THE ROLE OF MUSCLE AND NERVE IN

DISSERTATION

Presented in partial fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

By

Chitra C. Iyer

The Ohio State Biochemistry Graduate Program

The Ohio State University

2016

Dissertation Committee

Dr. Arthur H. M. Burghes, Advisor

Dr. Daniel J. Battle

Dr. Brian K. Kaspar

Dr. Jiyan Ma

Copyright by

Chitra C. Iyer

2016

ABSTRACT

Spinal Muscular Atrophy (SMA) is the leading genetic cause of infant death, affecting approximately 1 in 10,000 live births worldwide. SMA is caused due to decrease in levels of the ubiquitous Survival Motor Neuron (SMN) . SMN in humans is encoded by two SMN1 and SMN2. SMA is an autosomal recessive disease caused due to deletion or of SMN1 and retention of SMN2. Due to a

CT change in SMN2, the produces only small amounts of full-length SMN protein. In SMA patients, the low levels of SMN lead to degeneration of motor neurons and muscle atrophy. Since SMA is characterized by muscle atrophy, blocking of muscle ligases that degrade sarcomeric is a prospective therapy. We deleted two muscle-specific ubiquitin ligases, MAFbx and MuRF1, in severe SMA mice. Deletion of MAFbx did not improve the phenotype or survival of SMA mice, however a mild improvement in fiber size distribution was observed. MuRF1 deletion in the SMA mice turned out to be deleterious with majority of mice dying on the day of birth. MAFbx and

MuRF1 levels are upregulated in the skeletal and cardiac muscle in SMA mice whereas

MuRF2 and MuRF3 levels are unchanged. We conclude that deletion of muscle ubiquitin ligases in SMA does not improve the phenotype of SMA mice. Furthermore, it is unlikely that the beneficial effect of HDAC inhibitors in SMA is mediated through MAFbx and

MuRF1 as hypothesized in other studies.

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With various clinical trials in action in the field of SMA, understanding the spatial requirement of SMN protein is crucial. SMN is a ubiquitous protein but a decrease of

SMN in SMA causes degeneration of motor neurons and muscle atrophy. To study the relative importance of either tissue, we employed tissue specific Cre drivers and floxed mouse Smn alleles. We selectively decreased SMN in the neurons or the muscle and conversely restored SMN levels in the neurons or the muscle. Our studies show that 2 copies of SMN2 (and SMN∆7) produce sufficient SMN for the normal function of muscle. Decreasing SMN to SMA levels in skeletal muscle, using Myf5-Cre driver, does not affect the mice. Importantly, the force production capacity of the muscle, as measured by twitch, tetanic and eccentric forces, is unaltered. The muscle fiber size distribution and electrocardiogram of mice with low SMN in muscle is comparable to normal.

Replacement of SMN levels in solely the muscle does not improve the SMA phenotype.

In the neuron set of experiments, we found that decreasing or restoring SMN in only the motor neurons with ChAT-Cre significantly alters the functional output of the motor unit as measured by compound motor action potential and motor unit number estimation.

However ChAT-Cre alone did not alter the survival of SMA mice upon Smn-replacement and did not appreciably affect survival when used to delete Smn. On the other hand, replacement of Smn in both neurons and glia in addition to the motor neurons (Nestin-Cre and ChAT-Cre) resulted in the greatest improvement in survival of the mouse and in some instances complete rescue was achieved. These findings demonstrate that high expression of SMN in the motor neuron is both necessary and sufficient for proper

iii function of the motor unit. Furthermore, in the mouse high expression of SMN in neurons and glia, in addition to motor neurons has a major impact on survival.

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To all my teachers

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ACKNOWLEDGEMENTS

The Burghes Lab has been a perfect balance of work, entertainment and humor.

From 2009 to now, it has been an enriching journey and the Burghes Lab has been an active part of it, for I have spent more wakeful hours in the lab than in my apartment. I sincerely thank all the past and present members of the lab for all the help when needed, for the everyday interactions and for the beautiful friendships that have grown. I started my lab rotation with Thanh and he has been so helpful throughout, from all the little experiment-related technicalities to driving me around to pick my first car. Thank you,

Thanh. Special thanks to Kaitlyn; in the past year, she has been a huge help with the mice and genotyping. Corey, Anton, Isaac, Aurelie and Barb have provided the much-needed entertainment throughout while Ron cured all the hiccups of the instruments promptly. I thank Dave, Paul and Dr. Battle for insightful discussions every once in a while, especially during the preparation for candidacy. Vicki has been a second mentor to me.

She trained me in all the techniques, took care of my mice during my trips back home, and has critically read and corrected my manuscripts right from candidacy. Thanks a ton,

Vicki. I am grateful to my dissertation committee, Dr. Battle, Dr. Kaspar and Dr. Ma, for their encouragement. And now last and most important is King Arthur. Arthur has been a dedicated teacher, mentor and guide. Whether it is opening the tight liquid nitrogen tank for me or repairing my bicycle, Arthur has been eager to teach and help. Thank you for

vi enhancing my scientific thinking, allowing me to learn from mistakes and think on my own and of course, for honing my sarcasm skills.

