The Association of COG5 and SRD5A3 Gene Mutations to ER Stress Activation and Their Role in Retinal Degeneration

The Association of COG5 and SRD5A3 Gene Mutations to ER Stress Activation and their Role in Retinal Degeneration

Sami Tabbarah

A thesis submitted in conformity with the requirements for the degree of Master of Science Department of Physiology University of Toronto

The Association of COG5 and SRD5A3 Gene Mutations to ER Stress Activation and their Role in Retinal Degeneration

Sami Tabbarah

Master of Science

Department of Physiology University of Toronto

2018 Abstract

Five individuals were diagnosed with Leber’s congenital amaurosis, an early-onset inherited vision disorder. Two unrelated individuals have a homozygous mutation in SRD5A3 (p.W19X).

The remaining three are siblings that have compound heterozygous mutations in COG5

(p.M32R; p.S777Q_fs*14). Both SRD5A3 and COG5 play critical roles in glycosylation, which if disrupted could result in ER stress. Transfection of wild-type or mutant variants of SRD5A3 or

COG5 in vitro found no link between SRD5A3 and ER stress. However, PERK was significantly upregulated in response to compound transfection of the mutant COG5 variants.

Immunocytochemical staining revealed that the compound COG5 mutants lead to a significant increase in Golgi fragmentation and DNA damage. Treatment of the transfected cells with PERK inhibitor, GSK2606414, reduced DNA damage levels to that of the control. This study is the first to associate COG5 disruption with ER stress and has identified a potential therapeutic target for patients with COG5-related disorders.

Acknowledgments

My sincere thanks go towards my supervisor, Dr. Philippe Monnier, for his guidance and support throughout this project and for motivating me to expand the goals of this study. He has provided me with the opportunity to grow and develop as a scientist in his lab, and for that, I am very grateful. I would also like to greatly thank my co-supervisor, Dr. Elise Héon, for providing the basis through which this project was built on as well as her continued support, guidance, and words of encouragement entirely throughout. Many thanks additionally go to my supervisory committee members, Dr. Valerie Wallace and Dr. James Eubanks, for the time and care you have both put into shaping my project and for offering invaluable insight and direction during our meetings.

The time I spent working on this project was also shaped by the support I have received from my fellow lab members, many of whom have offered invaluable tips and support throughout my many trials and tribulations. I would like to especially acknowledge Dr. Jason Charish for helping me find my way around the lab and taking the time to guide me through my first experiments. I would also like to thank Dr. Hidekiyo Harada for his help on many occasions and his patience as I discuss my troubleshooting strategies with him. Moreover, many thanks to my colleagues at Krembil who have become my friends and first line of support.

Many thanks to my parents, family, and friends for your continued love and understanding.

I would also like to acknowledge the patients that formed the basis of this study, and their families, for offering their time, patience and consent to allow us to perform this investigation.

Finally, I would like to thank the generous funding support from the Vision Science Research Program.

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Table of Contents

Acknowledgments...... iii

Table of Contents ...... iv

List of Tables ...... vii

List of Figures ...... viii

List of Abbreviations ...... x

Introduction ...... 1

Background and Literature Review ...... 3

2.1 Retinal Dystrophy ...... 3

2.1.1 Leber’s Congenital Amaurosis ...... 4

2.2 Clinical Cases...... 5

2.2.1 Case A: Two unrelated individuals with a shared SRD5A3 variant...... 5

2.2.2 Case B: Three siblings with shared COG5 variants ...... 8

2.3 SRD5A3: Steroid 5-alpha-reductase 3 ...... 11

2.3.1 Role of SRD5A3 in N-glycosylation ...... 11

2.3.2 SRD5A3-associated congenital disorders of glycosylation ...... 12

2.4 COG: Conserved oligomeric Golgi complex ...... 14

2.4.1 The Golgi complex: Structure and function...... 14

2.4.1.1 Golgi dispersal and DNA damage ...... 15

2.4.1.2 Golgi trafficking machinery ...... 16

2.4.2 COG complex-associated congenital disorders of glycosylation ...... 20

2.4.3 The COG5 subunit ...... 20

2.5 ER stress and the Unfolded Protein Response ...... 22

2.5.1 Protein folding and aggregation ...... 22

2.5.2 UPR: Restoring proteostasis and signalling apoptosis...... 24

2.5.2.1 PERK activation ...... 26 iv

2.5.2.2 ATF6α activation ...... 28

2.5.2.3 IRE1α activation ...... 29

2.5.2.4 Apoptosis signalling ...... 30

2.5.3 ER stress and the UPR in retinal degeneration ...... 31

2.5.4 Therapeutic options targeting ER stress and the UPR ...... 32

Rationale and Hypothesis ...... 34

3.1 Hypothesis...... 35

3.1.1 Objectives ...... 35

Materials and Methods ...... 36

4.1 Mice ...... 36

4.2 DNA constructs ...... 36

4.2.1 Generation of mutant clones ...... 38

4.2.2 DNA plasmid preparation and purification...... 41

4.2.3 Subretinal injection of DNA followed by in vivo electroporation ...... 41

4.2.4 Harvesting, fixation, and sectioning of retinas ...... 42

4.2.5 Immunohistochemical staining of sectioned retinas ...... 42

4.3 In vitro assessment of gene mutations ...... 43

4.3.1 Culturing cells ...... 43

4.3.2 Transfection of cells with DNA constructs ...... 43

4.3.3 Cell lysis preparation and protein quantification ...... 44

4.3.4 Western blotting ...... 44

4.3.5 Fixation and immunocytochemical staining of transfected cells ...... 45

4.3.6 Imaging, analyses, and quantification ...... 46

Results ...... 50

5.1 SRD5A3: Expression and localization ...... 50

5.1.1 Constructs ...... 50 v

5.1.2 ER stress and UPR expression analysis ...... 52

5.1.3 In Vitro localization ...... 55

5.2 COG5: Expression, localization, & function ...... 57

5.2.1 Constructs ...... 57

5.2.2 ER stress and UPR expression analysis ...... 59

5.2.3 In Vitro localization and Golgi dispersal ...... 63

5.2.4 DNA damage induction ...... 67

Discussion ...... 75

6.1 SRD5A3 ...... 75

6.2 COG5 ...... 77

References ...... 85

A. Appendix ...... 105

A.1 Western blot expression of COG5 plasmids ...... 105

A.2 Co-localization analysis of COG5 with Giantin ...... 106

List of Tables

Table Description Page

Table 2.1 Phenotype summary for patients from Case A 7

Table 2.2 Phenotype summary for patients from Case B 10

Table 4.1 List of all DNA plasmids used along with their shortened name 37

Table 4.2 List of primers used for PCR, site-directed mutagenesis, and sequencing 40

Table 4.3 List of primary antibodies used and their respective applications 48

Table 4.4 List of secondary antibodies used and their respective applications 49

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List of Figures

Figure Description Page

Figure 2.1 Genetic pedigrees of the two families from Case A 6

Figure 2.2 Genetic pedigrees of the two families from Case B 9

Figure 2.3 Schematic demonstrating the role of SRD5A3 protein 13

Figure 2.4 Schematic demonstrating ER-Golgi trafficking 19

Figure 2.5 Schematic demonstrating the three UPR branches 25

Figure 5.1 Constructed plasmids for wildtype and mutant SRD5A3 51

Figure 5.2 Representative western blot images for SRD5A3-linked ER stress 53

Figure 5.3 Quantification for ER stress marker expression in SRD5A3 experiments 54

Figure 5.4 Representative images of WT and mutant SRD5A3 localization in vitro 56

Figure 5.5 Constructed plasmids for wildtype and mutant COG5 58

Figure 5.6 Representative western blot images for COG5-linked ER stress 61

Figure 5.7 Quantification for ER stress marker expression in COG5 experiments 62

Figure 5.8 Representative images of WT and mutant COG5 localization in vitro 64

Figure 5.9 Quantification of colocalization between COG5 constructs and GM130 65

Figure 5.10 Quantification of relative Golgi area dispersal in COG5-transfected cells 66

Figure 5.11 Representative images of DNA damage in COG5-transfected cells 69

Figure 5.12 Quantification of DNA damage levels in COG5-transfected cells 70

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Figure 5.13 Representative images of DNA damage in PERKi-treated cells 71

Figure 5.14 Quantification of DNA damage levels in PERKi-treated cells 72

Figure 5.15 Representative images of DNA damage in PERK-transfected cells 73

Figure 5.16 Quantification of DNA damage due to PERK overexpression 74

Figure 6.1 Schematic highlighting novel findings of research study 84

Figure A.1 Western blot confirming the expression of 6x-His tagged COG5 plasmids 105

Figure A.2 Representative images of GFP-tagged COG5 co-localization with giantin 106

Figure A.3 Quantification of colocalization between COG5 constructs and giantin 107

List of Abbreviations

4-PBA Sodium 4-phenylbutyrate

AAV Adeno-associated vector

ATF/CREB Activating transcription factor/cAMP response element binding protein

ATF4 Activating transcription factor 4

ATF6 Activating transcription factor 6

BCA Bicinchoninic acid

BiP Immunoglobulin binding protein

BSA Bovine serum albumin bZIP Basic leucine zipper

Caspase Cysteine-dependent aspartate-specific proteases

CDGs Congenital disorders of glycosylation

CHOP CCAATT-enhancer-binding proteins (C/EBPS) homologous protein

COG5 Conserved oligomeric Golgi complex, subunit 5

COP Coat protein

DAPI 4’,6-Diamidino-2-phenylindole ddH2O Double-distilled water

DHT 5-alpha-dihydrotestosterone

DMEM Dulbecco’s Modified Eagle’s Medium

DNA Deoxyribonucleic acid

DNA-PK DNA-dependent protein kinase

D-PBS Dulbecco’s phosphate buffered saline

DTT Dithiothreitol eIF2α Eukaryotic initiation factor-2α

EmGFP Emerald green fluorescent protein

ERAD ER-associated protein degradation

ERG Electroretinogram

ERGIC ER-Golgi intermediate compartment

Ero1 ER oxidoreductin 1

FBS Fetal bovine serum

GAPDH Glyceraldehyde 3-phosphate dehydrogenase

GFP Green fluorescent protein

GOLPH3 Golgi phosphoprotein 3

GRASP Golgi reassembly stacking protein

HSP Heat shock protein

IPTG Isopropyl β-D-1-thiogalactopyranoside

IRE1 Inositol-requiring kinase 1

KO Knockout

LAMP3 Lysosomal-associated membrane protein 3

LCA Leber’s congenital amaurosis

MEM Minimal essential medium

MRI Magnetic resonance imaging

MYO18A Myosin 18A

Nrf2 Nuclear factor (erythroid-derived 2)-like 2

NSF N-ethylmaleimide-sensitive factor

PBS Phosphate buffered saline

PBS-T Phosphate buffered saline with 0.1% Tween 20

PCR Polymerase chain reaction

PDI Protein disulfide isomerase

PEI Polyethylenimine

PERK Protein kinase RNA-like endoplasmic reticulum kinase

PFA Paraformaldehyde

PI Protease inhibitors

PLL Poly-L-lysine

PPI Peptidyl-prolyl isomerase

RFP Red fluorescent protein

RGC Retinal ganglion cell

RIPA Radio-immunoprecipitation assay

ROI Region of interest

ROS Reactive oxygen species

RP Retinitis pigmentosa

RPE Retinal pigment epithelium

SDS Sodium dodecyl sulfate

PAGE Polyacrylamide gel electrophoresis

S.E.M. Standard error of the mean

SNARE Soluble NSF attachment protein receptor

SRD5A3 Steroid 5-alpha reductase, subunit 3

TGN Trans-Golgi Network

TM Tunicamycin

TUDCA Tauroursodeoxycholic acid

UPR Unfolded protein response

WT Wild-type

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Introduction

Vision disorders and retinal degeneration affect millions around the world, and for many of these diseases, treatment is yet to be found. Recently, the FDA approved the first gene therapy to treat blindness caused by a form of retinal degeneration called Leber’s congenital amaurosis (LCA) that was due to mutations in RPE651. While this marks a significant milestone, LCA has been associated with 25 mutated genes, and the gene therapy approved targets only one of them2. Furthermore, the genetic cause of approximately 30% of LCA cases remains unknown3. It is crucial that genetic causes in LCA patients continue to be identified and investigated. Only then will it be possible to expand the therapeutic options for more types of LCA as well as other inherited disorders.

In this study, the genetic mutations found in five individuals with complex LCA, from three separate families, were investigated. In two unrelated individuals, the same homozygous mutation was identified in SRD5A3, and in three siblings, compound heterozygote mutations were identified in COG5. SRD5A3 plays a key role in N-linked glycosylation in the endoplasmic reticulum (ER), while COG5 is part of an eight-subunit complex that facilitates retrograde Golgi trafficking of vesicles, including those that contain proteins necessary for protein glycosylation and ER processes4,5. As a result, the phenomenon of ER stress and the subsequent activation of the unfolded protein response (UPR) was investigated in relation to the mutations found in SRD5A3 and COG5. These cases were combined as disturbance of those pathways were not documented in LCA, and the mutated genes usually lead to congenital disorders of glycosylation (CDG) without LCA.

Previous studies have documented the identified SRD5A3 mutation with a congenital disorder of glycosylation (type 1a) that involves retinal degeneration6. The mutation produces a non- functional protein, but the cellular effect of the truncated protein has not been examined. This study attempted to identify if the mutant SRD5A3 variant causes ER stress, but no link was found. In contrast, one of the identified COG5 mutations are novel, and COG5 mutation have been linked to CDGs without retinal degeneration. Additionally, investigations into the role of the COG complex has been confined to its role in Golgi trafficking and Golgi-mediated glycosylation, but its effect on the ER has not been studied. Through ER stress analysis, the COG5 mutations were found to have a direct effect on inducing ER stress. As a result, this study

1 2 will only briefly discuss the findings in relation to SRD5A3 and will place a greater emphasis on COG5. The investigation into the COG5 mutations further identified a novel link between upregulation of PERK – an arm of the UPR – and an increase in DNA damage. The findings presented here not only expand the roles that the COG complex and PERK play in pathogenicity, but expand the CDG phenotype spectrum, add a new category of LCA genes and provides an opportunity to identify potential therapeutic target for patients with COG5 mutations.

Background and Literature Review 2.1 Retinal Dystrophy

Retinal dystrophy is an umbrella term encompassing several diseases that cause retinal degeneration of the outer retina and that are usually genetically-determined. The retina is a multi- layered, light-sensitive structure at the back of the eye that plays a crucial role in vision transduction. Retinal dystrophies can occur as non-syndromic, meaning they are not accompanied by any other signs, or syndromic, as part of a larger group of signs affecting the patient to varying degrees. The prevalence of general retinal dystrophies was found to range between 1 in 750 individuals7 to 1 in 9000 individuals8 based on geographical location. The genetic heterogeneity of retinal dystrophies is great and encompasses many different patterns of inheritance, including autosomal dominant, autosomal recessive, or X-linked inheritance. The focus of this thesis relates to autosomal recessive inheritance.

A commonly diagnosed form of retinal dystrophy is retinitis pigmentosa (RP), an inherited vision disorder that is genetically and phenotypically heterogeneous but is primarily caused by the degeneration of rod photoreceptors. Common symptoms of RP include night blindness, loss of peripheral vision, and deteriorated visual acuity9. Various gene mutations have been associated with RP, and 30% of dominant RP cases result from a mutation in rhodopsin, a key G- protein-coupled receptor membrane protein that comprises the bulk of all proteins in the rod photoreceptor outer segments and is exclusively found in rod photoreceptor cells10,11. Rhodopsin absorbs light and initiates the visual transduction pathway, and mutations in the RHO gene can result in misfolded rhodopsin protein, which causes rod photoreceptor degeneration11. Ultimately, RP devolves from the selective variable degeneration of rod photoreceptors to the degeneration of the retinal pigment epithelium (RPE) and the cone photoreceptors, resulting in complete blindness in some cases.

Retinal dystrophies are genetically and clinically heterogeneous. Depending on the predominant photoreceptor involved, it may lead to forms of retinal disease such as rod-cone dystrophy or cone-rod dystrophy. Cone-rod dystrophy is marked with symptoms such as loss of central vision and colour vision defects, before its development to encompass loss of night and peripheral vision9.

2.1.1 Leber’s Congenital Amaurosis

Leber’s congenital amaurosis (LCA) is a severe and early-onset form of retinal degeneration. It manifests as a rapid and severe degradation of the rod and cone photoreceptors and may culminate to complete loss of vision2. Diagnosis of LCA can be made if three conditions are met: early and severe vision loss, nystagmus, and severely attenuated or non-detectable electroretinogram (ERG) response12. Its prevalence varies between 1 in 30,00013 to 1 in 81,00014 individuals.

LCA is genetically heterogeneous with over 20 disease-causing genes associated with it15. Its genetic heterogeneity results in heterogeneous expressions of the disease, which assist in identifying the underlying causative gene mutation16. A frequent clinical co-occurrence of LCA is nystagmus, which is an involuntary shaky movement of the eyes that disrupts vision. LCA is phenotypically heterogeneous and can also be syndromic and associated with non-ocular characteristics such as mental retardation17, olfactory dysfunction18, and renal failure19.

