASK

Though my initial hypothesis was for a cell autonomous role of CASK in ONH, there is evidence that CASK may also behave in a non-cell autonomous way. Apart from reduced optic disc size, one of the hallmarks for diagnosis of ONH is a tortuosity of retinal vasculature (Lambert et al., 1987). Using different markers for blood vessels, I have noted in mice that have a global reduction of CASK (Caskfl/fl) the presence of vascular tortuosity. How is it possible that reduction of CASK leads to alteration in this network? Is it due to the loss of RGCs, as we know that RGC health has an impact on vasculature? To date I have only looked at ages where there is already a loss of retinal ganglion cells. Thus, it is possible that the death of retinal ganglion cells may set off a cascade starting with a reactive astrogliosis commonly seen in Muller cells and astrocytes in response to a multitude of traumas (Bringmann et al., 2009). In Muller cells this reactive gliosis causes release of VEGF which at some doses provides neurotrophic effects (Zheng et al., 2012) as well as angiogenesis. Interestingly, prolonged and excessive expression of VEGF in the retina has been shown to have detrimental effects on vasculature as well as Muller cell proliferation, creating a negative feedback loop of cell death (Tolentino et al., 2002; Vecino et al., 2016). Exploring the developmental timeline of potential astrogliosis and vascular tortuosity would be insightful to understanding how these responses may play a role in ONH development and RGC death.

Alternatively, RNAseq of Cask+/- brains showed that the group of genes most altered by heterozygous deletion of CASK were extracellular matrix molecules such as laminins, collagens, and nidogens. To date there has been no research on potential

151 shared pathways of CASK and collagens in the central nervous system, however in a wound healing model knockdown of CASK in keratinocytes led to accelerated attachment to the collagenous matrix suggesting that CASK may have some role in modulating epidermal proliferation (Ojeh et al., 2008). Collagens, laminins, and nidogens are primarily important for the stability of the extracellular matrix and in some cases mutations of these genes can cause lethality in prenatal mice where there is a loss of basement membrane stability (Poschl et al., 2004, Yurchenco et al., 2011).

Interestingly, one of the genes that is altered in Cask+/-brain is Collagen IV, a major component of the basement membrane that has previously been implicated in ocular diseases with noted changes in vasculature (Roy et al., 1994; Bai et al., 2009; Alavi et al., 2016; Mao et al., 2015). It has even been suggested that Col4a1 mutations can cause ONH (Gould et al, 2007). Is the vascular tortuosity observed in Caskfl/fl retinas due to this change in Collagen IV? How is CASK involved in this process? Many questions remain to be investigated regarding how CASK may function in a non-cell autonomous manner.

4.2. Optic Nerve Atrophy or Optic Nerve Hypoplasia? Both ONH and Optic Nerve Atrophy (ONA) are diagnosed in humans via fundus photography. Historically, the separation of these two has been difficult for clinicians and scientists as there is degeneration, and often gliosis, present in both conditions (Kjer et al., 1983; Mosier et al., 1978). As there have been no other distinctions in pathogenesis, biomarkers, or etiogenesis of either disease, the main difference has been the timing of degeneration (Hoyt and Good, 1992). Thus, as the only distinction between the two

152 diseases is timing, I used the Cask+/-mutants to evaluate the developmental progression of optic nerve degeneration to assess whether we are modeling a true ONH phenotype, or mislabeling ONH as ONA. Interestingly, analysis of optic nerve area at P1 showed no difference in size between Cask+/- and female wildtype littermates, but by P6 the optic nerve was reduced in Cask+/- mice. At this age in mice, RGCs have already targeted retino-recipient areas and are in the process of maturing these synapses (Campbell et al., 1992; Lo et al., 2002; Hong and Chen 2011), this corresponds with third trimester development in humans (Dobbing and Sands, 1979; Graw et al., 2010). As this degeneration is occurring before the maturation of the retinofugal pathway, it could be said that this is a true ONH phenotype, not ONA. Alternatively, as the optic nerve is normal compared to wild-type at birth, this may suggest that there is a level of degeneration occurring. It has been argued that better diagnostic tools will differentiate

ONH from ONA, however, my data suggests that this may not be as clear cut as initially suggested.