I am grateful to all my family and friends for being there for me; I shall refrain from the names as it is a very long list. The family and friends on this continent however deserve a special mention for they made America a home away from home. My doting niblings, Shreya, Akshaya, Shruti and Raghav have been the color of my life. Having

Sangeeta, Anu, Karthik and Shashank some hours away is a blessing. I thank Janaki

Periamma for being my ‘periya amma’. In this journey, I consider myself fortunate to have found Amishaben as my guru; you have generated a lasting spark in me. Amma,

Appa and Uttara, you sent me so far to pursue my dreams, how do I thank you? I wish I could see you all more often. And Vikram, my soulmate, you have been my best discovery in graduate school. I could not have done this without you.

Chitra Iyer 4th December, 2015 Columbus, Ohio

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VITA

March 2004………………………………...Higher Secondary, Gujarat State Board, India

April 2007…………………..B.Sc. Biochemistry, St.Xavier’s College, Ahmedabad, India

April 2009…………...M.Sc. Biochemistry, Maharaja Sayajirao University, Baroda, India

September 2009 - December 2015………………………….Graduate Research Associate,

Ohio State Biochemistry Program,

Department of Biological Chemistry and Pharmacology,

The Ohio State University, Columbus OH, USA

PUBLICATIONS

1.) *Chitra C. Iyer, *Vicki L. McGovern, Jason D. Murray, Sara E. Gombash, Phillip

G. Zaworski, Kevin D. Foust, Paul M. Janssen, Arthur H.M. Burghes. Low levels of

Survival Motor Neuron protein are sufficient for normal muscle function in the

SMNΔ7 mouse model of SMA. Human Molecular Genetics 2015; 24 (21): 6160-

6173. PMID: 26276812. (*Co-first authors)

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2.) *Vicki L. McGovern, *Chitra C. Iyer, W. David Arnold, Sara E. Gombash, Phillip

G. Zaworski, Anton J. Blatnik III, Kevin D. Foust, Arthur H.M. Burghes. SMN

expression is required in motor neurons to rescue electrophysiological deficits in the

SMNΔ7 mouse model of SMA. Human Molecular Genetics 2015; 24 (19): 5524-

5541. PMID: 26206889. (*Co-first authors)

3.) Sara E. Gombash, Christopher J. Cowley, Julie A. Fitzgerald, Chitra C. Iyer, David

Fried, Vicki L. McGovern, Kent C. Williams, Arthur H.M. Burghes, Fedias L.

Christofi, Brian D. Gulbransen, Kevin D. Foust. SMN deficiency disrupts

gastrointestinal and enteric nervous system function in mice. Human Molecular

Genetics 2015; 24 (13): 3847-3860. PMID: 25859009.

4.) Chitra C. Iyer, Vicki L. McGovern, Dawnne O. Wise, David J. Glass, Arthur H.M.

Burghes. Deletion of atrophy enhancing genes fails to ameliorate the phenotype in a

mouse model of spinal muscular atrophy. Neuromuscular Disorders 2014; 24: 436-

444. PMID: 24656734.

5.) W. David Arnold, Paul N. Porensky, Vicki L. McGovern, Chitra C. Iyer, Sandra

Duque, Xiaobai Li, Kathrin Meyer, Leah Schmelzer, Brian K. Kaspar, Stephen J.

Kolb, John T. Kissel, Arthur H.M. Burghes. Electrophysiological biomarkers in

spinal muscular atrophy: proof of concept. Annals of Clinical and Translational

Neurology 2014; 1(1): 34-44. PMID: 24511555.

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Fields of Study

Major Field: Biochemistry

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TABLE OF CONTENTS

Abstract……………………………………………………………………………………ii

Dedication………………………………………………………………………………....v

Acknowledgements……………………………………………………………………….vi

Vita……………………………………………………………………………………...viii

List of Tables…………………………………………………………………………....xvi

List of Figures…………………………………………………………………………..xvii

List of Symbols and Abbreviations………………………………………………………xx

Chapters:

1. Introduction……………………………………………………………………………1

1.1 SMA – history, genetics and classification………………………………………..1

1.2 Diagnosis of Spinal Muscular Atrophy (SMA)…………………………………...6

1.2.1 Histology and electrophysiology in SMA patients………………………..6

1.2.2 SMN1 and SMN2 copy numbers…………………………………………..9

1.3 Survival Motor Neuron (SMN) protein and its roles…………………………….12

1.3.1 SMN is ubiquitously expressed and localized to gems in the nucleus…..12

1.3.2 Differential splicing of SMN2……………………………………………13 xi

1.3.3 Domains of SMN and the SMN complex………………………………..15

1.3.4 Role of the SMN complex in snRNP assembly………………………….19

1.3.5 Other postulated roles of SMN…………………………………………..22

1.4 Animal models of Spinal Muscular Atrophy…………………………………….24

1.4.1 C. elegans and Drosophila models of SMA……………………………...25

1.4.2 Zebrafish model of SMA………………………………………………...27

1.4.3 Mouse models of SMA…………………………………………………..30

1.4.4 Pig model of SMA……………………………………………………….37

1.5 Therapy for Spinal Muscular Atrophy…………………………………………...38

1.5.1 Increasing production of full-length SMN from SMN2………………….38

1.5.2 SMN Gene therapy………………………………………………………41

1.5.3 Additional strategies……………………………………………………..44

1.6 Significance of the study…………………………………………………………45

2. Deletion of atrophy enhancing genes fails to ameliorate the phenotype in a mouse

model of spinal muscular atrophy……………………………………………………48

2.1 Introduction………………………………………………………………………48

2.2 Materials and Methods…………………………………………………………...51

2.2.1 Mouse strains and breeding……………………………………………...51

2.2.2 Genotyping and weighing………………………………………………..52

2.2.3 Muscle fiber analysis…………………………………………………….52

2.2.4 Statistical Analyses….…………………………………………………...53

2.2.5 Droplet digital PCR (ddPCR)……………………………………………53

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2.3 Results……………………………………………………………………………54

2.3.1 Generation of the transgenic mouse lines………………………………..54

2.3.2 Weight and survival analyses……………………………………………55

2.3.3 Analyses of muscle morphology…………………………………………57

2.3.4 Expression of the muscle ubiquitin ligases in SMA mice……………….59

2.4 Discussion……………………………………………………………….……….63

3. Low levels of Survival Motor Neuron protein are sufficient for normal muscle

function in the SMN∆7 mouse model of SMA………………………………………67

3.1 Introduction………………………………………………………………………67

3.2 Materials and Methods…………………………………………………………...69

3.2.1 Mouse breeding…………………………………………………………..69

3.2.2 Genotyping, weighing and phenotypic assessment of mice……………..73

3.2.3 Hematoxylin and Eosin staining for fiber size distribution……………...74

3.2.4 Immunohistochemistry…………………………………………………..75

3.2.5 Muscle physiology tests………………………………………………….76

3.2.6 H&E staining for LCM…………………………………………………..77

3.2.7 Laser-capture Microdissection (LCM)…………………………………..77

3.2.8 Digital droplet PCR (ddPCR)……………………………………………78

3.2.9 ELISA on whole muscle sections………………………………………..78

3.2.10 Western blot analysis…………………………………………………….79

3.2.11 Statistical analyses……………………………………………………….79

3.3 Results……………………………………………………………………………80

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3.3.1 Testing the deletion and replacement Smn alleles……………………….80

3.3.2 Expression of Myf5-Cre driver…………………………………………..84

3.3.3 Deletion and replacement of Smn in muscle tissue in the SMN∆7 SMA

mouse model…………………………………………………………….88

3.3.4 Efficiency of Cre recombination in the muscle…………………………90

3.3.5 Determination of SMN protein levels in muscle………………………..91

3.3.6 Functional analyses of muscle force upon Smn-deletion………………...94

3.4 Discussion………………………………………………………………………..97

4. SMN expression is required in motor neurons to rescue electrophysiological deficits

in the SMN∆7 mouse model of SMA………………………………………………105

4.1 Introduction……………………………………………………………………..105

4.2 Materials and Methods………………………………………………………….107

4.2.1 Mouse breeding…………………………………………………………107

4.2.2 Genotyping……………………………………………………………...110

4.2.3 Phenotypic assessment of mice…………………………………………111

4.2.4 Immunohistochemistry…………………………………………………112

4.2.5 Muscle fiber size analysis………………………………………………113

4.2.6 Laser capture microdissection…………………………………………..113

4.2.7 Droplet digital PCR……………………………………………………..114

4.2.8 ELISA on LCM collected tissue………………………………………..115

4.2.9 Western blot analysis…………………………………………………...115

4.2.10 Gem counts……………………………………………………………..116

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4.2.11 Electrophysiological studies of CMAP and MUNE……………………116

4.2.12 Statistical analyses……………………………………………………...117

4.3 Results…………………………………………………………………………..117

4.3.1 Specific deletion and replacement of SMN upon Cre-mediated

recombination in neural tissue………………………………………….117

4.3.2 Characterization of Cre driver using tdTomato-RFP expression……….122

4.3.3 Survival and weight of mice are improved upon replacement of Smn in

neural tissue…………………………………………………………….131

4.3.4 Muscle fiber size is decreased upon deletion of Smn in neural tissue….135

4.3.5 Percent recombination events in LCM isolated motor neuron as

determined by ddPCR…………………………………………………..138

4.3.6 Determination of SMN protein levels in total spinal cord and motor

neurons………………………………………………………………….139

4.3.7 Electrophysiological studies of the functional output of the motor neuron

upon deletion and replacement of Smn…………………………………144

4.3.8 Other neural Cre drivers used in this study……………………………..148

4.4 Discussion………………………………………………………………………154

5. Conclusions and Future Directions…………………………………………………162

References…………………………………………………………..…………………..167

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LIST OF TABLES

Table 1.1. Classification of Spinal Muscular Atrophy .………………………………….5

Table 1.2. Electrophysiological findings in patients with SMA after onset of overt clinical signs of weakness…………………………………………………………………9