There were no treatments for LCA until very recently when gene therapy was approved to specifically treat LCA caused by a homozygous RPE65 mutation, which accounts for 8% of LCA cases. RPE65 is an RPE-resident protein that is required by rod-photoreceptors to supply 11-cis retinal, which is necessary for rhodopsin regeneration and the photoreceptors’ ability to respond to light20. LCA caused by mutations in RPE65 is more moderate than other forms, as it presents some visual impairment at infancy but does not progress to complete blindness until the patient is past middle-age. One of the main reasons that RPE65-associated LCA was a prime candidate for gene therapy was that its retinal cells remain relatively preserved16. Gene therapy is currently under investigation for several other inherited vision disorders such as achromatopsia21, retinitis pigmentosa22, and choroideremia23. Gene therapy remains in its infancy, but many clinical trials show great promise for a burgeoning market of approved gene therapy products that could one day treat many more forms of LCA. Studies such as this one are needed to illuminate the myriad causes of LCA and develop targeted treatment options as well as more general ones that encompass disruptions of similar nature.

2.2 Clinical Cases

Several patients with severe vision loss and retinopathy were diagnosed by Dr. Elise Héon to have LCA. The patients with LCA, and no mutation identified using clinical genetic testing, underwent whole exome sequencing (Illumina HiSeq). The sequencing results of roughly 5,000,000 variants were initially passed through a quality filter to ensure true heterozygous or homozygous mutations and then compared against various population databases to cross- reference with frequently-reported variants. The coding single nucleotide polymorphisms (SNPs) were kept and initially validated in silico through internal and publicly-available filtrate protocols. In the end, the shared variants between patients were identified. In Case A, for two unrelated individuals, the shared mutation in the SRD5A3 gene is a prime candidate for causing LCA. In Case B, two COG5 gene mutations were found to be shared by three siblings. In both cases, the variants identified were deemed pathogenic after undergoing a strict validation protocols. Details on patient demographics and symptoms for both cases as well as the identified gene mutations are discussed below (Tables 2.1, 2.2).

2.2.1 Case A: Two unrelated individuals with a shared SRD5A3 variant

Two unrelated individuals of different ethnic backgrounds presented with LCA symptoms and were found to have a homozygous SRD5A3 nonsense mutation (p.W19X) that was previously described (Figure 2.1)6. For both probands, the parents were heterozygous for the mutation and did not demonstrate any related phenotype. In Family 1, the proband, EPS_1.3 of Indian origin demonstrated at age 4 cognitive and motor delay, unsteady gait, microcephaly, and small optic chiasm. EPS_1.3 was diagnosed with LCA and demonstrated nystagmus, diffuse retinal atrophy, dull foveal reflex, and severely attenuated ERG results. In Family 2, the proband, EPS_2.3, is of Iranian origin and at the age of 18, presented cognitive and motor delay, some developmental delay, hearing loss in the left ear, hypotonia, short stature, and microcephaly. Along with the diagnosis of LCA, patient EPS_2.3 had nystagmus, bony spicule pigmentation of the retina, dull foveal reflex, cystic macular edema, and no recordable ERG. Neither child met the clinical diagnostic criteria of congenital disorder of glycosylation (CGD), previously associated with these mutations (OMIM 612379). The patients described here had severe vision loss from birth, while previously identified patients with the same mutations appear to have more early-onset retinal dystrophy. A summary of phenotypes for both probands is presented in Table 2.1.

Figure 2.1: Genetic pedigrees of the two families from Case A that both include a proband diagnosed with LCA and expressing various other severe symptoms. Family 1 has one daughter (EPS_1.3) who is homozygous for the autosomal recessive SRD5A3 mutation, p.W19X. Family 2 has a son and a daughter, but only the son (EPS_2.3) is homozygous for the p.W19X SRD5A3 mutation, while the daughter (EPS_2.4) did not inherit the mutant allele from either parent. The parents in both families are heterozygous for the SRD5A3 mutation. A filled-in shading of shape indicate patients with LCA diagnosis.

Table 2.1: Phenotype summary for patients EPS_1.3 and EPS_2.3 from Case A families with identified homozygous SRD5A3 (p.W19X) mutation. Proband EPS_1.3 EPS_2.3 Gender Female Male Ethnicity Indian (South Asian) Iranian Systemic Phenotypes Cognition Delayed Delayed Motor function Delayed Delayed Hearing ability Normal Sensorineural hearing loss in left ear Gait Unsteady Hypotonia, poor balance Stature Normal Thoracic kyphosis, short stature Dysmorphism Deep-set eyes, wide nasal None bridge, posteriorly-rotated pinnae Head diameter Microcephaly Microcephaly Head and Neck MRI Small optic chiasm Normal Ocular Phenotypes Clinical diagnosis LCA LCA Visual acuity Unable to see Teller chart 20/200 OD and 20/60 OS Nystagmus Yes Yes Anterior segment Normal Normal Retinal condition Diffuse retinal atrophy, mild Bone spicule pigmentation, dull optic nerve atrophy, mild RPE foveal reflex, attenuated vessels, changes in the periphery, dull cystic macular edema, rounded foveal reflex large variable-shaped atrophy outside the vascular arcades Visual field Not tested 25° horizontal and 20° vertical (diameter) (for III-4e) (at 23yrs) ERG Rod-isolated b-waves severely Extinguished reduced in amplitude. Mixed rod-cone responses are reduced, consistent with a rod-cone dysfunction (at 4yrs)

2.2.2 Case B: Three siblings with shared COG5 variants

Three siblings of Filipino origin presented with a complex phenotype that included LCA. Exome sequencing identified a compound heterozygous mutation in the COG5 alleles for all three probands, HMT_1.3, HMT_1.4, and HMT_1.5. A missense COG5 mutation (p.M32R) was inherited from the probands’ father, while a stop-gain frameshift COG5 mutation (p.S777Q_fs*14) was inherited from the probands’ mother. The stop codon that results from this frame-shift mutation appears 14 codons downstream of the mutation. Both parents were heterozygous for their particular COG5 mutations and did not display any remarkable phenotypes. The siblings HMT_1.3, HMT_1.4, and HMT_1.5 were assessed at ages 17, 18, and 20, respectively. All three probands had normal cognitive and motor abilities but had short stature and microcephaly. One of the siblings had bilateral sensorineural and conductive hearing loss. Along with their LCA diagnosis, the probands’ also presented with ocular phenotypes such as nystagmus, cone-rod dystrophy, low visual acuity, and limited visual field. None of the siblings met the clinical diagnosis criteria of congenital disorder of glycosylation (CDG), which is usually associated with COG5 mutations (OMIM 613612). A summary of assessed phenotypes for the three probands of Family 3 is listed in Table 2.2.

Figure 2.2: Genetic pedigree of the family from Case B with three probands diagnosed with LCA. Each of the three siblings (HMT_1.3, HMT_1.4, and HMT_1.5) are compound heterozygous for mutations in COG5. They all inherited one mutant allele with a missense COG5 mutation (p.M32R) from their mother, and a frameshift stop-gain mutation in COG5 (p.S777Q_fs*14) from their father. The parents are heterozygous for their respective COG5 mutations. A filled-in shading of a shape indicate the patients diagnosed with LCA.

Table 2.2: Phenotype summary for patients HMT_1.3, HMT_1.4, and HMT_1.5 from Case B (Family 3) with identified compound heterozygous COG5 mutations (p.M32R, and p.S777Q_fs*14). Proband HMT_1.3 HMT_1.4 HMT_1.5 Gender Male Female Female Ethnicity Filipino (East Asian) Filipino (East Asian) Filipino (East Asian) Systemic Phenotypes Cognition Normal Normal Normal Motor function Normal - Normal Hearing ability Normal Bilateral sensorineural Normal and conductive hearing loss Gait Normal Unsteady Normal Stature Short stature and Short stature Short stature, diffuse skeletal dysplasia platyspondyly, and hip dysplasia Dysmorphism None None None Head diameter Microcephaly (< 3 Microcephaly (< 2 Microcephaly (< 4 percentile diameter at percentile diameter at percentile diameter at age 9) age 18) age 19) Head and Neck Abnormal signal of Skull thickening, T2 Skull base sclerosis MRI the dentate nuclei hypointensity in bilaterally, short cerebellar dentate corpus callosum nuclei Ocular Phenotypes Clinical diagnosis LCA LCA LCA Visual acuity Low, “counting Low, “counting Low, “counting fingers” fingers” fingers” Nystagmus Yes Yes Yes Anterior segment Normal Normal Normal Retinal condition Diffuse mottling, but Diffuse mottling, and Diffuse mottling no maculopathy central pigmentation Visual field Peripheral island (for 40° central (for III-4e) 20-30° horizontal, 10° (diameter) IV-4e) ( at 18yrs) (at 19yrs) vertical (for IV-4e) (at 21yrs) ERG Severe cone-rod Cone-rod dystrophy Cone-rod dystrophy dystrophy

2.3 SRD5A3: Steroid 5-alpha-reductase 3

The steroid 5-alpha-reductase family is composed of five genes, SRD5A1, SRD5A2, SRD5A2L2, GPSN2, and SRD5A34. However, not all of the encoded proteins have been found to play a role in steroidogenesis. The most well-characterized of the encoded proteins are SRD5A1 and SRD5A2 as both are essential to the conversion of testosterone into 5-alpha-dihydrotestosterone (DHT) as well as reducing other steroids24. SRD5A3 has also been implicated in DHT production25, but conflicting accounts shed doubt on what role it plays in testosterone reduction if any at all26. SRD5A3 has in fact differentiated itself from its other family members for playing an early critical role in N-glycosylation.

2.3.1 Role of SRD5A3 in N-glycosylation

SRD5A3 is an endoplasmic reticulum (ER)-resident protein27 that reduces a polyprenol into a dolichol4. N-glycosylation is a posttranslational modification whereby a glycan is added to asparagine residues of nascent proteins. This process starts in the ER where a block of 14 monosaccharides is assembled on dolichol-phosphate (Dol-P), a lipid carrier, to form the N- glycan precursor28. This glycan is then bound onto the nascent protein which undergoes modification in the ER before being transported to the Golgi apparatus to complete its modification. Complete glycosylation assembly and modification is essential for protein folding, stability, and function29. The lipid carrier, Dol-P, is necessary for N-glycan production and SRD5A3 reduces the terminal bond of an alpha-unsaturated polyprenol to form dolichol, which is then phosphorylated to form Dol-P4 (See Figure 2.3). A disruption in this process will result in N-glycosylation defects, disrupting protein folding and stability.

However, recent evidence suggests that glycosylation defects are not due to dolichol deficit, but rather to an increase in the levels of polyprenol that is not reduced30. This was suggested through an investigation of the level of dolichol in fibroblasts obtained from a patient with a debilitating SRD5A3 mutation (p.W19X). The patient’s symptoms included nystagmus, psychomotor retardation, and muscle hypotonia. The investigation found that dolichol concentration levels matched those of healthy control fibroblasts, even though hypoglycosylation was still occurring30. This could potentially be the result of the unreduced polyprenol competing with dolichol in anchoring to N-glycan biosynthesis reactions. Another hypothesis suggests that while an alternative pathway is producing dolichol, it is not producing it in the right location for it to

12 complete all its tasks within the cell. Finally, the level of dolichol that is required for proper glycosylation may vary between various tissues and cells, and this may explain the selective phenotypes that result from SRD5A3 mutations if dolichol concentrations are not sufficient to meet the demands of those specific relevant cells30.

2.3.2 SRD5A3-associated congenital disorders of glycosylation

Glycosylation defects have been revealed to cause hereditary disorders at birth with wide- ranging symptoms that are usually debilitating. These groups of diseases are referred to as congenital disorders of glycosylation (CDGs) and are split into two types, each with separate groupings based on the mutation causing the glycosylation defects. Type I CDGs result from a disruption of the development and transfer of the glycan precursor onto the nascent protein, whereas type II CDGs result from defects in the subsequent processing of the N-glycan after attachment on the nascent protein. Due to SRD5A3’s role in producing the glycan precursor, mutations in SRD5A3 result in type I CDGs.

While the patients in Case A (section 2.2.1) did not meet the criteria of CDG, several mutations in SRD5A3 leading to CDGs have been reported in the literature with the most common symptoms including optic nerve atrophy, nystagmus, hypotonia, cognitive delays, and ataxia31. Recently, 7 patients from 4 unrelated families with early-onset retinal dystrophy have been reported to have the same homozygous SRD5A3 mutation that we have identified in our two patients (p.W19X) in Case A6. All the patients were of South Asian ethnicity, and their commonly shared phenotypes include cognitive delay, nystagmus, and slow-progressing retinopathy6. And while SRD5A3 is known to play a significant role in glycosylation, it likely has a partially redundant function seeing as it does not lead to lethality. There is a large gap in the literature on the direct effect of SRD5A3 protein disruption on cellular function and the compensatory mechanisms of the cell in producing dolichol.

Figure 2.3: Schematic demonstrating the role of SRD5A3 protein. In the ER membrane, the terminal (α) double-bond of a polyprenol is reduced by SRD5A3 into a dolichol. Dolichol is later phosphorylated (Dol-P) and becomes the site where the N-glycan precursor is assembled. The N- glycan is then transferred onto the nascent protein as it enters the ER from the cytosol. (Adapted from Cantagrel, et al. 2010. Cell)

2.4 COG: Conserved oligomeric Golgi complex

The conserved oligomeric Golgi (COG) complex is a Golgi-resident protein that facilitates retrograde trafficking between Golgi cisternae as well as from the Golgi to the ER. It is an eight- subunit structure with subunits, COG1 – 832. The complex is divided into two lobes, lobe A is comprised of COG1 – 4, and lobe B is of COG5 – 8. The two lobes are linked together through a bond between COG1 and COG833 (See Figure 2.4 A). While the subunits are dependent on each other for stability, each of the subunits binds to unique cargo-containing vesicles to be trafficked throughout the Golgi cisternae34. Many of these vesicles contain glycosylation enzymes and the COG subunits ensure that the proper glycosylation enzymes are recycled and properly sorted into the appropriate cisternae. These glycosylation enzymes play a key role in modifying nascent proteins during posttranslational modification, as they exit the ER and pass through the Golgi. After complete modification in the Golgi, the completely modified and folded protein is secreted for its localization, while defected proteins are sent to lysosomes for degradation5. Mutations in almost all the COG subunits have been linked to CDGs, causing wide-ranging disabilities35. Unfortunately, there are no treatments for any of the COG-CDGs, yet.

2.4.1 The Golgi complex: Structure and function

The Golgi complex is a protein and lipid modification and trafficking center, comprised of organized and stacked compartments, referred to as cisterna. Each cisterna houses a unique mixture of enzymes and proteins that are required for the sequential processing of proteins passing through the Golgi system. Nascent proteins are first folded in the ER where they start post-translational glycosylation, before being shuttled to the Golgi to complete their modification28.

The Golgi is classified into three segments, the cis-, medial-, and trans- Golgi layers, along with a pre-cis-layer and a trans-Golgi network (TGN) that flank the Golgi cisternae (Figure 2.4B). The pre-cis-layer is the closest to the ER and is derived from the ER-Golgi intermediate compartments (ERGICs) which are vesiculo-tubular clusters that facilitate ER-to-Golgi transport36. Coat protein (COP) complexes, or coatomers, are heavily involved in vesicle trafficking and fusion. COPII-coated vesicles transport cargo from the ER exit sites and then likely fuse together to form the ERGIC clusters. These ERGIC clusters form the pre-cis-layer and are thought to act as the first protein sorting stations for anterograde or retrograde

15 trafficking36. COPI-coated vesicles facilitate protein sorting at the pre-cis-layer, as well as retrograde trafficking from the Golgi and the ERGICs to the ER36,37. Retrogradely-trafficked components from the ERGIC include essential ER-resident proteins38. Proteins sorted for anterograde trafficking proceed to the cis-Golgi layer where the early glycosylation processes occur. As the protein progresses into the medial- and trans-Golgi cisternae, it encounters carbohydrate synthesis, with early-acting enzymes found in the medial layer, and late-acting enzymes localized in the trans layer38. In the TGN, proteins are sorted and packaged into ox- coated vesicles for secretion to subcellular destinations or to lysosomes for degradation (Figure 2.4B)39,40.

As the cell passes through the cell cycle and undergoes mitosis, the Golgi’s organized structure is disassembled to facilitate its equal separation into the resultant daughter cells. In the newly- formed cells, the Golgi is quickly reassembled to process proteins properly and quality-check their folding and modifications41. The phosphorylation of Golgi stacking proteins by kinases leads to the fragmentation of Golgi reassembly stacking proteins (GRASPs) which results in the fragmentation and dispersal of the Golgi structure. Phosphatases, on the other hand, facilitate the restacking of the Golgi, by dephosphorylating GRASPs and other stacking proteins and membrane tethers. One such membrane tether is p115, which interacts with GM130, a cis-Golgi matrix protein, to facilitate vesicle fusion with target membranes42.

2.4.1.1 Golgi dispersal and DNA damage

Golgi dispersal and fragmentation are not only characteristics of mitosis but are also observed in several diseases that range in severity and symptoms. These include neurodegenerative diseases such as Parkinson’s disease43, Alzheimer’s disease44, and Huntington’s disease45. For example, in Alzheimer’s disease, amyloid plaques aggregate in patients’ brains, exacerbating symptoms. The amyloid beta peptides that form the amyloid plaques have been found to cause Golgi dispersal. This occurs as a result of the amyloid beta peptides triggering the activation of cyclin- dependent kinase 5, which phosphorylates GRASP6544. This fragmentation disrupts the trafficking and modification of essential proteins for neuronal function and results in major alterations to the composition of lipids and proteins at the cell surface46. Thus, Golgi defects compromise neurons’ function and survival and likely contribute to the pathogenicity of neurodegenerative diseases46.