As CASK haploinsufficiency is seen in human patients diagnosed with ONH, and that we have now confirmed the same phenotype in Cask+/- mice, these mice satisfy the criteria for ‘face validity’ for an animal model of ONH. Having a face-validated model for

ONH allows us to ask questions about the developmental progression of ONH. One of the main questions that has been debated in regards to ONH is whether ONH is due to a primary failure of ganglion cell development (Whinery and Blodi, 1963; Walton and

Robb, 1970) or if degeneration of RGCs came secondary to ONH (Mann et al., 1957;

Mosier et al., 1978). This Cask+/- model of ONH supports the secondary degeneration

153 hypothesis of ONH as at P6 when there is a reduced optic nerve area, there is not yet a noted loss of cells in the GCL layer.

4.3. Developing therapeutics for ONH

Gene therapy has been a promising new approach for treating genetic diseases by delivering exogenous DNA to genetically modify patients. Different ways to target the genome include adeno-associated viruses (AAVs), lentiviruses, small-interfering RNAs

(siRNAs), and CRISPR-cas9. Because of the access point of the eye, its immune privileged nature (Niederkorn et al., 1990), and its relative isolation from the rest of the nervous system, diseases of the retina have been among the first diseases targeted for gene therapy. One such success of this system has been seen in patients with earlyonset retinal dystrophy, often called Leber Congenital Amaurosis (LCA). This disease is caused by deficiency in a retinal pigment epithelium gene (RPE65) necessary for function of rod and cone opsins (Bemelmans et al., 2006). Subretinal injection of

Rpe65 packaged in a lentiviral vector was sufficient to restore retinal function to a near- normal levels in mice that lack endogenous Rpe65 expression (Bemelmans et al.,

2006). Other groups have also shown success of rescuing RPE65-LCA in models using both lentiviral and AAV vectors (Acland et al., 2001, 2005; Bennicelli et al., 2008). Phase

I trials of subretinal AAV delivery in human patients with LCA showed remarkable results in improving visual acuity in these patients (Bainbridge et al., 2008; Maguire et al., 2008;

Hauswirth et al., 2008).

154 While the success of these trials is inspiring, there is still work to be done in developing gene therapy as a technique for other diseases, including ONH. Questions still remain about long term potential for these therapies as some patients develop inflammatory responses to vectors and there have been cases of vector-mediated insertional activation of proto-oncogenes (Baum et al., 2004; Ryu et al., 2008). Though these considerations are noted, is delivery of CASK a viable option for rescue of ONH in our mouse model or long-term for treatment of ONH in humans?

While there have been significant improvements in our ability to develop viral constructs to target specific groups of cells and we can now target RGCs using AAVs with success (McKinnon et al., 2005; Hellstrom et al., 2009; Harvey et al., 2002), AAV capsids have a limited capacity of cargo space as the AAV virus is ~4.7 kilobases (kb) in length. Genes that are shorter than 5 kb such as RPE65 (3.2 kb) (Hamel et al., 1993) are more likely to be successfully transferred. There have been difficulties in transfer of larger genes and in cases were larger genes were successfully packaged, there was a decrease in vector yield (Hermonat et al., 1997; Wu et al., 2010; Dong et al., 2010).

CASK is 404 kb (Moog et al., 2013) and, therefore, much larger than target size of 5 kb for AAVs. Thus, our attempts to package CASK have had limited success. Recently, one group has shown that using a triple vector approach and splitting the target transgene into three separate cassettes increases cargo capacity (Maddalena et al.,

2018). However, this approach requires the target cell to be successfully co-infected by all three vectors, and will still only reliably carry a gene of 15 kb. Alternatively, lentivirals have increased capacity compared to AAVs as one vector can carry 8 kb of genetic information (Greenberg et al., 2006). However, the lentiviral vector largely targets retinal

155 pigment epithelium cells and in some cases photoreceptors (Semple-Rowland et al.,

2007; Calame et al., 2011). Altering viral tropism via pseudotyping has shown limited success, and in one case it has been shown to be transduced by RGCs (van Adel et al.,

2003). Though this is an exciting advancement, ultimately, the lentiviral delivery system still faces the same rate-limiting factor as AAVs, a reduced amount of cargo space.