Table 1.3. Summary of the mouse models in SMA……………………………………..35

Table 3.1. Cre drivers……………………………………………………………………71

Table 3.2. Floxed alleles………………………………………………………………...71

Table 3.3. Primer sequences…………………………………………………………….74

Table 4.1. Cre drivers…………………………………………………………………..109

Table 4.2. Floxed alleles……………………………………………………………….110

Table 4.3. Primer sequences…………………………………………………………...111

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LIST OF FIGURES

Figure 1.1. Representative H&E stained muscle sections of SMA patients……………...7

Figure 1.2. SMN1 and SMN2 genes……………………………………………………..15

Figure 1.3. Diagram of SMN protein showing exons and domains………….…………17

Figure 1.4. A schematic illustration of the known components and interactions within the

SMN complex……………………………………………………………………………18

Figure 1.5. Simplified illustration of the role of SMN complex in snRNP assembly…..20

Figure 1.6. A photograph showing a non-SMA control mouse and two SMNΔ7 SMA mice at PND13…………………………………………………………………………...33

Figure 2.1. Weight and survival analyses of MAFbx-/--SMA and MuRF1-/--SMA animals…………………………………………………………………………………...56

Figure 2.2. Gastrocnemius muscle fiber of PND08 MAFbx-/--SMA and SMA animals..58

Figure 2.3. Quantification of MAFbx and MuRF1 transcripts (by ddPCR) at PND01,

PND 05, PND08 and PND14 of Smn+/+ Control and SMA animals……………………..61

Figure 2.4. Quantification of MuRF2p50A, MuRF2p60A and MuRF3 transcripts (by ddPCR) at PND01, PND 05, PND08 and PND14 of Smn+/+ Control and SMA animals..62

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Figure 3.1. Sequence of SmnINV and SmnRe alleles……………………………………...72

Figure 3.2. Smn deletion and replacement alleles used in the study…………………….82

Figure 3.3. Immunohistochemistry of Myf5-Cre; tdTomato; HB9:GFP mice

(PND12)………………………………………………………………………………….85

Figure 3.4. Immunohistochemistry of Myf5-Cre; tdTomato; HB9:GFP mice (PND12) indicating ectopic expression of the Myf5-Cre driver……………………………………86

Figure 3.5. Survival and weight analyses of muscle driver Myf5-Cre…………………..89

Figure 3.6. Immunohistochemical localization of SMN in gastrocnemius muscle sections

(PND10)………………………………………………………………………………….90

Figure 3.7. Analysis of Cre activity upon deletion of Smn in muscle…………………..93

Figure 3.8. Muscle physiology tests on mice with SMN reduction in the muscle, Myf5-

Cre; SmnD7/KO, at 8 weeks………………………………………………………………..95

Figure 3.9. Cardiac physiology tests on unanaesthetized mice with SMN reduction in the muscle, Myf5-Cre; SmnD7/KO, at 8 weeks………………………………………………...97

Figure 4.1. Diagram of Floxed Smn alleles and crosses used in this study……………120

Figure 4.2. Nestin-Cre and ChAT-Cre expression in the spinal cord………………….125

Figure 4.3. The expression of Nestin-Cre driver in the spinal cord……………………127

Figure 4.4. Limited expression of Nestin-Cre in several tissues………………………128

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Figure 4.5. Whole body sagittal section of Nestin-Cre + ChAT-Cre tdTomato RFP expression at P2………………………………………………………………………...130

Figure 4.6. Survival and weight of SMA mice is improved upon replacement of Smn with neuron specific Cre drivers………………………………………………………..133

Figure 4.7. Muscle fiber size upon deletion with Nestin-Cre and ChAT-Cre………….136

Figure 4.8. Determination of Cre activity to delete or replace Smn…………………..141

Figure 4.9. Motor neuron and spinal cord SMN levels in Cre modified SMNΔ7 mice……………………………………………………………………………………..143

Figure 4.10. Electrophysiological measures of motor unit function following targeted

Smn deletion and restoration……………………………………………………………146

Figure 4.11. Representative electrophysiological waveforms…………………………148

Figure 4.12. Survival and weight of mice upon deletion and replacement of Smn with

SYN1-iCre………………………………………………………………………………149

Figure 4.13. Ubiquitous RFP expression of SYN1-iCre……………………………….150

Figure 4.14. Survival and weight of mice upon deletion and replacement of Smn with rSyn1-Cre and Gad2-Cre……………………………………………………………….153

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LIST OF SYMBOLS AND ABBREVIATIONS