Recently, it has been found that DNA damage also leads to the fragmentation and dispersal of the Golgi, through a mechanism that involves Golgi phosphoprotein 3 (GOLPH3), myosin 18A (MYO18A), and F-actin47. This was found to occur independently of apoptosis47. GOLPH3 is a trans-Golgi peripheral membrane protein that assists in Golgi trafficking and maintaining its structure by linking trans-Golgi membranes with F-actin and MYO18A. The GOLPH3/MYO18A/F-actin interaction apply a tensile force on the Golgi membrane, resulting in its flattened shape of stacked cisternae48. However, GOLPH3’s overexpression in cells has been found to induce Golgi dispersal49. As previously mentioned, phosphorylation of Golgi integral proteins disrupts Golgi morphology, and as a phosphoprotein, GOLPH3’s contribution to Golgi fragmentation likely results from its phosphorylation. It was found that inhibiting or silencing DNA-dependent protein kinase (DNA-PK) prevented Golgi fragmentation under DNA damage- induction. It was confirmed that DNA-PK directly phosphorylates GOLPH3, which increases its affinity to interact with MYO18A. This enhanced interaction between GOLPH3 and MYO18A severely disrupts Golgi’s structural integrity and contributes to its dispersal under conditions of DNA damage47. Interestingly, GOLPH3 is an oncogene, and its overexpression has been found to increase tumor growth and lead to cancer50. Future studies must investigate the opportunity of targeting the Golgi and GOLPH3 function in developing therapeutic options for cancer, neurodegenerative diseases, and rare genetic disorders.

2.4.1.2 Golgi trafficking machinery

Trafficking between the Golgi and the ER is a bidirectional process, that occurs through a mirrored mechanism both ways. The donor organelle releases a cargo-carrying vesicle through budding, and the vesicle is then tethered to the acceptor organelle that strips it of its coatomer and fuses its contents into the acceptor organelle’s lumen51. COPII coats the vesicles moving from the ER to the Golgi through anterograde trafficking, while COPI coats vesicles for Golgi- to-ER retrograde trafficking (Figure 2.4B)51. The trafficked cargo is very diverse in nature and in destination, and so the trafficking machinery includes several adaptations and molecular chaperones to assist the Golgi in its role as a central trafficking hub52. One group of proteins that facilitate bidirectional Golgi trafficking are golgins, which are long coiled-coils, tethered to the Golgi’s membrane on the cytoplasmic side53. Golgins bind to incoming vesicles and have an affinity for Rab GTPases, which are important for vesicle formation and fusion54. Another vital

17 component for Golgi trafficking is the COG complex, which specifically facilitates retrograde trafficking55.

Retrograde Golgi trafficking is essential as it ensures that proteins used by the ER for export are returned to the ER to continue their function. Additionally, retrograde trafficking is employed by the Golgi to recycle proteins and enzymes that progress through the Golgi cisterna but need to be localized at specific cisternae for a nascent protein’s sequential processing and modification55. The COG complex has been nicknamed “the Golgi puppet master” due to its central role in Golgi trafficking and its wide-ranging interactions with various vesicle trafficking and fusion molecules such as: soluble N-ethylmaleimide-sensitive factor (NSF) attachment protein receptors (SNAREs), SNARE-interacting proteins, Rab proteins, coiled-coil tethers, coatomers, and molecular motors5. Long coiled-coil tethers largely play a role in binding to vesicles and facilitating their transport to their destination, as well as contributing to the architecture of the Golgi complex as peripheral membrane proteins. The COG complex interacts with a variety of these tethers, with COG5 having direct interaction with GM1305,56. Additionally, most of the COG subunits, including COG5, have been found to interact with COPI-coated vesicles, which are involved in retrograde intra-Golgi and Golgi-to-ER trafficking5.

There are several types of SNAREs and SNARE complexes that are involved in vesicle- membrane fusion. Donor membranes or exported vesicles contain a vesicle(v)-SNARE which binds with a target(t)-SNARE complex on an acceptor membrane, initiating fusion57. The COG complex has been found to interact with several SNAREs, but the subunits with the most SNARE-interactions are COG4, COG6, COG7, and COG858. COG5 has no known SNARE interactions, however, it likely has an indirect interaction through its strong linkage with and structural dependence on COG759. COG-SNARE interactions are necessary for Golgi trafficking, and any defects in the COG complex will severely affect this process60.

Rab proteins are a group of over 60 molecular switches that play an essential role in vesicle transport and Golgi trafficking. In their inactive GDP-bound state, Rab proteins are localized to the cytosol, however upon activation through the exchange of GDP to GTP they are localized to specific compartments where they interact with proteins that activate their downstream GTPase- dependent function61. Sometimes these interactions lead to the activation of other Rab proteins leading to a Rab cascade. The COG complex has been shown to interact with 12 Rab proteins,

18 with most interactions occurring with the COG4, COG5, and COG6 subunits. COG5 was found to specifically interact with Rab2a and Rab3956. Rab2a is a cytoplasmic protein that is required for Golgi-to-ER retrograde transport and has been found to coat approximately 60% of all cargo- containing vesicles budding from the Golgi62. Rab39 is also a Golgi-associated cytoplasmic protein that is involved in macroautophagy and facilitates vesicular transport for endocytosis63,64. The COG subunits all had a greater affinity for GTP-bound “active” Rab proteins than GDP- bound Rab proteins, indicating that the COG complex likely plays a role in actively initiating trafficking steps for the Golgi5,56.

While the COG complex’s key role in glycosylation and retrograde Golgi trafficking has been identified, the exact mechanism by which it carries out its function is yet to be confirmed. Several proposed models have been put forward that center on the different findings of the COG complex’s role, such as its spatial landmark formation for vesicular tethers, its stabilization of SNARE complexes, and its direct interaction with vesicle coatomers and Golgi-resident proteins60,65,66. In one of the proposed models, the COG complex is thought to tether and assemble various Golgi-trafficking molecules on the Golgi cisternae, with the COG subunits binding to different and specific SNARE and tethering proteins based on the cisternae the cargo is destined to be docked to66. In fact, it was found that lobe A COG subunits preferentially interact on the cis-Golgi segment, while lobe B COG subunits have been found to interact more predominantly with trans-Golgi factors5,66. While this model does not explain how these docking stations are signalled into formation, a different model sidesteps this issue by positing the idea that the COG complex directly recognizes specific protein sites or vesicular coats. Golgins and other coiled-coil tethers would attract various vesicles, and the COG complex subunits would only reel in proteins that have a direct connection with the acceptor cisternae5,56. A final model focuses on the role that SNARE complexes have on the fusion of vesicles with the acceptor membrane, and the COG subunits simply facilitate this role by stabilizing the SNARE complexes5,58. It is possible that the actual mechanism of COG complex-mediated Golgi trafficking is a hybrid of the proposed models, with a more complex and more specific interaction between the subunits, SNARE proteins, vesicular tethers, and the Golgi cisternal membranes.

Lobe A Lobe B B

Figure 2.4: Schematic demonstrating the COG complex and ER-Golgi trafficking. (A) Model of the COG complex demonstrating its two lobes and the subunits in each. Interactions between subunits are represented by connecting lines. (Adapted from Ungar, et al. 2005. J Biol Chem.) (B) Anterograde and retrograde trafficking between the ER and the Golgi. COPII-coated vesicles proceed through anterograde trafficking from the ER into the ERGIC before being passed along to the Golgi complex. COPI-coated vesicles undergo retrograde trafficking throughout the Golgi complex cisternae as well as from the Golgi to the ER. The COG complex facilitates the intra- Golgi and Golgi-ER retrograde trafficking by interacting with COPI. The trafficked vesicles at the Golgi have different contents that include glycosylation enzymes that need to be recycled and replenished throughout the Golgi cisternae (Adapted from Koegler, et al. 2010. Traffic67)

2.4.2 COG complex-associated congenital disorders of glycosylation

As mentioned previously, CDGs are a genotypically and phenotypically heterogeneous group of disorders that result from disruptions to the process of glycosylation. Due to the role of the COG complex in facilitating the glycosylation process at the Golgi, after the glycans have already attached to the nascent proteins and exit the ER, COG-CDGs are classified as type II68. Mutations in seven out of the eight subunits of the COG complex (excluding COG3) have been found to cause CDGs. The first identified subunit with a CDG-causing mutation was COG7, which demonstrated reduced glycosylation activity and altered Golgi trafficking in patient- derived fibroblasts69. A recently-generated COG7-CDG Drosophila model exhibited phenotypes similar to those of patients, such as neuromotor dysfunction, and was found to have defects in N- glycosylation70.

The first identified COG5-CDG case was of a patient that presented with mild cognitive delay and motor dysfunction, due to a homozygous intronic substitution in COG5 that led to a significantly reduced concentration of its protein71. Analysis on patient-derived fibroblasts revealed delayed retrograde Golgi-to-ER trafficking that was rescued by transfecting them with wild-type healthy forms of COG571. At least six more patients with COG5-CDG have been published since, demonstrating a range of mild to severe cognitive delay, delayed motor function, microcephaly, and hypotonia72. Most of the patients have short stature, and a couple presented with liver disorders, deafness, and blindness, however, retinal degeneration and ocular health were not mentioned72. One of the patients had a frame insertion mutation and the missense p.M32R mutation in COG5 that we identified as one of the two mutations inherited by the siblings in Family 3 (See section 2.2.2). The p.M32R missense mutation was predicted to be incredibly damaging due to its location in a highly conserved region of COG572. The protein concentration levels of COG5 in COG5-CDG patient-derived fibroblasts were found to be markedly reduced, along with concentration levels of COG7, emphasizing the strong bond that those two subunits share in lobe B of the COG complex72,59.

2.4.3 The COG5 subunit

COG5 was initially discovered as part of a Golgi transport complex (GTC) and was referred to as GTC-90 due to its size in kiloDaltons (kDa)73. It was found that inhibiting the activity of COG5 (or GTC-90) by using an antibody against it led to the inhibition of in vitro Golgi transport

21 between the cis- and medial-cisternae, which was the first indication of the broader role this protein plays in Golgi trafficking processes73. The same study additionally confirmed the colocalization of COG5 to the Golgi complex73. A Drosophila homolog of COG5 protein has been identified to be coded by the four-way stop (fws) gene, which plays a critical role in spermatogenesis74. Fws protein is also localized to the Golgi complex and is crucial for the formation of the Golgi-derived acroblast, which develops into the acrosome during spermatogenesis74,75. The acrosome is vital for fertility as it is the enzyme that allows the sperm to digest the outer membrane of the ovum75. Not only is the Drosophila COG5 homolog important for cytokinesis in spermatogenesis, but defects in Fws protein also disrupted the normal architecture of the Golgi74. The studies on the Drosophila homolog of COG5 protein emphasize the importance of the protein not only for Golgi transport but also maintaining Golgi structure and morphology. The Drosophila model also draws interest into why loss-of-function mutations in fws only result in spermatogenesis disruptions, while not presenting any other evident phenotypes.

In mammalian studies, validated COG5-deficient cells also demonstrated a marked reduction in COG6 and COG7, while COG1, COG2, and COG3 showed normal expression levels as well as normal Golgi colocalization76. This is in line with the prevailing view that the COG complex is separated into independently-functioning subcomplexes, lobe A (COG1-4) and lobe B (COG5- 8), that are interconnected through linkage of COG1 with COG8. Previous knockout studies of COG1 and COG2 demonstrated widespread expression level reductions of a subset of Golgi integral membrane proteins called GEAR proteins. However, COG5-deficient cells only resulted in a reduction of one Golgi SNARE protein, GS15. Additionally, this reduction of GS15 was rescued through the supplementation of COG5-deficient cells with wild-type COG5 cDNA76. This demonstrates an aspect of specificity for the different COG subunits in their interactions with Golgi components.

COG5-deficient cells also presented a more dilated architecture of the cisternae but did not demonstrate any global glycosylation defects as were presented in COG1- and COG2-deficient cells76. This is despite the finding that mutations in the yeast homolog of COG5 protein showed abnormal glycosylation patterns of a cell wall glycan77. These results hint at the possibility that COG5’s role in Golgi function may not be solely linked to the Golgi’s glycosylation and posttranslational modification function. Indeed, a recent study highlighted the COG complex’s

22 glycosylation-independent role in cellular function78. While investigating COG-knockout(KO) cell lines for lobe A and lobe B, the expression levels of many glycosylation enzymes were significantly decreased for all KO groups. However, the knockout of those same glycosylation enzymes, such as MGAT and GALE, did not mimic some of the phenotypes displayed by the COG KO cells. For instance, COG-KO cells demonstrated severe fragmentation of the Golgi complex and formed enlarged vacuolar endo-lysosomal structures, both of which were not present in MGAT- and GALE-KO cells. Protein sorting and secretory trafficking were also impaired only in the COG-KO cells. Finally, retrograde Golgi trafficking was significantly more impaired in COG-KO cells than in MGAT- and GALE-KO cells78. These findings all point to a role played by the COG complex that is beyond its effect on glycosylation. Considering that previous studies did not show glycosylation defects when only COG5 was affected, that would indicate that COG5 likely plays a more significant role in maintaining Golgi structure and facilitating retrograde trafficking. However, studies looking into the specific role the COG5 subunit plays are missing from the literature and the only studied effects are linked to the general disruption of the COG complex’s lobe B.

2.5 ER stress and the Unfolded Protein Response

Several components within the cell work together to ensure the proper processing and folding of newly-synthesized proteins, including molecular chaperones such as immunoglobulin binding protein (BiP), lectins such as calnexin, and foldases such as protein disulfide isomerase (PDI)79. A third of all cellular proteins are synthesized in the ER80, and so it hosts most of the processes responsible for proper protein folding, including the pathway to degrade improperly-folded proteins referred to as the ER-associated degradation (ERAD) pathway81. If the ER is unable to maintain protein homeostasis (proteostasis) due to an abundance of aggregated, incorrectly- folded proteins, it will undergo ER stress. This will activate the unfolded protein response (UPR) with the goal of restoring proteostasis81.

2.5.1 Protein folding and aggregation

A nascent protein is not functional until it achieves proper folding. This folding is driven by hydrophobic interactions that occur as the polypeptide is placed in an aqueous environment after translation82,83. Several safeguards to maintain native protein folding are put in place as proteins generally undergo various disruptions to their structure throughout their lifespan83. Disruptions to

23 protein folding can result due to mutations or a deficiency in the protein’s binding partners or molecular chaperones84. The cell relies on molecular chaperones in the lumen of the ER to assist in folding and trafficking of nascent proteins85,86. Other factors such as radiation, reactive oxygen species, and hypoxia, can also disrupt protein modification and processing in the ER, leading to ER stress87.

Unfolded or partially unfolded proteins have exposed hydrophobic patches that molecular chaperones recognize and bind to88. Due to their protective function of proteins during times of cellular stress, molecular chaperones have been referred as heat shock proteins (HSP), as they were discovered to be upregulated in response to heat shock stress83. HSPs that bind to unfolded proteins to stabilize them and prevent their aggregation are referred to as holdases. HSPs that bind to proteins and assist with their folding are called foldases, and those that help untangle the aggregation of misfolded proteins are called disaggregases89.

One of the most important holdases is BiP, an HSP that binds to exposed hydrophobic patches of partially unfolded proteins to reduce aggregation-prone sequences. It enacts its function by binding to and subsequently releasing the unfolded protein, in a cycle. With each release, the protein is allowed the opportunity to correctly fold in the aqueous environment through hydrophobic interactions of its exposed hydrophobic patches90.

In addition to stabilization by holdases, many proteins require an active participant in their folding and the two most prominent foldases are PDI and peptidyl-prolyl isomerase (PPI). PDI catalyzes the formation of disulfide bonds, whereas PPI is responsible for the isomerization of peptide bonds near proline residues91. PDI actually encompasses a group of at least 19 PDI family members that reside in the ER, but many of them remain elusive with regards to their function92. PDI members differ with regards to the types of proteins they interact with, where some, like PDIA1, interact with glycosylation and non-glycosylated proteins alike, while members such as PDIA3 exclusively interact with glycosylated proteins92. HSP70 chaperones also have disaggregase activity as they have been found to disaggregate unfolded, inactive, insoluble proteins and reactivate their function93. However, the mechanism by which this happens remains unknown. If protein misfolding persists, the proteins are targeted by the ER for degradation.

Due to the sensitive nature of protein folding, the ER has a regulatory quality control mechanism that mitigates problems arising due to misfolded proteins, called the ER-associated protein degradation (ERAD) pathway84. Misfolded proteins are recognized by their exposed hydrophobic patches or by an attached glycan that has not been adequately modified84,94. These proteins are then tagged for degradation through the attachment of ubiquitin by ubiquitin ligase. This acts as a signal for further ubiquitin tags to be added, creating a polyubiquitin chain that is recognized by the proteasome, a cytosolic protein complex responsible for protein degradation95. The polyubiquitin chain drives the protein to exit the ER into the cytosol and enter the proteasome, which proteolytically degrades the protein into amino acids that can be recycled by the cell to translate new proteins. If the amount of misfolded proteins is too high for the ER to send to the ERAD pathway efficiently, then this will cause the ER to stress, activating the UPR96.

2.5.2 UPR: Restoring proteostasis and signalling apoptosis

Under cellular and ER stress, resulting from the accumulation of toxic, misfolded proteins, the UPR is initiated to restore proteostasis96. The UPR activates molecular chaperones and transcriptional factors to achieve three goals, which are to stop further protein translation from proceeding, to increase production and recruitment of molecular chaperones to assist with protein folding and to degrade and eliminate the accumulated misfolded proteins properly97. There are three separate arms of the UPR that act in parallel and are activated independently of one another. Those three arms are referred to by the name of their initiating proteins or sensor domains: protein kinase RNA-like ER kinase (PERK), inositol-requiring protein 1α (IRE1α), and activating transcription factor 6 (ATF6)96 (See Figure 2.5). The three UPR stress sensors are activated through various mechanisms described below but are thought to be inhibited from activation through interactions with BiP98. BiP binds to the luminal domains of the UPR sensors but has a strong affinity for misfolded proteins. As a result, when the levels of misfolded proteins rise in the ER, BiP dissociates from the UPR sensors, resulting in UPR sensor activation98. However, another proposed model of activation characterizes the UPR sensors as ligand-driven, having a strong affinity themselves to misfolded proteins87. It is proposed that a hybrid model of both direct BiP dissociation and indirect peptide binding regulate and mediate UPR sensor activation99.