Theoretically, if there was an advancement in these delivery systems, would

ONH be an ideal target for genetic therapy? ONH is a disease of development (Dobbing and Sands, 1979), seen as early as P6 in mice, corresponding to third trimester of human gestation. Is delivery of CASK at birth in humans too late to rescue the phenotype? Potentially not, as at P6 in mice there is not yet a loss of RGCs. Thus, as there is a delay in the loss of RGCs, delivery of CASK at birth in humans may still have success in preventing this secondary loss of RGCs. However, ONH is rarely diagnosed at birth. The average age of diagnosis for ONH in humans is 2 years old, considerably later in the development of this disease and the subcortical visual system. Increasing the chances of success for gene therapy would therefore require a system of screening for visual deficits earlier. Furthermore, the success of gene therapy so far for retinal diseases has been narrowly focused on photoreceptors. RGCs are much larger cells with axons from mouse retina to dLGN about 15 mm long (LaVail et al., 2003) and therefore a much more difficult target for restoring function of the cell by gene therapy.

4.4. Other causes of ONH and models to understand them

There have been both environmental (non-genetic) links to ONH as well as a number of genetic links. Though my work has focused very narrowly on one genetic link,

156 it is important to gain larger perspective about the pathogenesis of ONH in these different models, if there is any variation of CASK in these models, and how my research contributes to understanding of ONH in general.

4.4.1. Non-Genetic causes of ONH

ONH has historically been considered a sporadic disease with causes attributed to maternal recreational drug use, age of mother at gestation, and other such gestational affects (Garcia-Filion and Borchert, 2013). Maternal alcohol drug use has been reported as teratogenic in the development of visual system (Stromland, 1985;

Flanigan et al., 2008), with up to 90% of children affected by fetal alcohol syndrome

(FAS) showing some variation of ophthalmological defects including strabismus and abnormalities of outer segment (Stromland, 1985, 1987). The two most described visual defects of children with FAS are ONH and increased tortuosity of retinal vessels, both described in ~50% of children with FAS that have noted ophthalmological defects

(Stromland 1994). To understand the consequences of ethanol particularly on retinal development, a variety of models have been developed including rats, mice, and zebrafish. Early groups were able to replicate the ONH phenotype in rat models, showing a reduced optic nerve diameter in 73% of rats born from an ethanol exposed dam (Stromland and Pinazo-duran, 1994). However, they also showed that these rats had other ophthalmic defects including delayed eye weight gain in 80% of rats, anophthalmia, and reduced retinal thickness in some percentage. These phenotypes together suggest that ethanol exposure has a larger effect in eye morphogenesis and ocular globe growth. Analysis of gene expression in ethanol-exposed zebrafish embryos

157 showed an alteration in genes necessary for eye development, including homeobox genes six3a and rx3, both transcription factors (Santos-Ledo et al., 2013). Furthermore, the timing of the ethanol exposure plays a large role as mice exposed to ethanol prenatally show a reduction in retinal area within 48 hours of exposure (Kennedy and

Elliott, 1986), but mice exposed to ethanol postnatally do not show a difference in retina area by P20 (Dursun et al., 2011). Ultimately, pre-natal exposure to ethanol is shown to have profound effects on the development of the eye, cell-fate determination, and migration of retinal cells (Cook et al., 1987) and thus ONH in these models is secondary to more devastating developmental defects. An interesting angle that should be pursued further is the possibility that this prenatal exposure to ethanol may alter expression or distribution of CASK and ONH is in this condition may still be as a result of CASK alterations.

4.4.2. Genetic variants in ONH

Large scale registries of children with ONH have noted the low occurrence of siblings (Ryabets-Lienhard et al., 2016). Interestingly, we were contacted by a family in which all 3 daughters were diagnosed with ONH. Historically, the lack of molecular diagnostic strategies and the seemingly sparsity of familial cases suggested that ONH was a largely sporadic disease. As more cases of familial ONH have been reported

(Hackenbruch, 1975; Cidis et al., 1997; Benner et al., 1990), and as molecular diagnostic tools have improved, cohorts of patients have been identified with similar mutations. As such, there have been multiple genes indicated as causative of ONH including PAX6, SOX2, HESX1. The majority of these genes are transcription factors