ASO anti-sense oligonucleotide

BBB blood-brain barrier

BCA bicinchoninic acid

ChAT choline acetyl transferase

CMAP compound motor action potential ddPCR droplet digital PCR

DNA deoxyribonucleic acid

ECL electrochemiluminiscence

EDL extensor digitorum longus

EDTA ethylene diamine tetraacetic acid

EIM electrical impedance myography

ELISA enzyme-linked immunosorbent assay

EMG electromyogram

H&E hematoxylin and eosin xx

FL-SMN full-length Survival Motor Neuron

GFP green fluorescent protein

HDAC histone deacetylase

HRP horseradish peroxidase

ICV intra-cerebroventricular

ISS-N1 intronic splice silencer N1

LCM laser-capture microdissection

MAFbx Muscle Atrogin F-box

MN motor neuron

MO morpholino

MUNE motor unit number estimate

MUP motor unit potential

MuRF1 Muscle RING Finger 1

NMJ neuromuscular junction

PCR polymerase chain reaction

PBS phosphate buffered saline

PND post-natal day xxi

RFP red fluorescent protein scAAV self-complementary adeno-associated virus

SDS sodium dodecyl sulphate shRNA short-hairpin ribonucleic acid

SMA Spinal Muscular Atrophy

SMN Survival Motor Neuron

SMN∆7 SMN with exon 7 deletion

SMUP single motor unit potential snoRNA small nucleolar RNA snRNA small nuclear RNA snRNP small nuclear ribonucleoprotein

RNA ribonucleic acid

TRIM tripartite motif

TSA trichostatin A

UPS ubiquitin proteasome system

WT wild-type

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CHAPTER 1

INTRODUCTION

1.1 SMA – history, genetics and classification

Proximal Spinal Muscular Atrophy of 5qSMA, simply known as SMA, presents as degeneration of spinal alpha motor neurons and muscle atrophy. SMA, an autosomal recessive disease, has a wide spectrum of onset and severities. SMA is the leading genetic cause of infant death affecting 1 in 10,000 live births (Pearn, 1978). With large scale

DNA screening the pan-ethnic carrier frequency in United States has been estimated to be

1 in 54 (Prior et al., 2010; Sugarman et al., 2012).

The early onset infantile SMA, also called acute SMA or Type I SMA, was first described by Werdnig in 1891 and Hoffmann in 1893; hence it is also known as

Werdnig-Hoffmann disease. They noted the progressive loss of muscle strength, with histology revealing severely atrophic muscle fibers and sparing of the diaphragm.

Autopsy also revealed loss of anterior horn cells of the spinal cord. Further studies characterized abnormalities in the electromyogram (EMG) indicative of motor horn disease, in line with the histology (Brandt, 1950; Buchthal and Clemmesen, 1941;

Buchthal and Olsen, 1970). Phenotypically the babies present with a bell-shaped chest and characteristic frog-like posture due to weak musculature; the former due to

1 diaphragmatic breathing (Brandt, 1950; Byers and Banker, 1961). Sensory or cognitive functions are not impaired. Death occurs in infancy due to respiratory infection.

Kugelberg and Welander in 1956 described a benign, juvenile form of hereditary muscle atrophy, quite distinct from muscle dystrophy (Kugelberg and Welander, 1956). The age of onset is between 2 to 17 years, with atrophy progressing from legs and pelvic girdle to arms and pectoral girdle. These patients can walk, and the disease course is usually benign with little tendency of deterioration. Fasciculations and EMG defects indicative of neurogenic atrophy, are present (Buchthal and Olsen, 1970). This form of SMA is called

Type III SMA or chronic SMA. However Byers and Banker’s analysis categorized the chronic spinal muscular atrophy of Kugelberg-Welander disease as a benign form of the

Werdnig-Hoffmann disease (Byers and Banker, 1961). Additionally, an intermediate form of spinal muscular atrophy was observed, with onset at 2-18 months of age, survival till adolescence and symptoms in between the acute and chronic form (Buchthal and

Olsen, 1970; Byers and Banker, 1961; Dubowitz, 1964; Fried and Emery, 1971). The intermediate form of SMA came to be known as Type II. Type II patients can sit but not walk, and can maintain head posture. This often leads to scoliosis. Fasciculation is occasionally observed in Type II SMA patients. In all three types of SMA, the legs are weaker than the arms. The apparent variation in the age of onset and severity of SMA within siblings and within the same family, accompanied by similar physical, histopathology and electrophysiology findings, suggested it to be a spectrum of the same genetic disease (Byers and Banker, 1961; Dubowitz, 1964; Munsat et al., 1969).