Figure 2.5: Schematic demonstrating the three branches of the unfolded protein response (UPR) and their respective function. Upon an aggregation of misfolded proteins in the endoplasmic reticulum (ER), the ensuing ER stress activates the UPR, which is composed of three interdependent branches that act in parallel96. The first two arms, PERK and IRE1, are activated when misfolded proteins bind to their luminal domain87. IRE1 acts to degrade ER-localized mRNA to reduce entry of nascent proteins in the ER100. PERK works towards reducing global protein synthesis101. ATF6 is transported to proteasomes at the Golgi apparatus, where it is cleaved and activated. ATF6 upregulates transcription of chaperones that assist with degradation of misfolded proteins102. Under severe ER stress, all three UPR arms initiate the termination response or apoptosis, which involves the upregulation and activation of pro-apoptotic factors.

2.5.2.1 PERK activation

While the particular pathways and temporal activation of UPR sensors are still under investigation, recent evidence has identified a peptide substrate, P16, that binds to the PERK luminal domain, and competitively competes with misfolded proteins for PERK-binding87. PERK is a transmembrane ER-protein with domains on both the luminal and cytosolic side of the ER101. Upon an increase in abundance of misfolded proteins in the ER lumen, PERK’s luminal domain dissociates from P16 and binds to misfolded proteins. This will initiate the oligomerization of PERK that is succeeded by the autophosphorylation of its cytosolic kinase domain87. As a result, its kinase activity functions to phosphorylate the eukaryotic initiation factor-2α (eIF2α), which regulates transcription pathways to stop global protein translation87,101. The reduction in protein synthesis also prevents nascent proteins from entering the ER until the UPR can restore proteostasis103. Under instances of oxidative stress, PERK also phosphorylates Nrf2 transcription factor which acts to upregulate factors that help reduce intracellular reactive oxygen species (ROS) that induce genotoxic stress104. The silencing of PERK was found to trigger DNA damage induced by oxidative stress and an accumulation of ROS104.

Phosphorylation of eIF2α results in the prioritized translation of activating transcription factor 4 (ATF4), which in turn, regulates the transcription of various genes responding to ER stress. One such gene is CHOP, which suppresses pro-survival molecules and activates ER oxidoreductin 1 (Ero1)α, triggering calcium-mediated programmed cell death (apoptosis)105. The ER is a prominent storage site for calcium, which contributes to protein folding and acts as a signalling messenger for various processes in the body such as hormone secretion, synaptic transmission, and gene transcription106. An imbalance in ER calcium levels can trigger ER stress and lead to an oversaturation of calcium in the p, resulting in its rupture and release of pro-apoptotic factors106. In tumor cells, the downstream activation by PERK of eIF2α and ATF4 also results in the induction of LAMP3 (lysosomal-associated membrane protein 3), a lysosomal-membrane- resident protein that is involved in autophagy107,108. The PERK/ATF4/LAMP3-arm was found to be integral to the recruitment of DNA damage repair signalling of γ-H2A.X and promotes resistance of cancer cells to radiotherapy109. The recruitment of γ-H2A.X downstream of PERK activation, however, conflicts with an earlier study that found that silencing of PERK leads to increase of γ-H2A.X recruitment104. This may hint at a modulatory effect of PERK signalling on DNA damage repair signalling.

Another gene upregulated by phosphorylated eIF2α is growth arrest and DNA-damage-inducible 34 (GADD34), which acts in a negative-feedback loop to dephosphorylate eIF2α and reduce transcription of CHOP110. The knockdown of GADD34 was also found to upregulate LAMP3 expression in cancer cells109. ATF4 has also been found to facilitate the trafficking of another UPR sensor, ATF6, from the ER to the Golgi where it is cleaved and activated111. Activation of ATF6 is discussed in more detail below, but this finding challenges the view that the three arms of UPR are activated in parallel independently and explores an integrated model of the UPR in responding to ER stress and facilitating apoptosis.

PERK’s role in disease models is under investigation, mainly as a possible therapeutic target. In autosomal dominant RP, PERK has been found to reduce photoreceptor cell death that occurs due to mutant rhodopsin112. PERK inhibition in a transgenic rat model of the mutant rhodopsin- caused RP led to increased photoreceptor death and reduced ERG activity112. In the same study, it was also found that inhibiting the dephosphorylation of eIF2α, which would maintain phosphorylated-eIF2α’s UPR activation, improved photoreceptor survival112. In line with these findings, it was found that a deficiency in PERK due to autosomal recessive mutations, results in wide-ranging developmental defects referred to as the Wolcott-Rallison syndrome113. Symptoms of Wolcott-Rallison include ones that are shared by the Clinical Case B patients, such as skeletal dysplasia, growth retardation, and microcephaly114. However, Wolcott-Rallison syndrome also commonly manifests other symptoms including diabetes, liver failure, hypothyroidism, and intellectual delays114. Loss of PERK function in cell culture was found to impair ER-to-Golgi anterograde trafficking as well as the transport of misfolded proteins from the ER into the cytosol for degradation. Additionally, PERK loss-of-function results in impairment of proteasomal degradation of proteins under the ERAD pathway115.

More success has been demonstrated in treating neurodegeneration by inhibiting PERK activity. For instance, elevated levels of phosphorylated-eIF2α have been found in brains of Alzheimer’s patients in association with an increase in amyloid precursor proteins. Through the suppression of PERK kinase activity, the phosphorylation of eIF2α is prevented, and this has demonstrated an improvement in spatial memory and synaptic plasticity in mouse models of familial Alzheimer’s disease116. Another therapeutic investigation on PERK was performed on a mouse model of prion disease, a neurodegenerative disease caused by abnormal misfolding of prion proteins in the brain. It was found that oral treatment of a blood-brain barrier-permeable PERK

28 inhibitor (GSK2606414) to the prion disease-mouse models led to the rescue of cognitive deficits and stopped the clinical progression of the disease117. Some side effects of PERK-inhibitor treatment did arise in the mice including weight loss and dysregulation of insulin117. Nevertheless, these investigations provide a promising prospect of targeting the PERK/phosphorylated-eIF2α pathway as a therapeutic option for diseases led by ER stress activation118.

2.5.2.2 ATF6α activation

ATF6 is a transcription factor bound to the ER membrane and acts as a sensor for ER stress. As a member of the activating transcription factor/cAMP response element binding protein (ATF/CREB) family, ATF6 incorporates a basic leucine zipper (bZIP) DNA-binding domain119. A functional bZIP domain permits cleaved ATF6 to upregulate transcription of ER stress genes such as CHOP120. The activation of ATF6 commences as it is transported from the ER membrane to the Golgi complex, where it gets cleaved by two proteasomes111. The cleaved ATF6 subsequently enters the nucleus and functions to upregulate transcription of ER chaperones such as PDI and several UPR target genes, including those for BiP and ERAD pathway-associated factors99,102,119. BiP takes on many roles in the process of ATF6 activation, the first of which is binding to the luminal domain of ATF6, inhibiting its activity and preventing it from being translocated to the Golgi121. As misfolded proteins increase, BiP dissociates from ATF6 to bind to the proteins and assist with their folding99.

ATF6 mutations have been found in patients diagnosed with achromatopsia and cone-rod dystrophy. Achromatopsia is an inherited vision disorder that results in a complete loss of colour vision, photophobia, and decrease visual acuity, due to the specific dysfunction of cone photoreceptors122. Identified ATF6 mutations in the luminal domain were found to impair ER-to- Golgi trafficking while mutations that cause defects in ATF6’s bZIP domain results in loss of ATF6 function and subsequently, an increase in apoptosis120. Additionally, achromatopsia patients due to ATF6 loss-of-function mutations have also shared features such as malformation of the fovea and retinal developmental deficits102. It was found that ATF6 plays an embryonic developmental role in differentiating stem cells towards a mesodermal differentiation as well as playing an essential role in ER maturation102. This expands the role of the UPR beyond restoring ER proteostasis to potentially playing another pivotal role in early embryonic development and

29 differentiation. Investigations into potential therapeutic approaches for achromatopsia symptoms include rescuing ATF6’s transport to the Golgi for luminal domain mutations and gene therapy to introduce a functional bZIP domain in mutations that lead to bZIP domain disruptions120.

2.5.2.3 IRE1α activation

IRE1 is a UPR stress sensor with two isoforms, α and β. However, this review will only focus on IRE1α, which is ubiquitously expressed, while IRE1β is only expressed in respiratory and gastrointestinal tracts100. IRE1 is an ER-transmembrane protein with a luminal domain that, similarly to PERK, has an affinity for misfolded proteins123. Specific activation of IRE1 or PERK by binding to misfolded proteins is further complicated by the affinity of BiP to misfolded proteins. However, their specific activation likely depends on varying preferences to the type of peptides they bind to. For example, IRE1 tolerates prolines and histidine, while BiP was found to avoid interactions with those amino acids. Moreover, BiP tolerates specific amino acids such as serine and threonine, which IRE1 does not approach100,123.

The binding of IRE1 to misfolded proteins initiates its oligomerization which subsequently leads to the auto-phosphorylation of its kinase domain. This activation of IRE1 regulates transcription pathways to over-express molecular chaperones to assist with ER protein folding. To that end, IRE1’s activation results in cleavage of many mRNAs in a process referred to as the regulated IRE1-dependent decay of messenger RNAs (RIDD). RIDD’s goal is to reduce protein entry into the ER to lessen the burden of misfolded proteins. Specifically, IRE1 also initiates the splicing of XBP1 mRNA, which leads to the encoding of XBP1s, a transcription factor that results in downstream transcription of genes that assist with restoring ER proteostasis100. Some of the UPR genes upregulated by XBP1s are involved in the ERAD pathway, in protein quality control, and in disulfide linkage111.

As a potential therapeutic target, IRE1 activation was investigated on mammalian cells expressing both wild-type and P23H mutant rhodopsin. The P23H rhodopsin mutation causes the most common form of autosomal dominant RP, with the misfolded mutant rhodopsin resulting in photoreceptor cell death. While ER stress-induced activation of all three arms of the UPR decreased levels of both wildtype rhodopsin and misfolded mutant rhodopsin, the selective activation of IRE1 signalling only promoted degradation of misfolded P23H mutant rhodopsin10. However, this does not represent a comprehensive therapeutic finding since IRE1 signalling was

30 not able to restore delivery of the mutant rhodopsin to its target to prevent photoreceptor death10. It is interesting to note that overexpression of BiP in a transgenic rat model with P23H mutant rhodopsin restored its visual function by reducing levels of cleaved ATF6, as well as both phosphorylated eIF2α and CHOP, which are upregulated under PERK activation124. BiP overexpression was also found to reduce apoptosis, likely by forming a complex with proapoptotic factors caspase-7 and caspase-12 and preventing their activation124,125. These findings support an integrated system model between the arms of the UPR and their molecular components, as all three of the IRE1, ATF6, and PERK arms contribute to cell survival, protein folding, and maintaining normal function.

2.5.2.4 Apoptosis signalling

Apoptosis is an essential and highly conserved process of programmed cell death that protects organisms from abnormal cell behaviour or DNA damage that may lead to severe mutations or tumor development126. Defects in the process of apoptosis have been linked to cancer, neurogenerative diseases, and various other disorders. All three arms of the UPR are involved in signalling programmed cell death, or apoptosis. Activated IRE1 results in splicing of XBP1 mRNA and an increase of its RNase activity, both of which activate apoptosis. PERK activation also induces apoptosis by upregulating proapoptotic factors like CHOP and GADD34127. ATF6 mediates apoptosis by upregulating transcription of proapoptotic factors such as WW domain binding protein 1 (WBP1) and CHOP128,129. While CHOP is transcriptionally upregulated for apoptotic signalling, it also upregulates downstream genes that can facilitate apoptosis, or act as an inhibitory feedback loop and promote survival129.

Apoptosis almost always begins with the activation of caspases (cysteine-dependent aspartate- specific proteases), which are protease enzymes that cleave target proteins and sometimes other caspases in order to activate them126. During apoptosis, caspases disrupt mitochondrial function and actin polymerization, leading to the disruption of the cell’s structural morphology and integrity126. As mentioned previously, the disruption of the mitochondrial function results in its swelling and rupture, releasing pro-apoptotic factors into the cytosol like cytochrome c, which subsequently activates more caspases. ER stress also initiates a cytochrome-c-independent pathway of caspase-mediated apoptosis whereby it initiates cleavage of the ER-membrane-bound caspase 12, releasing it into the cytosol. Activation of caspase 12 leads to a cascade of caspase 9

31 then caspase 3 activation130. Caspase 3 activation marks the beginning of the “execution pathway” whereby endonucleases degrade chromosomal DNA and proteases degrade nuclear and cytoskeletal proteins. The resultant morphological changes will cause nuclear fragmentation and the formation of apoptotic bodies which are eventually phagocytosed131.

2.5.3 ER stress and the UPR in retinal degeneration

The ER plays an essential role in protein synthesis, folding, and modification to ensure its proper function. It is thus likely to play a significant role in causing many diseases, including those of retinal dystrophy and degeneration. In fact, ER stress has been identified to be involved in a type of retinal degeneration that results from a T17M-mutation in rhodopsin, which is associated with autosomal dominant RP132. As mentioned earlier in section 2.1, rhodopsin is the most abundant protein in the outer segments of rod photoreceptors, and plays a critical role in the visual transduction pathway, so its proper folding and function are critical. In transgenic mouse models expressing the T17M rhodopsin mutation, rhodopsin was found mislocalized in the outer nuclear layer, unable to be shuttled into the outer segments132. Additionally, several UPR factors and chaperones such as BiP, ATF4, eIF2α as well as apoptotic-inducer proteins such as CHOP were all upregulated. Apoptosis was activated as indicated by the release of mitochondrial proapoptotic factors and the increased expression of caspase-3 and -12132. These results indicate that the T17M rhodopsin mutation prevented its proper folding and localization and this correlated with induced UPR in photoreceptors, culminating in their death and causing retinal degeneration. Similar results were found in an analysis of other autosomal dominant RP models expressing the S334X rhodopsin mutation and the P23H mutation133,134.

Mutations in tubby gene family members have been found to cause retinal dystrophies such as RP, LCA, and rod-cone dystrophy. An investigation of mice expressing mutant Tub gene demonstrated a mislocalization of both rhodopsin and M-opsin, which is found in cone photoreceptors135. This mislocalization also correlated with the upregulation of several UPR markers including IRE1, eIF2α, ATF6, and BiP, indicating the involvement of ER stress in other forms of retinal dystrophy135. A mutation of another tubby gene family member, TULP1, which causes LCA, has also linked the misfolding of the mutant TULP1 protein with activation of the PERK and IRE1 arms of UPR and upregulation of proapoptotic gene CHOP136.

In an animal model where UPR was actively induced through the injection of tunicamycin intraocularly, UPR markers were upregulated in addition to inflammatory markers and caused a marked loss of ERG activity and a significant amount of photoreceptor death137. Additionally, tunicamycin-induced UPR activation led to the activation of microglia, which are macrophage cells that are presumably acting to reduce inflammation but could lead to phagocytosis of photoreceptors137,138. In cultured porcine retinal explants, ER stress was induced by prolonged dim-light illumination and resulted in increased expression of BiP and proapoptotic factors CHOP and caspase-12139. This activation of ER stress resulted in reduced levels of rhodopsin and increased photoreceptor degeneration through autophagy139. These studies present a myriad of ways through which ER stress can cause retinal dystrophy but highlight the prevalence of ER stress as a shared cause for retinal degeneration. The recent push towards having ER stress and the UPR as a possible therapeutic target is thus warranted, but it comes with its own set of challenges.

2.5.4 Therapeutic options targeting ER stress and the UPR

The possibility of therapeutic options targeting ER stress and the UPR has expanded over the last few years due to the development of various methods gene therapy delivery, including lentivirus, adenovirus, and adeno-associated vectors (AAV). As mentioned previously (see section 2.5.2.3) overexpression of BiP was found to restore visual function in a model of autosomal dominant RP caused by a P23H mutation in rhodopsin124. A recent study delivered BiP through an AAV cultured human RPE cells and found that it promoted RPE survival by upregulating the adaptive PERK and ATF6 arms of the UPR while downregulating CHOP140. PERK regulation, as mentioned in its titular section, has also been of great interest as a therapeutic target and has shown success in prion and Alzheimer’s disease models116,117.

Therapy through chemical modulation of ER stress and the UPR has also been investigated through the use of modulators such as sodium 4-phenylbutyrate (4-PBA) and tauroursodeoxycholic acid (TUDCA)141. The use of PBA and TUDCA were able to control UPR expression by reducing the expression of BiP, CHOP, and XBP1s while reducing photoreceptor degradation in mutant photoreceptor-derived cells142. In retinal explant models of Bardet-Biedl syndrome, a form of retinal degeneration, it was found that photoreceptor death resulted from overexpression of UPR-induced caspase-12143. Through the use of a caspase-12 inhibitor, Guanabenz,

33 photoreceptors were rescued from degradation by upregulating expression levels of BiP, phosphorylated-eIF2α, and CHOP, which reduced caspase-12-mediated apoptosis143. In another study, BiP was directly induced in vitro and in vivo in cells after UPR was activated with tunicamycin and it was found that BiP activation reduced CHOP protein expression and the levels of retinal cell death144.