158 with roles eye and brain morphogenesis. Human patients with mutations in these genes present with a variety of ophthalmic and neurological defects from aniridia to olfactory dysfunctions (Malandrini et al., 2001; Sisodiya et al., 2001). Null mutations in mouse models of these genes often present devastating developmental defects such as anophthalmia, cranio-facial defects, and often perinatal lethality (Sale et al., 2002, Chao et al., 2003; Wang et al., 2017). Interestingly, as there is a comorbidity of ophthalmological and neurological defects in mouse models with null deletions of these genes, an argument could be made that though these models are not accurate representations of ONH in its isolated form, they may be better suited for understanding diseases such as SOD and septo-optic-pituitary dysplasia. Conditional deletions of

PAX6 and SOX2 from the ectoderm primarily affects the development of the lens (Smith et al., 2009) as retinal lamination remains largely intact, though the retina is misplaced

(Ashery-Padan et al., 2000). There has been little reported about the presence of ONH or the morphological integrity of the brain in these conditional models or if these mice are more viable than the null mutations of these genes. As these studies have primarily focused on deleting these genes prior to lens development at E8.5-9, finding a driver line to delete these conditional alleles after the development of the lens would be helpful to understand the role of SOX2 and PAX6 on RGCs and their axons later in development.

4.4.3. How other causes of ONH relate to CASK

Though I have presented them as separate factors, both environmental factors as well as non-CASK genetic links may still alter CASK expression or function either by

159 directly interacting with CASK or by altering pathways associated with CASK. Thus, I will discuss the potential of using non-CASK models (such as PAX6, etc.) to further understand CASK’s role in ONH.

Though models of ethanol exposure have effects on eye morphogenesis that predate ONH, it is important to consider the possibility that different doses of ethanol at different gestational times may have preferential effects on different genes. Thus, it is still possible that at some point ethanol exposure may alter expression of CASK.

Interestingly, though total retinal area is not altered by postnatal exposure to ethanol, there is a reported loss of neurons in GCL (Dursun et al., 2011). This is similar to the phenotype I have reported in Cask variant mice. Proteomics for CASK in any of these models of ethanol exposure would be interesting, particularly if ethanol exposure occurs after optic vesicle evagination has already taken place. There is also evidence that mutations in the hedgehog signaling pathway can exacerbate effects of prenatal ethanol exposure (Kietzman et al., 2014), which begs the question is there an interaction such as this with CASK as well? Are there patients who receive a “double hit” of CASK mutation and prenatal ethanol exposure at higher risk for developing ONH?

Given that ONH can occur in an isolated or syndromic fashion, it is likely that

ONH is a multigenic disease (as the number of patients with CASK mutation is considerably lower than those with ONH). Furthermore, a majority of genes that have been associated with ONH have been transcription factors and thus may function to repress CASK expression either via direct interaction or by downstream targeting. In fact, both PAX6 and SOX2 have been indicated as potential transcription factors for

CASK (Matys et al., 2003, 2006). Though mouse models of PAX6 and SOX2 are difficult

160 to study as they often have anophthalmia and perinatal lethality, it would be interesting to note if there was an alteration in the expression of CASK that could explain why these genes have been linked to ONH. It is also possible that mutation in one of these transcription factors may affect expression of the other transcription factors, creating a cycle where it is difficult to elucidate the culprit behind a detected phenotype.

Interestingly, PAX6 and HESX1 have both been independently associated with ONH but

Hesx1-/-mice show reduced PAX6 expression in the neural plate (Andoniadou et al.,

2007) showing an interaction of these two genes in brain morphogenesis, raising the question whether phenotypes seen in HESX1 animal models may be due to alterations in PAX6 expression.

Normal expression and function of CASK is necessary for the development of the subcortical visual circuitry. I have shown here two different mutations of CASK in mouse models that result in ONH, supporting literature in human patients of CASK’s genetic link to ONH. Interestingly, due to the nature of the other known environmental (ethanol) and genetic (PAX6 and SOX2) risk factors for ONH, it is possible that these links are still the result of aberrations in CASK function. This possibility should be explored further.

Furthermore, though I hypothesized that CASK behaves in a cell autonomous manner to produce ONH via RGC-derived CASK, my evidence suggests that there are non-cell autonomous roles for CASK in ONH, particularly regarding changes in retinal vasculature, a feature commonly described in patients with ONH. How does this change in retinal vasculature alter support for RGCs, which are already preferentially affected by loss of CASK?

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