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The first study on incidence of Type I SMA studied in north-east English population revealed an incidence of 1 in ~25,000 births, accompanied by a carrier frequency of 1 in

80 (Pearn, 1973). The incidence of Type II and Type III was determined to be 1 in

~24,000 births with a carrier frequency of 1 in 76-111 in the same population (Pearn,

1978). However, there was still a debate as to whether the continuum of phenotypic presentations of the disease, roughly categorized in three groups, shared genetic homogeneity. The bone of contention was laid to rest with genetic mapping studies. The gene causing types II and III SMA was mapped to a on 5q, in proximity to type I SMA-causing gene (Brzustowicz et al., 1990; Burghes et al., 1994a;

Gilliam et al., 1990; Melki et al., 1990a; Melki et al., 1990b; Simard et al., 1992; Wirth et al., 1994). The chromosomal region was further narrowed down by mapping of multicopy dinucleotide markers closer to the known markers on the 5q SMA-region

(Burghes et al., 1994b; DiDonato et al., 1994). The gene Survival Motor Neuron (SMN) was discovered to be deleted or mutated in all SMA patients (Lefebvre et al., 1995).

Furthermore, a centromeric copy of SMN was found on 5q, which produced SMN transcripts lacking exon 7 (Lefebvre et al., 1995). However the exact mechanism of how

SMN deletion leads to varied severities of SMA was unclear. Further studies revealed that Type I patients have a deletion of telomeric SMN (SMNT) on both ; and Type II and III SMA patients have a deletion of SMNT or a gene conversion of

SMNT to SMNC (centromeric copy), while also possessing extra copies of SMNC gene

(Burghes, 1997; Campbell et al., 1997; McAndrew et al., 1997). The 5q12 SMA region was found to be complex, with multiple copies of genes and markers (Burghes, 1997).

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Type II and III patients and parents were found to have multiple copies of SMNC

(Campbell et al., 1997; McAndrew et al., 1997). In some Type II and III cases, the SMNT had converted to SMNC giving rise to a chimeric allele (Burghes, 1997; van der Steege et al., 1996). Copy number and protein analyses in SMA patients revealed that more the

SMNC copies, more the full-length SMN protein produced and the milder the phenotype

(Coovert et al., 1997). Interestingly, SMN2 copy number varies in the population from 1 to 8, with 10-15% of the population having no SMN2 at all (Mailman et al., 2002; Prior et al., 2010; Vitali et al., 1999). To add to that, unaffected individuals with homozygous

SMN1 deletion and increased copies of SMN2 (up to five) have been found to exist (Prior et al., 2004). Study on discrepant SMA patients in whom the low SMN2 copy number does not explain the mild phenotype has revealed an SMN2 variant, 859GC (Prior et al., 2009). The SMN2 859GC in exon 7 creates an exonic splice enhancer and hence this variant of SMN2 produces more full-length SMN transcripts (Prior et al., 2009). Thus the data so far confirm SMN2 gene to be a modifier of the SMA phenotype.

In summary, humans have two forms of SMN gene, SMN1 (telomeric) and SMN2

(centromeric). SMA is caused by deletion or mutation of SMN1 and retention of SMN2, the copy number of SMN2 being a determinant of severity (Burghes and Beattie, 2009).

The exons of SMN2 are different from SMN1 by 2 nucleotides which do not alter an amino acid, the critical one amongst them is a CT change in exon 7 (Lorson et al.,

1999; Monani et al., 1999). Because of this nucleotide change, SMN2 can produce only small amount of full-length transcripts, while the majority of its transcripts are SMNΔ7, lacking exon 7 (Burghes and Beattie, 2009).

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Based on onset of symptoms and ambulation, SMA can be classified into 4 broad categories as shown in Table 1 (Kolb and Kissel, 2011). Briefly, Type I is the most severe form with onset at 6 months of age and the patients never being able to sit. Death usually occurs by two years of age. This is followed by Type II SMA, patients who can sit but never stand and with onset before 18 months. Type III SMA (patients can stand but not walk) can be further divided into types IIIa and IIIb based on onset of symptoms at 18 months – 3 years and beyond 3 years respectively (Zerres and Rudnik-Schoneborn,

1995). An adult onset category (> 21 years), Type IV SMA was added to the original 3 broad classifications (Zerres and Rudnik-Schoneborn, 1995). Subsequent modifications to the classification added a fetal SMA, Type 0, category wherein the patients show decreased fetal movements before birth and present with asphyxia and severe weakness at birth (Dubowitz, 1999). It is to be noticed that more the copies of SMN2, less severe is the phenotype.

Table 1.1 Classification of Spinal Muscular Atrophy

Natural Age Type Age at Onset Highest function SMN2 no. at Death

0 Prenatal Respiratory support <1 month 1 I 0-6 months Never sit <2 years 2 II <18 months Never stand >2 years 3, 4 III >18 months Stand alone Adult IIIa 18 mo to 3 years Stand alone Adult 3, 4 IIIb >3 years Stand alone Adult 4 IV >21 years Stand alone Adult 4-8

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1.2 Diagnosis of Spinal Muscular Atrophy (SMA)

1.2.1 Histology and electrophysiology in SMA patients

Beyond the standard phenotypic presentation, muscle histology and electrophysiology have historically been the diagnostic procedures for SMA before the discovery of the

SMN gene. All SMA patients present with simple atrophic muscle fibers, both type 1 and