Direct chemical induction of the ATF6 or IRE1 arms of the UPR in cells expressing wild-type or mutant rhodopsin found their activation reduced the expression levels of mutant rhodopsin but maintain steady-state levels of the wildtype form145. ATF6 activation thus helped reduce misfolded protein levels that may cause apoptosis of cells but did not help facilitate the transport of misfolded rhodopsin to the plasma membrane145. The direct induction of the PERK arm, however, reduced both mutant and wildtype forms of rhodopsin, as well as other proteins in the cell, in accordance with its role in reducing global protein synthesis145. These studies hint towards a potential therapeutic role for targeting ATF6 and IRE1 arms of the UPR in reducing the amount of misfolded or aggregate rhodopsin in certain disorders of retinal degeneration, but that the targeting of PERK may lead to unintended side effects. Due to the complexity of the ER stress and UPR pathway, future research should focus on therapeutic options that can target specific components of the UPR, to alleviate symptoms and improve cellular function, while minimizing side effects.

Rationale and Hypothesis

Five patients are presenting with a multi-systemic disorder that is affecting their development and most prominently, their vision. The causal mutations in SRD5A3 and COG5 could be passed down or shared with other individuals and establishing screening methods for these genes is necessary. The five patients are a small representation of a larger group of individuals suffering from a similarly wide-ranging disorder affecting their vision and living without treatment or a known mechanism to target.

Whether it is Leber’s congenital amaurosis, retinitis pigmentosa, or congenital disorders of glycosylation, all of these diseases have been associated with other mutations found in SRD5A3 and COG5, but a molecular mechanism of disease has not been established for any of them. Additionally, many studies have linked ER stress and activation of the unfolded protein response to a wide range of diseases such as retinal degeneration, Alzheimer’s disease, Parkinson’s disease, bipolar disorder, diabetes, and inflammation, but a targeted approach for treatment have yet to be developed.

As a polyprenol reductase, the role that SRD5A3 plays in N-glycosylation has been clearly established, and the identified nonsense mutation p.W19X has already been associated with retinal dystrophy. However, despite SRD5A3’s ubiquitous expression, the homozygous expression of the mutation is not lethal, and only affects a limited number of organs. The mutated SRD5A3 is likely to induce the UPR through the disruption of N-glycan processing in the ER. Assessing the effect of expressing the mutated SRD5A3 protein in vitro on the regulation of the UPR arms can help indicate the particular defects caused by the mutation. Identifying the UPR arms and markers that are activated or inhibited by a mutated SRD5A3 will hint at how proteins are being disrupted and assist future investigations into assessing targeted therapeutic options.

The role that the COG complex plays in Golgi trafficking and assisting with protein glycosylation has also been well-characterized; however, the direct interactions of the COG5 subunit remain understudied. Most investigations categorically look at broad effects of lobe A or lobe B disruption of the COG complex, due to the structural interdependence of the subunits in each lobe. However, in this study, the goal is to study the COG5 subunit independently of other subunits, to categorize the direct role of mutations in this protein on disrupting cellular function.

This is further of importance due to the absence of a phenotype in the parents expressing a heterozygous mutation, but the presence of severe and wide-ranging phenotypes in the children with a compound heterozygous mutation in COG5. Any mutation in the COG complex is likely to cause a disruption to Golgi trafficking and glycosylation, leading to an aggregation of misfolded proteins at the ER and perhaps an absence of essential ER-resident proteins due to mistrafficking. This will likely result in ER stress and the activation of the UPR, but no study has looked at the role of the COG complex on UPR induction. This investigation will not only assess the effect of the COG5 mutations in upregulating various UPR and ER stress markers but will also investigate the interplay and combinatory effect of both COG5 mutations being expressed in vitro together. These experiments will offer illuminating insights into an essential Golgi trafficking molecule and provide potential therapeutic targets to investigate.

3.1 Hypothesis

SRD5A3 and COG5 proteins play an integral role in protein glycosylation and trafficking, and mutations in their respective genes will lead to an increase in ER stress and activation of the UPR, resulting in photoreceptor cell death and retinal degeneration.

3.1.1 Objectives

1. Generate DNA constructs expressing wildtype and mutant forms of COG5 and SRD5A3, tagged on the N-terminus with GFP and another set tagged with 6x-His.

2. Assess in vitro localization of wildtype and mutant COG5 and SRD5A3 proteins.

3. Investigate activation of ER stress and UPR markers in cells transfected with wildtype or mutant COG5 or SRD5A3 plasmids through Western blotting.

4. Measure levels of Golgi dispersal resulting from mutant COG5 protein expression and expression of both COG5 mutants, compared with wildtype COG5 expression.

5. Assess and compare levels of DNA damage in cells transfected with wildtype, mutant, or double-mutant COG5 plasmids.

6. Target the ER stress or UPR marker that is most upregulated in double-mutant COG5 condition and assess rescue of the phenotype.

Materials and Methods 4.1 Mice

Mice used throughout this project were all of the C57BL/6J strain (Charles River). They were handled in accordance with the ethical and legal requirements outlined by Ontario’s Animals for Research Act and the federal Canadian Council on Animal Care. All experiments performed were approved by the University Health Network Research Ethics Board and was overseen by the Animal Resources Centre at the Krembil Research Institute, under protocol 3381.5.

4.2 DNA constructs

Constructs of human COG5 (cDNA clone MGC: 87389 IMAGE: 4374289) and human SRD5A3 (cDNA clone MGC: 3099 IMAGE: 3350037) that were tagged at the N-terminus with Green Fluorescent Protein (GFP) were obtained from The SickKids Proteomics, Analytics, Robotics & Chemical Biology Centre (SPARC BioCentre). A complete list of DNA plasmids used is listed in Table 2.1, along with the shortened name they are referred to throughout this manuscript.

Table 4.1: List of all DNA plasmids used along with the shortened name they are referred by.

DNA Plasmid Description Shortened Name pcDNA6.2-N-EmGFP-hCOG5(wt) GFP-COG5 pcDNA6.2-N-EmGFP-hCOG5(M32R) GFP-COG5(M32R) pcDNA6.2-N-EmGFP-hCOG5(S777Q_fs*14) GFP-COG5(S777Q) pcDNA6.2-N-EmGFP-hSRD5A3(wt) GFP-SRD5A3 pcDNA6.2-N-EmGFP-hSRD5A3(W19X) GFP-SRD5A3(W19X) pcDNA6.2-N-6xHis-hCOG5(wt) His-COG5 pcDNA6.2-N-6xHis-hCOG5(M32R) His-COG5(M32R) pcDNA6.2-N-6xHis-hCOG5(S777Q_fs*14) His-COG5(S777Q) pcDNA6.2-N-6xHis-hSRD5A3(wt) His-SRD5A3 pcDNA6.2-N-6xHis-hSRD5A3(W19X) His-SRD5A3(W19X) pT2K-CAGGS-eGFP 146 GFP [control]

4.2.1 Generation of mutant clones

GFP-tagged COG5 and SRD5A3 constructs underwent site-directed mutagenesis to insert mutations identified in human patients: COG5 c.95T>G (p.M32R), COG5 c.2327dupT (p.S777Q_fs*14), and SRD5A3 c.57G>A (p.W19X). This was done by preparing a mutant strand synthesis reaction mixture of 10μl 5X GC reaction buffer, 100ng of dsDNA template, 2.5μl of 10μM forward primer, 2.5μl of 10μM reverse primer, 1ul of 10mM dNTP, 1.5μl of DMSO, 0.5μl of Phusion DNA polymerase (NEB), and up to 50μl of nuclease-free, sterile, double-distilled water (ddH2O). Polymerase chain reaction (PCR) cycle was run in two segments: 1) 1 cycle of 95°C for 30 seconds; 2) 18 cycles of a) 98°C for 30 seconds, b) 60°C for 1 minute, c) 68°C for 1 minute per kilobase of plasmid length. At the end of the reaction, samples were placed on hold at 4°C or on ice until cool. The parental DNA was digested by adding 1μl of Dpn I restriction enzyme (NEB) to the 50μl reaction mixture and incubated at 37°C for 1 hour. DNA was transformed into competent DH5ɑ Escherichia coli (E. coli) cells and plated on LB-agar plates as described in section 4.2.2. below.

For Western blotting applications, the GFP was excised out and replaced with a 6xHistadine tag instead. This was performed by creating primers that include a restriction enzyme as well as the 6xHistadine sequence and running a PCR to amplify the gene insert (COG5 or SRD5A3), attached to the 6xHis tag. The PCR ran in three segments: 1) 1 cycle of 98°C for 30 seconds; 2) 35 cycles of a) 98°C for 30 seconds, b) 71°C for 30 seconds, c) 72°C for 30 seconds per kilobase of plasmid length; 3) 72°C for 10 minutes. The PCR product was run on a 1% agarose gel at 120V for 20 minutes, and a band of correct size was confirmed using a UV gel imager. The band was excised out of the gel, and the DNA was extracted using QIAquick Gel Extraction Kit (Qiagen). The extracted DNA was polyadenylated by mixing 30μl of the DNA with 5μl of 10X ThermoPol® Buffer (NEB), 1μl of 10mM dATP (Thermo Scientific), 1μl of Taq DNA polymerase (NEB), and 13μl of ddH2O, and incubating the mixture at 72°C for 30 minutes. The DNA was purified using the QIAquick PCR Purification Kit (Qiagen). The PCR-purified product was ligated into pGEM®-T Easy Vector by mixing 6μl of DNA insert with 1μl of the vector, 1μl of T4 DNA ligase (NEB), and 2μl of rapid ligation buffer (Thermo Scientific). The ligation product was transformed into competent DH5ɑ E. coli cells and plated on LB-agar plates that have been smeared with a mixture of 80μl X-Gal and 16μl IPTG for white-blue selection. White

39 colonies, indicating that DNA was inserted into the vector, were picked and inoculated in LB broth for DNA plasmid preparation, as described below in section 4.2.2. After confirming the addition of the 6xHis tag in the DNA sequencing results, the insert was digested out of the vector, using the restriction enzymes flanking the insert, Xba1 and Pme1, and the digested product was run on a 1% agarose gel. The DNA band was extracted from the gel and ligated, as described earlier, into the original pcDNA6.2 vector that has been emptied of both the insert and the GFP tag. The ligation product was transformed into DH5ɑ cells, inoculated in LB media and the DNA was purified to be sequenced. Verification of all mutants and constructs were done through sequencing of DNA at ACGT Corporation. All primers used are listed in Table 4.2.

Table 4.2: List of primers used for PCR, site-directed mutagenesis, and sequencing.

Primer Target/Name Primer Sequence (5’ – 3’) CGC TGC GCG CGG TGT GAC TCA CGC TGA SRD5A3, 57G>A, Forward CCG CCG CGG CGG TCA GCG TGA GTC ACA CCG CGC SRD5A3, 57G>A, Reverse GCA GCG CGG CAC CTG CCA ACA GGG AAG GTG GCG COG5, 95T>G, Forward GCG G CCG CCG CCA CCT TCC CTG TTG GCA GGT COG5, 95T>G, Reverse GCC G GGG GAT GTG ATT CCG TTT CAG CAT CAT COG5, 2327dupT, Forward TAT TCA G CTG AAT AAT GAT GCT GAA ACG GAA TCA COG5, 2327dupT, Reverse CAT CCC C TAA TAC GAC TCA CTA TAG GG T7 Forward Primer ATT TAG GTG ACA CTA TAG SP6 Reverse Primer CGC AAA TGG GCG GTA GGC CMV Promoter Forward Primer CTT CCG TGT TTC AGT TAG C TK PolyA Reverse Primer ACA AGG GCT CGA GCC CAT CAA FP1 Forward Primer AAA ATC TAG AGC CAC CAT GCA TCA TCA His-tag for COG5, Xba1, Forward TCA TCA TCA TGG CTG GGT GGG CGG GCG G TTT TGT TTA AAC TTA CCT GAT GAG GAG COG5, Pme1, Reverse GAG CCT GTC AAA ATC TAG AGC CAC CAT GCA TCA TCA His-tag for SRD5A3, Xba1, Forward TCA TCA TCA TGC TCC CTG GGC GGA GGC C TTT TGT TTA AAC AAA CAA AAA TGG TAG SRD5A3, Pme1, Reverse GAA AGC TTT

4.2.2 DNA plasmid preparation and purification

DNA plasmid was transformed into high-efficiency, competent DH5ɑ E. coli cells. 1μl of DNA (~100ng) was mixed in 30μl of competent cells and incubated on ice for 20 minutes, before being heat-shocked in a 42°C water bath for 45 seconds, and placed back on the ice for 2 minutes. 500μl of lysogeny broth (LB) medium is pipetted into the DNA-competent cell mixture and incubated at 37°C while shaking at 225rpm for 30 minutes. 250μl of the LB-competent cell outgrowth media is plated on pre-warmed LB-Agar plates containing the appropriate antibiotic the DNA plasmid is resistant to, as a selective marker. The plate is incubated overnight in a 37°C chamber and placed at 4°C the following morning. Colonies that grew on the plate are picked using a sterile pipette tip and dropped into 5mL (for mini-prep) or 250mL (for maxi-prep) of LB media that includes the selective antibiotic, for example, ampicillin. The LB media is incubated at 37°C while shaking at 225rpm overnight and DNA purification commences the following morning.

Purification of a small yield of DNA plasmid for purposes of sequencing and ensuring accurate and precise mutagenesis was done using a Plasmid Mini Kit (QIAGEN), according to the manufacturer’s outlined procedure. Preparation of a high yield of DNA plasmid for purposes of in vivo and in vitro transfection was done using a Plasmid Maxi Kit (QIAGEN), according to the manufacturer’s outlined procedure. DNA plasmid is solubilized in 100 - 200μl of Tris-EDTA buffer and stored at -20°C. DNA plasmid concentration is measured through NanoDrop 2000c (ThermoFisher Scientific) nucleic acid quantification.

4.2.3 Subretinal injection of DNA followed by in vivo electroporation

For successful transfection of DNA in neonatal mice, the progenitor cells were targeted in pups of age P0-P1. The procedure was adapted from de Melow & Blackshaw (2011)147, with a mixture of 2-5ug/mL of DNA solution and 0.1% Fast Green dye prepared for injection. Neonatal pups were anesthetized on ice for 5 minutes before opening one of the eyelids using a sharp 30- gauge needle. The eye is punctured by inserting a sharp 30-gauge needle midway behind the lens. A syringe with a blunt-end 33-gauge needle is inserted through the puncture until the resistance of the retina is felt, and 0.4μl of the DNA mixture is gently injected subretinally. Tweezer-style electrodes, which were soaked in PBS, are placed against both eyes of the pup with the positive pole against the injected eye. Five pulses of 80 volts that are 50 milliseconds in

42 duration with 950-millisecond intervals are applied using an electroporator. The pup is warmed up before being returned to its parents, and its eyes are harvested at a later timepoint, as per experimental requirements.

4.2.4 Harvesting, fixation, and sectioning of retinas

At the desired age of assessment, the pups are sacrificed using cervical dislocation to collect the retinas for analysis preparation. If not naturally open, the eyelids are opened using a 30-gauge sharp needle, and the eye is extracted using curved forceps and immediately placed in PBS where it is punctured using a 30-gauge needle to allow paraformaldehyde (PFA) to enter the eye and replace the vitreous fluid. The eye is placed in 4% PFA (in PBS) for 1 hour at 4°C. To enhance sectioning, the lens is micro-dissected out of the eye. To protect from freezing damage, the tissue is cryo-protected by being placed in 15% sucrose for 2-3 hours, followed by 30% sucrose for 2-3 hours. The retina is embedded in a mixture of 30% sucrose and O.C.T. freezing media in a 1:1 ratio and frozen at -80°C. The retina is sectioned at a cryostat with a thickness of 12 microns at a chamber temperature of approximately -25°C. Sections are placed on slides and stored in a light-protected slide-box at -80°C.

4.2.5 Immunohistochemical staining of sectioned retinas

Slides with retinal sections are taken out of storage from the -80°C freezer and allowed to dry at room temperature in the dark for several hours. The slides are washed for 5 minutes with PBS. For certain antibodies, as indicated in Table 4.3, antigen-retrieval is required. Antigen-retrieval is performed by immersing slides in citrate buffer (made of 10mM citric acid, pH 6.0 with 0.05% Tween 20) at room temperature for 10 minutes and then placing them in a 95°C water bath in a container of pre-heated citrate buffer for 20 minutes. The slides are allowed to cool at room temperature for 40 minutes and then rinsed 3 times, for 5 minutes each, with PBS + 0.01% Tween 20 solution. If antigen-retrieval is not required, the slides are permeabilized by washing with PBS + 0.1% Triton™ X-100 solution for 10 minutes. After rinsing with PBS, the slides are blocked with 10% FBS for 1 hour at room temperature. Afterward, the primary antibody, diluted in 10% FBS + 0.1% Tween 20 solution, is applied onto the slide within an area enclosed by a liquid barrier PAP pen, and incubated in a humidified chamber overnight at 4°C. The primary antibody solution is washed away with PBS + 0.1% Tween 20 solution 3 times, 5 minutes each. The secondary antibody, diluted in 10% FBS + 0.1% Tween 20 solution, is applied to the

43 sections for 1 hour at room temperature, in a dark humidified chamber. See Table 4.4 for a list of secondary antibodies used. The slides are washed with PBS, and the sections are counter-stained with 0.1ug/mL of DAPI (DNA stain) for 10 minutes. The slides are rinsed with PBS before being mounted with a coverslip using Mowiol® mounting medium and allowed to dry overnight. Slides are stored in the dark at room temperature and are imaged using a fluorescence microscope.