2 muscle fibers. There are muscle fibers at all stages of atrophy, with atrophy in a fascicular pattern indicative of neurogenic atrophy (Byers and Banker, 1961; Dubowitz,

1964). SMA Type I and II show atrophic fibers that are occasionally angulated

(Dubowitz, 1985). The extent of atrophy varies within a muscle, with varying proportions of atrophic, normal and large-sized fibers. Later stages of the disease reveal large hypertrophic fibers interspersed with the atrophic fibers, almost three to four times the normal size for the patient’s age. Fiber type staining shows that the hypertrophic fibers are of the same type indicating they are from sprouting of remaining motor neurons, as a means to compensate for the atrophied fibers (Dubowitz, 1985). Additionally histology also shows fibrosis between the fascicles and absence of central nuclei in muscle fibers

(Dubowitz, 1985). Further, severe SMA cases present with group atrophy of the same fiber type implying loss of innervating motor neuron. Severe cases also show an increase in number of muscle spindles with no pathological defect per se (Dubowitz, 1985).

Figure 1.1 shows representative images of muscle histology (Dubowitz, 1985).

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Figure 1.1. Representative H&E stained muscle sections of SMA patients (Dubowitz, 1985). (A) Severe SMA case (autopsy at 6 months; H&E x 200): Severely atrophied muscle fibers can be seen at the center of the section surrounded by larger re-innervated fibers. (B) Intermediate SMA case (H&E x 330): Large hypertrophic fibers are seen adjacent to the atrophied fibers. There were also some fibers with large unknown inclusions (center). (C) Mild SMA case (H&E x 270): Large group atrophy and clusters of enlarged fibers can be observed.

In their study of milder form of SMA, Kugelberg and Welander observed bundles of atrophic fibers surrounded by normal sized fibers in muscle biopsy studies (Kugelberg and Welander, 1956). The atrophic fibers here are mostly in small clusters and large group atrophy is uncommon (Dubowitz, 1985). Again, fiber type grouping is observed in the large fibers. Dubowitz has clarified the diagnoses further saying that fasciculations of tongue and skeletal muscles accompanied by normal intellectual development and absence of cardiac defects distinguish it from muscle dystrophy (Dubowitz, 1964).

Importantly, serum levels of creatine kinase, transaminase and aldolase are normal in all patients (except a few mild cases), ruling out dystrophy as a diagnosis (Brandt, 1950;

Dubowitz, 1964).

Along with histology, electromyographic (EMG) methods have been and still continue to be a useful diagnostic and prognostic tool in SMA (Arnold and Burghes,

7

2013). A characteristic EMG finding in Type I SMA patients is spontaneous activity in relaxed muscle, even in sleep (Buchthal and Clemmesen, 1943; Buchthal and Olsen,

1970). Other signs of motor neuron disease are also present, namely increase in mean amplitude of motor unit potentials (MUPs), increase in territory or increase in amplitude within a territory (Buchthal and Olsen, 1970). Furthermore, the increase in abnormality seen in needle EMG correlates with the worsening of the disease in patients. Also, in line with histology, the more the hypertrophic fibers and the higher the amplitude and territory of motor unit potential (Buchthal and Olsen, 1970). Hausmanova-Petrucewicz and colleagues classified the three types of SMA and gave a comprehensive account of the differentiating parameters of EMG in the types of SMA (Hausmanowa-Petrusewicz and Karwanska, 1986). The main differentiating factor in EMG is the mean amplitude of single motor unit potential (SMUP). Fibrillations and in particular, fasciculations are more prevalent in Type II and III patients (Hausmanowa-Petrusewicz and Karwanska,

1986). The Kugelberg-Welander form shows a higher amplitude and longer duration of potentials which can be explained by the presence of fewer atrophic fibers and more reinnervation (Hausmanowa-Petrusewicz and Karwanska, 1986; Kugelberg and

Welander, 1956).

Electrophysiological studies after the discovery of SMN gene revealed that the compound motor action potential (CMAP) and the motor unit number estimate (MUNE) correlate to the copy number of SMN2, age and functional capacity of patient (Swoboda et al., 2005). CMAP measures the total functional output of motor units supplying a muscle upon maximum stimulation; while MUNE, as the name suggests, is the estimate

8 of functional motor units in a given muscle (Arnold and Burghes, 2013). In the severe

Type I patients, there is a precipitous decline in CMAP and MUNE, which then plateaus

(Swoboda et al., 2005). Type II patients on the other hand exhibit a modest decline in both parameters, with Type III patients essentially stabilizing after a certain decline

(Swoboda et al., 2005). Studies in pre-symptomatic severe SMA patients show preserved motor unit function which plummets as the symptoms set in (Finkel, 2013; Swoboda et al., 2005). Thus electrophysiology studies are in concert with severity of phenotype and serve as a good marker for disease progression. Table 1.2 summarizes the electrophysiological measures in SMA patients (Arnold and Burghes, 2013).

Table 1.2 Electrophysiological findings in patients with SMA after onset of overt clinical signs of weakness.