4.3 In vitro assessment of gene mutations 4.3.1 Culturing cells

Both N1E-115 (mouse neuroblastoma) and HEK-293 (human embryonic kidney) cells were cultured in DMEM (Sigma-Aldrich) supplemented with 10% FBS (Gibco) and 01% Penicillin- Streptomycin (Gibco). Cells were passaged twice weekly by aspirating the old media and washing the plate with pre-warmed D-PBS (Sigma-Aldrich) and then applying pre-warmed 0.25% Trypsin with EDTA 4Na (Gibco) for approximately 3 minutes to suspend the adherent cells. Pre-warmed, supplemented, DMEM was applied on top of the Trypsin and manual force by pipetting the media several times to completely suspend the cells before transferring to a conical tube. The tube was spun in a centrifuge at 1,100 rpm for 5 minutes. The pellet was resuspended in fresh, supplemented, DMEM and equally rationed among several tissue culture plates at an optimal seeding ratio for an experiments’ objective. The passage number was labelled, and the plates were incubated in a 37°C, 5% CO2 chamber. N1E-115 cells were used due to being morphologically similar to neuronal cells in the retina, while HEK-293 cells were used for increased transfection efficiency for assessing global changes to ER stress and UPR markers in cell lysates through Western blotting.

4.3.2 Transfection of cells with DNA constructs

Lipofectamine 3000 reagent (ThermoFisher Scientific) was used to maximize transfection efficiency of DNA constructs in N1E-115 and polyethyleneimine (PEI) was used for HEK-293 cells. For 10 cm plates, as used for Western blot lysis preparation, 14μg of DNA was mixed with 28μl of P3000 reagent in 500μl of Opti-MEM (Gibco) and incubated with a solution of 21.7μl of Lipofectamine 3000 reagent in 500μl of Opti-MEM. For 12-well plates, as used for immunocytochemistry, 1μg of DNA was mixed with 2μl of P3000 reagent in 50μl of Opti-MEM

44 and incubated with a solution of 1.5μl of Lipofectamine 3000 reagent in 50μl of Opti-MEM. For HEK-293 transfection with PEI, 9μg of DNA was mixed with 27μl of PEI in 500μl of Opti- MEM. The transfection mixtures for both PEI or Lipofectamine 3000 were left to incubate at room temperature for 15 minutes before being added on top of the supplemented growth media for cultured cells. In PEI-treated cells, the media was replaced with fresh supplemented media 6 – 16 hours after transfection. For GFP or RFP (red fluorescent protein) tagged constructs, transfection success was assessed under a fluorescence microscope. Transfected cells were fixed or lysed 24 – 48 hours after transfection, as per experimental objectives.

4.3.3 Cell lysis preparation and protein quantification

For the purposes of Western blotting, transfected HEK-293 cells in 10-cm dishes were lysed using 1X RIPA lysis buffer (Cell Signaling Technology) with 1:100 dilution of protease inhibitor (PI) cocktail (Sigma-Aldrich). Growth media was aspirated from cell culture plates, and the cells were washed with ice-cold D-PBS twice. D-PBS was aspirated out, and 150μl of 1xRIPA lysis buffer with PI was added on top of the cells. The cells were scraped out and pipetted into a pre- chilled 1.5mL Eppendorf tube and placed on ice for 30 minutes while being vortexed every 5 minutes. The cell lyses were centrifuged for 30 minutes at 4°C at 13,200 rpm in a benchtop microcentrifuge. The pellet was discarded, and the supernatant containing the lyses was collected, aliquoted, and stored at -20°C. Total protein concentration in the cell lyses was measured using the Pierce BCA Protein Assay Kit (ThermoFisher Scientific). Standards were prepared through a serial dilution of bovine serum albumin (BSA) in 1xRIPA lysis buffer with PI. 96-well plate was used to add 200μl of BCA working reagent onto a 10-time dilution of each cell lysis, which was then incubated for 30 minutes in a 37°C chamber. The absorbance of each cell lysis sample was measured at 560nm on a plate reader. Protein concentration was measured using the standard curve plotted using the absorbance of each BSA standard against its concentration in μg/mL.

4.3.4 Western blotting

Specific protein concentration levels in cell lyses are analyzed through sodium dodecyl sulfate – polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto a nitrocellulose membrane for antibody staining and imaging. Cell lyses loading samples are prepared and diluted using 1xRIPA with PI to equalize all samples’ concentrations, according to results of the Pierce BCA

Protein Assay Kit. Laemmli buffer with dithiothreitol (DTT) is added to each sample from a 6X stock solution. The cell lysis sample is then heated on a 95°C heat block for 5 minutes and then placed on ice for 2 minutes before loading. The SDS-PAGE gel is prepared according to the appropriate pore size for clear visualization of protein (usually between 10 – 15%) and topped with a 3% stacking gel. After the samples are loaded, the gel is run at 180 volts for 45 – 50 minutes in Tris-Glycine SDS running buffer.

The separated protein on the SDS gel were transferred onto a nitrocellulose membrane that was activated in 20% ethanol. The gel and membrane are pressed against each other and tightly sandwiched using filter paper and sponges. The gel-membrane sandwich is placed in a transfer chamber that is filled with Tris-Glycine Western transfer buffer and is enclosed in an ice bucket to keep it cool. The transfer is run either for one hour at 80 volts (for smaller proteins), or overnight at 11 volts (for larger proteins). The membrane is air-dried for 30 minutes in a laminar flow hood and then re-hydrated for 5 minutes with PBS. The membrane is blocked at room temperature with either 5% skim milk (in PBS) for 1 hour or with 2.5% BSA (in TBS) for 30 minutes, depending on primary antibody used. The primary antibody is diluted in blocking buffer with 0.1% Tween 20 and incubated with the membrane overnight at 4°C. See Table 4.3 for a list of primary antibodies used and respective conditions applied. The membrane is washed with PBS + 0.1% Tween 20 (PBS-T) 3 times, 10 minutes each, and then incubated with the secondary antibody diluted in blocking buffer with 0.1% Tween 20. See Table 4.4 for a list of secondary antibodies used. The membrane is washed with PBS-T 3 times, 10 minutes each, and can be stored in PBS at 4°C. The membrane is imaged using Li-Cor Odyssey fluorescence imaging system. Quantifications are made using Li-Cor’s Imaging Studio™ software and normalized to quantifications of glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as a housekeeping protein.

4.3.5 Fixation and immunocytochemical staining of transfected cells

To allow adhesion of cells, round glass coverslips are coated with 0.01% poly-L-lysine (PLL) solution (Sigma-Aldrich) and incubated overnight at 4°C or for 1 hour in a 37°C chamber. The coated coverslips are rinsed 3 times with sterile ddH2O and then allowed to dry for at least 1 hour. Cells are cultured on each coverslip in supplemented DMEM growth media and transfected using Lipofectamine 3000 as described above. 24 – 48 hours after transfection, media is

46 aspirated, and the cells are immediately fixed using 4% PFA (in PBS) for 10 minutes at room temperature. Cells are washed 3 times, 5 minutes each with PBS and then permeabilized in PBS + 0.1% Triton™ X-100 at room temperature for 10 minutes. Cells are washed 3 times, 5 minutes each and blocked for 30 minutes at room temperature using 10% FBS (in PBS-T). Primary antibody (see Table 4.3 for details), diluted in blocking buffer, is applied on the cells and incubated overnight at 4°C. The primary antibody is washed away with PBS-T 3 times, 5 minutes each. The secondary antibody (see Table 4.4 for details), diluted in blocking buffer, is applied on the cells and incubated for 1 hour at room temperature. The cells are washed with PBS-T and counter-stained with DAPI. The cells are rinsed with PBS and then mounted with Mowiol® mounting medium on a glass slide. Slides are allowed to dry in the dark, at room temperature, overnight before imaging using a fluorescence microscope.

4.3.6 Imaging, analyses, and quantification

Image acquisition of the retina and cells were obtained using a Carl Zeiss LSM700 laser- scanning confocal microscope and Olympus BX61 epifluorescence microscope. Images taken using the confocal microscope were exported from Zen Black to Zen Blue and processed using the in-house software. Images for DNA damage and colocalization analysis were taken using the epifluorescence microscope at 20x magnification and analyzed and processed using the in-house CellSens Dimension software. A region of interest (ROI) was drawn around each transfected cell and a scatterplot was generated between the intensity of the two fluorophore wavelengths to be assessed for colocalization. The background levels were subtracted from the scatterplot area of measurement and a colocalization coefficient was generated. The colocalization coefficient was transformed to a Z factor for averaging and the averaged total was transformed back to the R2 colocalization coefficient.

Levels of DNA damage in cells were quantified by measuring the fluorescence intensity within a drawn ROI around each cell for γH2A.x antibody staining of double-stranded DNA breaks. Exposure time and laser intensity were kept constant among all measured samples, and background fluorescence was subtracted from every image taken. Colocalization analysis was performed within ROIs drawn around every transfected cell, obtaining correlations between GFP intensity and RFP intensity, representing the transfected construct and the Golgi marker, respectively.

Western blots were imaged using the Li-cor Odyssey fluorescence scanner. The in-house software, ImageStudio, was used for Western blot analysis and processing, including measuring protein band intensity and subtracting background intensity from quantification. The protein band intensity was normalized by measuring the intensity of GAPDH bands. The analysis was performed using Microsoft Excel and GraphPad Prism. Statistical significance for comparing multiple groups was determined by applying one-way ANOVA analysis followed by Tukey post- hoc test. Comparison between two groups was performed by applying Student’s t-test.

Table 4.3: List of primary antibodies used and their respective applications.

Primary Host Dilution Application Supplier Product Antibody Species Code 6xHis-Tag Mouse, 1:200 (IF) WB, ICC, Invitrogen MA1-21315 monoclonal 1:4000 (WB) IHC ATF6 Rabbit, 1:1000 WB Novus NBP1-75478 polyclonal Biologicals BIP Rabbit, 1:1000 WB Cell Signaling C50B12 monoclonal Technology Calnexin Rabbit, 1:1000 WB Cell Signaling C5C9 monoclonal Technology COG5 Rabbit, 1:600 WB, ICC, Sigma- SAB4200440 polyclonal IHC, AR Aldrich GAPDH Mouse, 1:4000 WB Invitrogen ZG003 monoclonal GFP Mouse, 1:500 (IF) WB, ICC, Cell Signaling 2955S monoclonal 1:1000 (WB) IHC Technology GFP Mouse, 1:500 (IF) WB, IHC, Invitrogen GF28R monoclonal 1:1000 (WB) ICC Giantin Rabbit, 1:4000 ICC Biolegend 19243 polyclonal GM130 Rabbit, 1:1000 ICC Cell Signaling 12480S polyclonal Technology IRE1α Rabbit, 1:1000 WB Cell Signaling 3294S polyclonal Technology PDI Rabbit, 1:1000 WB Cell Signaling 2446S polyclonal Technology PERK Rabbit, 1:1000 WB Cell Signaling D11A8 monoclonal Technology Phospho- Mouse, 1:1500 ICC Millipore- 05-636 Histone H2A.X monoclonal Sigma SRD5A3 Rabbit, 1:200 WB, ICC, Abcam Ab104063 polyclonal IHC

WB: Western blotting; ICC: Immunocytochemistry; IHC: Immunohistochemistry; IF: Immunofluorescence (includes ICC and IHC); AR: Antigen-retrieval required

Table 4.4: List of secondary antibodies used and their respective applications.

Secondary Antibody Host Species Dilution Application Supplier

Alexa Fluor 488 Rabbit Donkey 1:500 ICC, IHC Life Technologies

Alexa Fluor 555 Rabbit Donkey 1:500 ICC, IHC Life Technologies

Alexa Fluor 647 Rabbit Donkey 1:500 ICC, IHC Life Technologies

Alexa Fluor 488 GFP Rabbit 1:500 ICC, IHC Life Technologies

Alexa Fluor 568 Mouse Goat 1:500 ICC, IHC Life Technologies

Alexa Fluor 555 Rabbit Donkey 1:500 ICC, IHC Life Technologies

Alexa Fluor 647 Mouse Fab2 1:500 ICC, IHC Life Technologies anti-Rabbit IRDye 680 Donkey 1:20,000 WB Li-cor anti-Rabbit IRDye 800 Donkey 1:20,000 WB Li-cor anti-Mouse IRDye 800 Donkey 1:20,000 WB Li-cor anti-Mouse IRDye 680 Donkey 1:20,000 WB Li-cor

WB: Western blotting; ICC: Immunocytochemistry; IHC: Immunohistochemistry

Results 5.1 SRD5A3: Expression and localization

5.1.1 Constructs

To conduct in vivo and in vitro experiments that investigate the expression and localization of the mutated SRD5A3 protein, a pcDNA 6.2 plasmid was obtained that contains the human-SRD5A3 gene tagged at the N-terminus with an Emerald Green Fluorescent Protein (EmGFP), for clear localization analysis (Figure 5.1A, left). Through site-directed mutagenesis, as described in section 4.2.1, the constructs were mutated to include the identified human mutations, represented in Figure 5.1B. For alternative applications and to rule out any secondary effects that may have been caused by the large size of the EmGFP tag, the constructs were cloned to replace the EmGFP tag with a smaller 6x-His tag, as previously described (Figure 5.1A, right).

Figure 5.1: Constructed plasmids for wildtype and mutant SRD5A3 (A) Simplified vector map of the pcDNA 6.2 plasmids harbouring the human-SRD5A3 gene in its wild-type form or its mutated form (p.W19X). One of the plasmids (left) includes an N-terminus EmGFP tag for SRD5A3, and the other (right) has an N-terminus 6x-Histidine tag instead. (B) Alignment of the SRD5A3 gene segment where the identified mutation arises with the wildtype SRD5A3 gene sequence. The p.W19X nonsense mutation results in guanine being replaced with an adenine, resulting in an immediate premature stop codon.

5.1.2 ER stress and UPR expression analysis

The concentration levels of several ER stress components and UPR markers were measured in HEK-293 cells after transfection with either the 6xHis-tagged plasmid containing wild-type (WT) SRD5A3 or one containing the mutant SRD5A3 (p.W19X). A plasmid with a blank insert was transfected into cells as a negative control, and another group of cells were treated with Tunicamycin (TM), which induces ER stress by blocking N-glycosylation, as a positive control. The measured ER stress markers include calnexin, IRE1, ATF6, PERK, PDI, and BiP. GAPDH, a housekeeping protein, was also measured for normalization. A sample of representative western blots is presented below in Figure 5.2. The concentration levels were quantified from n ≥ 3 experiments and are represented in the bar graphs of Figure 5.3.

The quantification of most of the ER stress markers showed a wide range of variability and a standard error margin that did not result in any statistically significant difference between any of the WT-, W19X-, control-transfected groups. Statistical significance was only found in comparison with the TM-treated positive control groups in protein expression analysis of calnexin, BiP, PDI and IRE1α. For calnexin and BiP, protein expression levels in the TM group were significantly higher than all three remaining groups. PDI protein levels of expression in the TM-treated group were only significantly higher compared to the control- and WT-transfected groups. IRE1α protein expression was only significantly higher in the TM-treated group compared with the WT-transfected group.

Figure 5.2: Representative western blot images comparing the protein expression levels of six ER stress or UPR markers in HEK-293 cells transfected with wildtype SRD5A3 plasmid (WT), mutant SRD5A3 plasmid (W19X), a plasmid with a blank insert (control), or cells treated with tunicamycin (TM). The GAPDH staining is displayed below each respective blot and was used for normalization of band intensity quantification.

Figure 5.3: Expression of mutant SRD5A3 did not induce UPR or ER stress. Quantification of the western blots (as represented in figure 5.2) measuring the protein abundance levels of six ER stress or UPR markers in cells transfected with wildtype SRD5A3 plasmid (WT), mutant SRD5A3 plasmid (W19X), a plasmid with a blank insert (control), or treated with tunicamycin (TM). The protein abundance levels were quantified from n ≥ 3 independent experiments, and band intensities were normalized by measuring GAPDH band intensity. One-way ANOVA analysis followed by post-hoc Tukey test was used to assess levels of significance between compared groups. * = p < 0.05, ** = p < 0.01, *** = p < 0.001. Error bars = S.E.M.

5.1.3 In Vitro localization

SRD5A3 is an ER-resident protein and thus should have a perinuclear distribution and localization within the cell. Mouse neuroblastoma N1E-115 cells were transfected with the GFP- tagged wild-type (WT) or mutant (W19X) SRD5A3 plasmids and the protein’s distribution was assessed under an epifluorescence microscope after nuclear DAPI staining. Visualization of the transfected cells displayed an apparent and stark difference in localization and distribution of the WT and W19X variant forms of SRD5A3 protein. As represented in Figure 5.4, wild-type SRD5A3 was more tightly packed and localized in a perinuclear fashion within the cell. The mutant p.W19X variant of SRD5A3 does not seem to be tightly localization but is instead expressed across the cytosol. This mislocalization of the mutant SRD5A3 protein is likely due to the early stop mutation that results in its truncation.

Figure 5.4: Mutant SRD5A3 is mis-localized within the cell. Representative images of the localization and distribution of the transfected GFP-tagged wild-type SRD5A3 (WT) protein or mutant SRD5A3 (p.W19X) protein in N1E-115 cells. WT SRD5A3 protein shows tightly-packed distribution and perinuclear localization, while p.W19X mutant SRD5A3 protein shows a cytosolic distribution and complete mislocalization within the cell. DAPI is used as a nuclear stain. Scale bar = 50μm.

5.2 COG5: Expression, localization, & function 5.2.1 Constructs

To conduct in vivo and in vitro experiments that investigate the role of COG5 mutations, a pcDNA 6.2 plasmid was obtained that contains the human-COG5 gene tagged at the N-terminus with EmGFP, for clear localization analysis (Figure 5.5A, left). Through site-directed mutagenesis, as described in section 4.2.1, the construct was mutated to include one of the identified human mutations, represented in Figure 5.5B. For alternative applications and to rule out any effects caused by the EmGFP tag, the constructs were cloned to replace the EmGFP tag with a 6x-His tag, as previously described (Figure 5.5A, right).

Figure 5.5: Constructed plasmids for wildtype and mutant COG5. (A) Simplified vector map of the pcDNA 6.2 plasmids harbouring the human-COG5 gene in its wild-type form or with any one of its two mutations (p.M32R or p.S777Q_fs*14). One of the plasmids (left) includes an N- terminus EmGFP tag for COG5, and the other (right) has an N-terminus 6x-Histidine tag instead. (B) Alignment of the COG5 gene segments where the identified mutations arise with the wildtype COG5 gene sequence. Top: the p.M32R missense mutation results in thymine being replaced with guanine. Bottom: the p.S777Q_fs*14 nonsense mutation results in a duplication of thymine, which results in a premature stop signal 14 codons downstream of the mutation.

5.2.2 ER stress and UPR expression analysis

The concentration levels of several ER stress components and UPR markers were measured in HEK-293 cells after transfection with one of the 6xHis-tagged plasmids containing wild-type (WT) COG5, COG5 with missense (M32R) mutation, COG5 with stop-gain duplication (S777Q) mutation, or transfected with both COG5 mutant plasmids. A plasmid with a blank insert was transfected into cells as a negative control, and another group of cells were treated with Tunicamycin (TM), which induces ER stress by blocking N-glycosylation, as a positive control. The measured ER stress markers include calnexin, IRE1, ATF6, PERK, PDI, and BiP. GAPDH, a housekeeping protein, was also measured for normalization. A sample of representative western blots is presented below in Figure 5.6. The protein abundance levels were quantified from n ≥ 6 independent experiments and are represented in the bar graphs of Figure 5.7.

A significant difference in protein levels of two of the three UPR branches was found between the WT and mutant COG5 transfection groups. IRE1α was the only branch that was not influenced and had a consistent and similar level of protein abundance between all four COG5 treatment groups (WT, M32R, S77Q, and M32R+S77Q) and the control.

The second UPR arm, ATF6, presented a wide range of variability across experiments for the mutant COG5 groups, leading to a large S.E.M., and no statistically significant difference of protein abundance compared with the co-transfection group. However, the missense-mutant- transfected group (M32R) had a significantly higher abundance of full-length ATF6 compared with the WT.

The protein abundance of the third UPR arm, PERK, was significantly elevated in the cells transfected with both mutants (M32R + S777Q), compared with the WT- and M32R-transfection groups. Additionally, only the double-mutant-transfected group demonstrated a significant difference compared with the TM-treated group. This highlights an important effect of having both COG5 mutants on the abundance of PERK.

The remaining ER stress markers are BiP, PDI, and calnexin. Both BiP and PDI were expressed in similar levels across the four transfection groups but were significantly upregulated under the treatment of TM. Calnexin was expressed at a similar level for all four COG5-transfected groups (WT, M32R, S77Q, and M32R + S777Q) but only the single-mutant transfection groups (M32R

60 and S777Q) had significantly lower calnexin abundance compared with the TM positive control group.

Figure 5.6: Representative western blot images comparing the protein expression levels of six ER stress or UPR markers between cells transfected with wildtype COG5 plasmid (WT), COG5 plasmid containing a missense mutation (M32R), COG5 plasmid containing a stop-gain duplication mutation (S777Q), both mutant COG5 plasmids co-transfected (M32R + S777Q), a plasmid with a blank insert (control), or cells treated with tunicamycin (TM). The GAPDH staining is displayed below each respective blot and was used for normalization of band intensity quantification.

Figure 5.7: Co-transfection of COG5 mutants activated the PERK branch of the UPR. Quantification of the western blots (as represented in figure 5.6) measuring the protein abundance of six ER stress or UPR markers in cells transfected with wildtype COG5 plasmid (WT), single-mutant COG5 plasmids (M32R; S777Q), co-transfected with both COG5 mutants (M32R + S777Q), or treated with tunicamycin (TM), relative to the protein abundance in a plasmid with a blank insert (control). The concentration levels were quantified from n ≥ 6 independent experiments, and band intensities were normalized by measuring GAPDH band intensity. One-way ANOVA analysis, followed by Tukey post-hoc test was used to assess levels of significance between compared groups, * = p < 0.05, *** = p < 0.001. Error bars = S.E.M.

5.2.3 In Vitro localization and Golgi dispersal

The GFP-tagged constructs were used to transfect mouse neuroblastoma (N1E-115) cells in order to visualize the localization of the wild-type and mutant COG5 proteins within the cell and to assess its co-localization with the Golgi apparatus. GM130, a golgin, was used as a Golgi marker and DAPI was used for nuclear staining. Figure 5.8 displays representative photos of the GFP- tagged wild-type (WT), M32R-mutant, and S777Q-mutant COG5 proteins and their localization within the cell. Additionally, a group of cells were transfected with both M32R- and S777Q- mutant COG5 variants to assess any influence on Golgi co-localization. The S777Q mutation in COG5 is a stop-gain frameshift mutation, meaning that insertion mutation results in a stop codon to occur prematurely downstream. This mutation does not likely lead to nonsense-mediated decay as the GFP-tagged S777Q-mutant COG5 protein is expressed in transfected cells and has a perinuclear localization (Figure 5.8), indicating that it is undergoing a normal translation process.

Figure 5.9 shows the quantified co-localization analysis of each of the four groups in a bar graph. In all groups (WT, M32R, S77Q, and M32R + S77Q), COG5 was co-localized with GM130 at a similar level, as no statistical significance was found between any of the groups. Interestingly, the quantifications showed that COG5, in all groups, was more mislocalized with GM130, noting that the colocalization coefficient was in the negative. This is likely due to the close proximity of COG5 to the Golgi, but its existence in the cytosol, rather than the luminal area.

As can be noted in Figure 5.8, however, the appearance of the Golgi, as stained by GM130, appears to be influenced differently by each of the four transfection groups. The apparent dispersal of the Golgi also corresponds with a noticeable dispersal of the GFP-tagged COG5 protein as well. To assess the level of Golgi fragmentation or dispersal, its area was measured relative to the area of non-transfected Golgi and compared across the four groups. These quantification results are presented in Figure 5.10 and indicate that level of Golgi dispersal was most severe in the group co-transfected with M32R- and S777Q-mutant variants of COG5. In fact, the level of Golgi dispersal in the co-transfected group was significantly higher than all other transfection groups. Both single-mutant transfection groups (M32R; S777Q) also showed a significantly higher amount of Golgi dispersal compared with the wild-type transfected group.

Figure 5.8: Representative images of N1E-115 cells transfected with GFP-tagged wild-type (WT) COG5, mutant (M32R; S777Q) COG5, or co-transfected with both mutant COG5 plasmids (M32R + S777Q). The cells were stained for the Golgi using GM130 antibody, a cis-Golgi marker. DAPI was used for nuclear staining. Arrow points towards an example of a normal Golgi structure in un-transfected cells. Scale bar is 20μm.

Figure 5.9: COG5 mutants did not disrupt colocalization of COG5 with the Golgi. Quantification of the colocalization between the GFP that is tagged to the COG5 constructs and GM130, a cis-Golgi marker, in cells transfected with wild-type (WT) COG5, mutant (M32R; S777Q) COG5, or co-transfected with both mutant COG5 plasmids (M32R + S777Q). For each group, >350 cells were quantified across n = 6 independent experiments. Statistical analysis was performed using one-way ANOVA analysis. Error bars = S.E.M.

Figure 5.10: COG5 mutants led to increased Golgi fragmentation. Quantification of the relative Golgi area, indicating the level of Golgi dispersal, compared to control across four groups of transfected cells. Cells were transfected with wild-type type (WT) COG5, mutant (M32R; S777Q) COG5, or co-transfected with both mutant COG5 plasmids (M32R + S777Q). For each group, >150 cells were quantified across n = 3 independent experiments. Results were normalized to the level of Golgi area in the untransfected control cells. Statistical analysis was performed using one-way ANOVA analysis followed by Tukey post-hoc test, * = p<0.05, ** = p<0.01 *** = p<0.001. Error bars = S.E.M.

5.2.4 DNA damage induction

Due to the increase in levels of Golgi dispersal observed (Figures 5.8 and 5.10) in cells transfected with both COG5 mutants (M32R + S777Q), and to a lesser extent in cells transfected with the single mutants (M32R; S777Q), the level of DNA damage in each of the four transfection groups was measured. This is because previous studies have demonstrated a clear link between Golgi fragmentation or dispersal and DNA damage47.

Figure 5.11 displays representative images of the different COG5 plasmids transfection groups (WT, M32R, S777Q, and M32R + S777Q) in mouse neuroblastoma N1E-115 cells, and immunostained for double-stranded DNA breaks using anti- gamma-phospho-histone H2A.X antibody. DAPI was used for nuclear staining. Assessment of γ-H2A.X-indicated DNA damage has been performed by measuring the fluorescence intensity within a region of interest drawn around the cell and averaged through a Z-stack, which has proven to be of comparable reproducibility to other methods involving foci counting and flow cytometry148. The quantification of γ-H2A.X fluorescence intensity in each of the four transfection groups is presented in Figure 5.12. The fluorescence intensity of γ-H2A.X in cells co-transfected with both M32R and S777Q mutant COG5 plasmids was found to be significantly higher than all other transfection groups, including the control of GFP-transfected cells.

The double-mutant co-transfected group (M32R + S777Q) of cells was the only group that had a significantly higher level of measured DNA damage than all other groups including the control, and previously it was demonstrated that only the M32R + S777Q group had a significantly higher level of PERK protein expression (Figure 5.7) than the control and the WT and M32R groups. As a result, a PERK inhibitor, GSK2606414 (PERKi), was used to treat the transfected cells to assess whether there is a link between PERK upregulation and DNA damage. Indeed, it was found that PERKi treatment significantly reduced the level of γ-H2A.X fluorescence intensity only in the M32R + S777Q group and reduced it to a level that was similar to the fluorescence intensity of the control and the S777Q-mutant groups (Figure 5.14).

The significant decrease in fluorescence intensity of the WT and M32R groups compared with the double-mutant group remained significant after PERKi treatment. PERKi treatment also resulted in levels of DNA damage for the WT and M32R groups that were significantly less than the S777Q mutant group and resulted in the WT group have significantly less DNA damage than the GFP

68 control as well (Figure 5.14). This may speak to an unintended effect of GFP transfection on causing DNA damage. It is of note that PERKi treatment decreased the levels of DNA damage of the M32R + S777Q group to reach similar levels as DNA damage in the S777Q-only group but was still significantly higher than DNA damage found in the WT and M32R-only groups, after and before PERKi treatment. This may hint at a predominant role that the S777Q mutant variant of COG5 plays in causing DNA damage.

Representative images of the N1E-115 cells transfected with WT, M32R, S777Q, or M32R + S777Q COG5 plasmids and immunostained for γ-H2A.X after PERKi treatment are presented in Figure 5.13. DAPI was used as a nuclear stain. Quantifications of fluorescence intensity in the transfection groups after PERKi treatment and the comparison of these levels against measurements of untreated cells are presented in Figure 5.14.

To confirm the association of PERK upregulation leading to an increase in DNA damage, a group of N1E-115 cells were transfected with a PERK plasmid to overexpress the protein, and then assessed for γ-H2A.X fluorescence intensity (Figure 5.15). This was compared to the γ- H2A.X fluorescence intensity in a group of cells that were transfected with a control GFP. Additionally, the specificity of the PERK inhibitor was confirmed by treating both transfection groups with PERKi (Figure 5.15). The quantification of the fluorescence intensity measurements are presented in Figure 5.16, which display a significant increase in DNA damage in the PERK- transfected groups compared with the GFP-transfected group. Additionally, the PERK inhibitor treatment reduced the levels of DNA damage in the PERK-transfected group to levels that were comparable to that of the control (Figure 5.16).

Figure 5.11: Representative images of cells transfected with GFP-tagged WT, M32R, S777Q, or co-transfected with M32R + S777Q variants of COG5 plasmids in N1E-115 cells. The concentration of the DNA plasmid transfected in cells is represented in brackets. The cells were immuno-stained against γ-H2A.X and DAPI was used for nuclear staining. Scale bar is 20μm.

Figure 5.12: Co-transfection of COG5 mutants induced DNA damage. Quantification of γ- H2A.X fluorescence intensity, in arbitrary units (AU), measured in cells transfected with WT, M32R, S77Q, or both M32R + S777Q variants of GFP-tagged COG5 plasmids. A GFP-only plasmid was also transfected in cells as a control and was measured for γ-H2A.X fluorescence intensity as well. For each group, n>300 cells were measured and pooled from n ≥ 4 independent experiments. The concentration of the DNA plasmid transfected in cells is represented for each group in brackets. Statistical significance was assessed using one-way ANOVA analysis followed by Tukey post-hoc test. * = p < 0.05, *** = p < 0.001. Error bars = S.E.M.

Figure 5.13: Representative images of cells that were treated with PERKi and transfected with GFP-tagged WT, M32R, S777Q, or co-transfected with M32R + S777Q variants of COG5 plasmids in N1E-115 cells. The concentration of the DNA plasmid transfected in each group is represented between brackets. The cells were immuno-stained against γ-H2A.X and DAPI was used for nuclear staining. Scale bar is 20μm.

A B

Figure 5.14: Treatment of transfection groups with PERK inhibitor reduced DNA damage in double-mutant group only. Quantification of DNA damage levels after PERKi treatment. (A) Quantification of γ-H2A.X fluorescence intensity, in arbitrary units (AU), measured in cells treated with PERKi and transfected with WT, M32R, S777Q, or both M32R + S777Q variants of GFP-tagged COG5 plasmids. A GFP-only plasmid was also transfected in cells as a control and was measured for γ-H2A.X fluorescence intensity. Statistical significance was assessed using one-way ANOVA analysis followed by Tukey post-hoc test. ** = p < 0.01, *** = p < 0.001. Error bars = S.E.M. (B) Comparison of quantified γ-H2A.X fluorescence intensity between PERKi treated (coloured) and untreated (uncoloured) groups. For each group, n > 300 cells were measured and pooled from n ≥ 4 independent experiments. For the PERKi-treated control group, n = 192 cells were measured and pooled from three experiments. The concentration of DNA plasmid transfected in cells are represented in brackets. Statistical significance was assessed using Student’s t-test. * = p < 0.05. Error bars = S.E.M.

Figure 5.15: Representative images of N1E-115 cells that were transfected with a PERK- expressing plasmid or a control GFP plasmid and untreated (top), and cells that were transfected with PERK or a control GFP as well as treated with PERKi (bottom). The cells were immunostained against γ-H2A.X and DAPI was used for nuclear staining. Scale bar is 20μm.

Figure 5.16: Overexpression of PERK induces DNA damage. Quantification of γ-H2A.X fluorescence intensity, in arbitrary units (AU), measured in cells transfected with PERK or GFP and treated or untreated with PERK inhibitor (PERKi). For each group, n>150 cells were measured and pooled from n ≥ 3 independent experiments. Statistical significance was assessed using Student t-test. * = p < 0.05, ns = not significant. Error bars = S.E.M.

Discussion

This project started with the diagnosis of five individuals with LCA and their categorization of having either a homozygous mutation in SRD5A3 or compound heterozygous mutations in COG5. The association of glycosylation-related genes with LCA is uncommon, if not novel. Due to the critical role that SRD5A3 plays in forming the precursor for N-glycosylation, and the role that COG5 plays in in retrograde Golgi trafficking, an investigation into the gene mutations with relation to ER stress was commenced. A link between SRD5A3 and UPR activation was not found, but the UPR was found to be upregulated in cells transfected with mutant COG5 constructs. This link led to several other experiments and measurements to be conducted for COG5 variants and led to the identification of a link between the expression of the compound heterozygous variants and increase in DNA damage. In the following discussion, the findings and interpretation with regards to SRD5A3 will be briefly addressed first and will be succeeded by a lengthier discussion into the results pertaining to COG5.

6.1 SRD5A3

It was previously reported4,30, and it was evident through the in vitro visualization of WT and W19X SRD5A3 localization, that the mutant variant was non-functional, due to its early stop codon and truncation (Figure 5.4). Western blot analysis of ER stress and UPR markers in lysates of cells transfected with WT or W19X-mutant SRD5A3 plasmid did not demonstrate an explicit activation of any of the three UPR arms (Figure 5.3). In fact, there was considerable variability in the ER stress/UPR markers across experiments (even though cell age at lysis was the same across groups) that even an indicated average increase in abundance was not statistically significant. The variability is likely normal as the cell undergoes natural levels of ER stress or UPR activation through its quality control mechanism, but with more time available, perhaps a higher number of experiments performed may have yielded results of significant difference.

Tunicamycin treatment did not significantly increase the level of ATF6 or PERK protein abundance, likely because tunicamycin’s mode of ER stress activation is through the blocking of N-glycosylation. However, there are various other forms of post-translation modifications and protein folding disruptions that could occur that may activate other arms of the UPR149,150. Tunicamycin-treatment, however, did elevate the protein level of calnexin and BiP compared to

76 all other groups, indicating that both Calnexin and BiP are likely not upregulated as a result of mutant-SRD5A3 expression. IRE1α protein level was only significantly upregulated in the tunicamycin group compared with the wild-type SRD5A3-transfected group. However, this does not indicate activation of the IRE1α arm in the mutant-transfected group because the level of IRE1α protein in the mutant-transfection group was not significantly different from the levels expressed in the negative control group (Figure 5.3).

The only ER stress marker that may have been activated due to mutant-SRD5A3 transfection is PDI. PDI’s levels of protein were significantly lower in the wildtype and control-transfected groups compared with the tunicamycin-treated group, however, there was no significant difference between protein levels in the mutant-transfected group and the tunicamycin-treated group. This does not definitively indicate an activation of PDI due to mutant-SRD5A3 expression because there was also no significant difference found between the wildtype-, mutant-, and control-transfected groups (Figure 5.3). The lack of statistical significance between the mutant and tunicamycin groups may be linked to the large variability in abundance across experiments or might be due to the PDI’s response to the truncated SRD5A3 protein and its attempt at helping it fold properly. However, due to SRD5A3’s short length, it is unlikely to aggregate at the ER, and it was in fact found to be free-floating in the cytosol (Figure 5.4), decreasing the likelihood of the UPR to be activated.

It should be noted that this study regarding SRD5A3 had the limitation of endogenous SRD5A3 being already expressed in the cultured cells. Thus, the exact effect of a homozygous nonsense mutation in SRD5A3 cannot be assessed without creating a transgenic cell line expressing the mutant variant only or conducting experiments on patient-derived cells. Additionally, subretinal in vivo electroporation of the WT and mutant SRD5A3 constructs in mice only resulted in the successful expression of the mutant non-functional variant, likely due to not having any disruptive effect on the cell. Transfection of GFP-tagged wild-type SRD5A3 was not observed, possibly due to the degradation of cells overexpressing a functional form of the protein (results not shown). This is outside of the scope of this project and so was not followed further but would form the basis of an interesting independent study looking at the effect of having an abundance of SRD5A3 protein and an increase in dolichol synthesis.

While the phenotypes presented by the two patients from Case A who also had LCA were not found to be associated with ER stress or UPR activation in our experimental model, they join at least seven other identified individuals who share the same homozygous SRD5A3 p.W19X mutation and have early-onset retinal dystrophy6. The identification of the two unrelated individuals in this project reaffirms the role that this autosomal recessive mutation in SRD5A3 has in causing retinal degeneration. Future investigations should further look into how this disruption in a critical glycosylation component does not result in embryonic lethality, but negatively affects the retina and causes other phenotypes such as developmental delay and motor deficits. The truncating mutation leading to a non-functional SRD5A3 protein would also make diseases caused by the W19X mutation a prime candidate for gene therapy to rescue the phenotypes caused by loss of SRD5A3 function.

6.2 COG5

The objectives of this project were to understand how the compound heterozygous mutations in COG5 result in LCA and the various other symptoms expressed by the three patients of Case B (Figure 2.2, Table 2.2). By evaluating six ER stress and UPR markers, it was found that the COG5 mutants and the co-transfection of both mutants do have an influence on some ER stress activation and UPR upregulation (See Figure 5.7). Similar to what was found in the SRD5A3 investigation, however, many ER stress markers were only found to be significantly elevated in the tunicamycin-treatment groups. PDI and BiP were both found to be significantly reduced in all transfection groups in comparison to the TM-treated group. Calnexin was found to be significantly upregulated in the TM-treated group only compared with the single-mutant transfection groups. However, there was no significant difference amongst the four COG5 variant transfection groups, indicating a similar level of protein abundance and perhaps a generalized effect of COG5 overexpression on calnexin regulation. Tunicamycin, however, did not demonstrate a significant increase for any of the three UPR branches (PERK, ATF6, and IRE1α). This is likely due to the fact that tunicamycin induces ER stress by stopping N- glycosylation, which is only one type of several modifications that an occur on nascent proteins, such as O-glycosylation, phosphorylation, acetylation, among many others149,150. Many proteins do not undergo N-glycosylation and will likely not be affected by tunicamycin treatment, and as a result, ER stress markers and the UPR sensors will not be globally activated. Other positive

78 control treatments that can induce ER stress through other mechanisms include thapsigargin, dithiothreitol (DTT), and brefeldin A151.

Despite the proposed integrated UPR model of the three UPR arms being inter-dependent, the results obtained paint a more independent view of the UPR arms, as each of the three arms had a different activation pattern. Of the three UPR arms, IRE1α was the only one that did not show a significant level of activation for any of the WT or mutant COG5 transfection groups (Figure 5.7). Since IRE1α is considered to be the regulator between cell survival and apoptosis152, its steady-state levels across transfection groups potentially signify that the COG5 mutants do not cause retinal degeneration by directly signalling cell apoptosis – or at least, they do not signal cell apoptosis through IRE1α activation.

The second UPR arm was assessed by measuring full-length ATF6, which is the inactivated form of the protein. To be activated, ATF6 is transported to proteasomes at the Golgi where it is cleaved and then is transported into the nucleus for transcriptional regulation of ER-resident protein folding chaperones111,119. The protein level of full-length ATF6 in cells transfected with WT COG5 was found to be significantly less than its abundance level in the M32R-mutant COG5 transfection groups (Figure 5.7), which implies that either the wild-type COG5 protein reduces ATF6 production, which seems unlikely, or that it results in the over-activation of the protein, resulting in its cleavage. The decrease in full-length ATF6 abundance in the WT group was not significant compared with the S777Q-mutant or double-mutant group, but the S777Q- mutant and double-mutant group’s ATF6 abundance was also not significantly reduced compared with the M32R-mutant groups. The interpretation of this finding is limited by the lack of staining for the cleaved ATF6 protein by the antibody used.

The third UPR arm, PERK, provided the most notable result, as it was found to have the highest level of protein abundance in the group co-transfected with both M32R and S777Q mutant COG5 plasmids, compared with the wildtype and M32R transfection groups, as well as the TM- treated cells (Figure 5.7). The upregulation of PERK in the co-transfected group was not significant compared with protein level in the S777Q-transfected group, but the S777Q group was not significantly upregulated compared with any other group as well. This perhaps implies that the S777Q mutation is the leading source of pathogenicity in the compound heterozygous context of Case B (See Figure 2.2, Table 2.2), but further studies are required to confirm this

79 notion. Since the double-mutant-transfected groups were meant to mimic the compound heterozygous genetics of the patients in Case B as close as possible, this provides us with a possible agent responsible for causing the pathogenic phenotypes of the probands. PERK is responsible for stopping global protein translation, and its activation also upregulates proapoptotic factors87,105. However, this conflicts with the findings that PERK protects cells with mutant rhodopsin from dying and that its inhibition led to increased photoreceptor death112. In this study, PERK upregulation either ultimately leads to cell death, which is responsible for retinal degeneration, or it promotes survival of toxic cells harbouring defected proteins because of COG5 mutations, that indirectly result in retinal degeneration. To investigate these ideas, the effect of mutant COG5 genes on proper Golgi colocalization and inducing DNA damage were studied.

Co-localization analysis revealed that GFP-tagged COG5 protein was co-localized with the Golgi at a similar rate across all transfection groups (Figure 5.9). This did not correspond or explain the observed phenotypic effect of mutant COG5 transfection on Golgi and GFP-tagged COG5 distribution around the nucleus. GM130 was used as a cis-Golgi marker for this analysis due it being insensitive to COG deficiency or overexpression in mutant cell lines153. A COG- sensitive cis-Golgi marker, giantin, was also used to measure co-localization and while it also did not show a significant difference between transfection groups, the average colocalization coefficient in the WT group was notably higher than the single-mutant or double-mutant transfection groups (Appendix, Figures A.2 and A.3). There is an inherent limitation to any colocalization analysis of COG5 with the Golgi apparatus because while the COG complex is associated to the Golgi, it is not found in the Golgi lumen, but is cytosolic in nature. Additionally, the COG complex tethering stability and location is influenced by various interactors, and it is constantly shuttled along the Golgi cisternae and to neighbouring organelles154. A better method of co-localization might be to immuno-stain for another lobe B COG subunit, such as COG7, as they are likely to co-localize due to their strong bond59. However, it is unknown how the transfection of the wildtype and mutant COG5 plasmids would influence the localization of other COG subunits and this must be assessed.

The observed Golgi distribution phenotype was quantified by measuring the relative Golgi area compared to a control, and it was found the mutant transfection groups showed a significantly higher level of dispersal compared to the wild-type transfected group. Moreover, the double-

80 mutant transfection group was significantly higher than the single-mutant transfection groups, signifying a compounded effect of the COG5 mutation on Golgi dispersal and fragmentation (Figure 5.10). While Golgi dispersal has been witnessed in various diseases, as mentioned previously, a landmark study found that DNA damage induces Golgi fragmentation47. DNA damage, as a result, was investigated in the different transfection groups by measuring the fluorescence intensity level of γ-H2A.X, a marker for double-stranded breaks in DNA and is involved in the repair process155,156. In studies that used UV irradiation to induce DNA damage, γ-H2A.X recruitment was found to occur within minutes, rendering it a very sensitive marker for double-stranded DNA breaks155,157. γ-H2A.X recruits other repair factors for assembly at the site of DNA damage and is essential for maintaining chromosomal stability158. In a pattern resembling that of PERK protein expression (Figure 5.7), γ-H2A.X fluorescence intensity was found to be the greatest in the double-mutant transfection group compared with all other transfection groups, including the GFP-control group (Figure 5.12). While some DNA damage is naturally occurring and can be repaired, severe levels of DNA damage can result in apoptosis.

The double-mutant COG5 transfection group demonstrated the highest level of DNA damage (Figure 5.12) and demonstrated the highest level of PERK protein expression (Figure 5.7). The double-mutant group was also the only transfection group to have a significant difference in DNA damage and PERK protein expression compared with the negative and positive controls, respectively. The similarity in expression patterns between γ-H2A.X and PERK led to the assessment of whether both effects are directly related, and whether PERK leads to increased DNA damage, or if DNA damage leads to increased PERK upregulation. This was studied by using a proven PERK inhibitor, GSK2606414, in treating the different transfection groups and assessing whether it has an effect on DNA damage levels159. The PERKi treatment led to a general decrease in the average fluorescence intensity of γ-H2A.X that was only statistically significant for the double-mutant-transfected group (Figure 5.14). This presents a direct link between PERK activation and increase in DNA damage and offers a mode of action for the compound heterozygous COG5 mutations in causing pathogenicity.

Of note is that after PERKi treatment, a significantly higher level of γ-H2A.X fluorescence intensity was observed in the PERKi-treated S777Q-mutant-transfected and double-mutant- transfected groups compared with the PERKi-treated wildtype-transfected and M32R-mutant- transfected groups (Figure 5.14A). This increase was not significantly higher than the level

81 observed in the control group and does not likely indicate a pathogenic level of DNA damage. One possibility is that the S777Q-mutation of COG5 might be the driving force behind the pathogenic nature of the compound heterozygotes. In fact, the M32R mutation has been identified previously as part of another compound heterozygous mutation resulting in COG5- CDG72. That patient had milder symptoms than the probands in Case B and did not present with any observed form of retinal degeneration nor did the patient share any of the phenotypes found in the Family 3 probands. While counting foci is the ideal way of assessing DNA damage, it is a laborious process and is impossible to perform in cells that have an abundance of γ-H2A.X expression where foci cannot be differentiated. Nevertheless, foci counting and measurement of fluorescence intensity were both found to be reproducible and produce comparable results148.

This study has found several new links between a COG complex subunit, UPR activation, and DNA damage that have not been identified before. The presented findings open the gates to new possibilities and investigations that can and should be conducted to better understand the role of COG5 in affecting retinal integrity. On a structural level, the increase in Golgi fragmentation corresponding with the mutant COG5 transfections should be further assessed to identify whether its dispersal is caused by PERK upregulation, DNA damage, or the inherent existence of a mutant COG5 subunit. This will elucidate the structural interaction between the COG5 subunit and the Golgi considering that a definitive model of the COG complex’s retrograde Golgi trafficking and cargo tethering is yet to be definitively established and agreed upon.

Future experiments must also incorporate assessment of patient-derived primary cells, to assess whether the phenotypes documented here are reproducible in cells that do not have an endogenous wild-type copy of COG5. Patient-derived cells will allow us to definitively assess the mode of action that the compound heterozygous COG5 mutations have towards causing pathogenicity, and the extent of DNA damage in the patients that may result in retinal degeneration. Alternatively, transgenic cell lines that only express the mutant variants of COG5, or cells that are silenced for endogenous COG5 and transfected with the mutant variants can also be assessed. However, the patient-derived cells will allow the testing and development of the PERK inhibitor as a potential therapeutic drug to combat the effect of the compound heterozygous COG5 mutations. The COG5 mutations described are not good candidates for gene therapy since the mutant COG5 proteins will remain present alongside the supplemented healthy COG5 protein. The interactions and disruptions of the mutant COG5 protein will likely continue

82 to proceed alongside the normal interactions of the healthy COG5 protein. However, the healthy COG5 protein could alleviate some of the phenotypes presented as could have potentially happened by the endogenous COG5 present within the transfected cells. Future studies can investigate the effect of overexpressing the wildtype COG5 plasmid alongside the mutant COG5 plasmids to assess if it reduces Golgi dispersal, PERK upregulation, or levels of DNA damage.

In conclusion, this study is novel in identifying a direct effect between expressing mutated COG5 in cells and both upregulating UPR stress marker PERK and inducing DNA damage. A schematic summarizing the novel findings of this study is presented in Figure 6.1 This schematic begins with the hypothesized effect of the mutant COG5 protein in causing aggregation of misfolded proteins and inducing ER stress and the UPR. It highlights the identified upregulation of PERK in the double-mutant transfection group (Figure 5.7) followed by the increase in DNA damage (Figure 5.12) that was alleviated through PERKi treatment (Figure 5.14B). While previous studies have identified integral components downstream of PERK activation that are necessary for γ-H2A.X recruitment and DNA damage109, the exact mechanism of how PERK upregulation causes DNA damages remains unknown. The schematic also posits the observed Golgi fragmentation (Figure 5.10) as an effect of DNA damage due to a previous study that showed that DNA damage triggers Golgi dispersal47. However, it must be noted that in this context, we have not presented evidence of DNA damage directly causing Golgi fragmentation. Golgi dispersal may have directly resulted from the mutant COG5 subunit disrupting Golgi’s structural integrity or due to the aggregation of misfolded proteins leading to Golgi structural deficits.

While this study looks at specific COG5 mutations relevant to the patients in Case B with LCA, the findings of the study explore the broader role that COG5 plays in maintaining proteostasis and sensitizing the cell to DNA damage. Previous studies have demonstrated that COP1 mediates the sensitization of DNA damage in the cell as a result of ER stress, and that COP1 works with the COG complex in retrograde Golgi trafficking and localization of Golgi-resident proteins153,160. While these studies were investigated with regards to cancer and tumorigenesis, the findings presented here further contribute to this body of knowledge and expand its application to neurodegenerative and vision loss disorders. Mutations in the COG complex have been found to cause several, debilitating CDGs and it is known that a disruption in one COG subunit, will disrupt the integrity and function of other subunits within the same lobe or

83 subcomplex161. Thus by understanding the specific effect of COG5 disruption, we can also understand the effect of mutations in other subunits involved in causing diseases, such as mutations or deficiencies in COG7 and COG669,162. Ultimately, the continued identification of novel mutations and subsequent investigations into the molecular effects of such mutations will lead to increased development of therapeutics and targeting molecular effectors to alleviate the disruptions and restore function.

Figure 6.1: Schematic summarizing the findings of the current study and positing potential links between the observed results. The hypothesis of the study suggested that due to COG5’s key role in Golgi retrograde trafficking and localization of glycosylation enzymes, a disruptive mutation in the subunit will likely lead to several impairments that could lead to an aggregation of misfolded proteins. The aggregation of misfolded proteins is known to lead to ER stress and results in the activation of the UPR. The UPR has three interdependent branches that are activated to restore proteostasis, and this study found that only the PERK arm is activated when the identified compound heterozygous COG5 mutations are both expressed. After finding that PERKi-treatment reduces DNA damage, we found a direct link between PERK upregulation in this context and the induction of DNA damage. However, PERK’s mechanism of action towards causing double-stranded DNA breaks remains unknown. Due to a previous study that directly links DNA damage to trigger Golgi dispersal47, the schematic here draws a connection between DNA damage and the observed Golgi fragmentation. Nevertheless, the possibility of Golgi dispersal to have been caused by a different mechanism, such as directly due to the COG5 mutation or aggregation of misfolded proteins remains and must be considered.

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A. Appendix A.1 Western blot expression of COG5 plasmids

Figure A.1: Image of a western blot confirming the expression of the wildtype and mutant variants of 6x-His-tagged COG5 plasmids when transfected in vitro. The same blot was immunostained for the 6x-His tag using an anti-6x-His antibody and then stripped and re-stained with an anti-COG5 antibody. GAPDH was used to confirm equal loading between samples.

105 106

A.2 Co-localization analysis of COG5 with Giantin

Figure A.2: Representative images of N1E-115 cells transfected with GFP-tagged wild-type (WT) COG5, mutant (M32R; S777Q) COG5, or co-transfected with both mutant COG5 plasmids (M32R + S777Q). The cells were stained for the Golgi using COG-sensitive cis-Golgi marker, giantin. DAPI was used for nuclear staining. Scale bar is 20μm

107

Figure A.3: Quantification of the colocalization between the GFP that is tagged to the COG5 constructs and giantin, a COG-sensitive cis-Golgi marker, in cells transfected with wild-type (WT) COG5, mutant (M32R; S777Q) COG5, or co-transfected with both mutant COG5 plasmids (M32R + S777Q). For each group, >85 cells were quantified across n = 3 experiments. Statistical analysis was performed using one-way ANOVA analysis followed by Tukey post-hoc test. Error bars = S.E.M.