Electromyography SMA Decreased Enlarged CMAP MUNE Fibrillations Type recruitment motor units Type I ↓↓↓ ↓↓↓ ++ ↑↑↑ ↑ Type II ↓↓ ↓↓ + ↑↑ ↑↑ Type III ↓ or normal ↓ +/- ↑ ↑↑↑

Another technique called electrical impedance myography (EIM) is being developed to study SMA disease progression and efficacy of therapy. EIM is a non- invasive technique of passing low-intensity, high-frequency electrical current through

9 skin and measuring the resulting surface electrical patterns (Rutkove et al., 2010). SMA

Type II and Type III patients have distinct EIM parameters that are significantly different from healthy controls (Rutkove et al., 2010). Also, SMA patients do not exhibit any change in EIM parameters over the study time of 1.5 years indicating no significant disease progression; whereas healthy children show an alteration corresponding to maturing and growing muscle (Rutkove et al., 2012). Though the mechanism underlying

EIM remains to be understood, EIM does hold potential as a biomarker in SMA and has the advantage of being painless and easily tolerated in children.

1.2.2 SMN1 and SMN2 copy numbers

Studies showing that SMA patients have either a loss of SMN1 or a conversion of

SMN1 to SMN2 established SMN2 as a modifier of SMA (Campbell et al., 1997; Lefebvre et al., 1995; McAndrew et al., 1997). It necessitated the gene dosage analyses of the two

SMN genes. The first diagnostic technique used PCR amplification of SMN exon 8, followed by restriction enzyme digestion that would cut only SMN2 exon 7 or exon 8

(van der Steege et al., 1995). Later, Campbell et al. used pulse field gel electrophoresis of restriction-digested genomic DNA, followed by probe hybridization to distinguish SMN1 and SMN2 (Campbell et al., 1997). While SMN1 is mostly absent in patients, this method is not adequate to indicate the copy number of SMN2 or gene conversion events. Hence a quantitative multiplex PCR to amplify SMN1 and SMN2 with respect to the housekeeping gene CFTR was developed (Burghes, 1997; McAndrew et al., 1997). Moreover, in patients who do not have homozygous deletion of SMN1, further analysis needs to be done to look for within the SMN1 gene (Rochette et al., 1997). A fluorescent 10 allele specific PCR for the detection of the seven most common intragenic SMN1 mutations has been developed (Mailman et al., 2002). Aside from diagnosis, the location of missense mutations in SMN1 and their relation to severity of SMA sheds important light on the roles of SMN protein. More sensitive and specific techniques of SMA screening have been developed over the years with their own pros and cons (Prior et al.,

2011). As an example, oligonucleotide tagged-bead arrays on DNA extracted from blood spots can detect exon 7 deletion in SMN1 (Pyatt et al., 2007). Since SMN1 exon 7 deletion is found in ~95% SMA patients, it can be used for carrier and newborn screening in general population (Prior et al., 2010).

In conclusion, the clinical diagnosis of SMA is not straightforward. Occurrence of chimeric SMN alleles, de novo SMN1 mutations, two SMN1 genes on the same chromosome, and multiple copies of SMN2 can confound the diagnostic testing (Mailman et al., 2001; McAndrew et al., 1997; Prior et al., 2004; Wirth et al., 1999). Ironically, often the test may be ordered to exclude proximal SMA (Mailman et al., 2002). Secondly, rapid denervation after the onset of symptoms offers a very small time window for the administration of effective therapy to salvage the remaining motor neurons. Hence a newborn screening for SMA, in effect, will aid not only in pre-symptomatic administration of available therapy but also pro-active care, especially in milder cases with later onset.

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1.3 Survival Motor Neuron (SMN) protein and its roles

1.3.1 SMN is ubiquitously expressed and localized to gems in the nucleus

Immunological staining of SMN protein in HeLa cells revealed SMN localization in distinct dot-like structures in the nucleus and diffused staining throughout the (Liu and Dreyfuss, 1996). In newly divided cells, the staining showed SMN diffused in the cytoplasm, but absent in the nucleus suggesting a lag phase before SMN assembles in the nucleus (Liu and Dreyfuss, 1996). These dot-like structures with SMN in the nucleus were similar to coiled bodies in size and number and were termed ‘gems’ for ‘Gemini of coiled bodies’ (Liu and Dreyfuss, 1996). Gems are found often adjacent to coiled bodies but they are distinct from coiled bodies. Interestingly, gems respond to various different metabolic conditions in the same way as coiled bodies and thus are dynamic nuclear structures (Liu and Dreyfuss, 1996). Indeed the number of gems directly correlates with the SMA phenotype (Coovert et al., 1997). The presence of gems in SMA patient fibroblast cultures with 0 copies of SMN1 and 2 copies of SMN2 demonstrates that

SMN2 does produce SMN protein (Coovert et al., 1997). Further, examination of SMN protein levels in patient-derived fibroblast cultures show decreased SMN levels in patients. Quantitative western analyses show that SMN is expressed ubiquitously; high levels in brain, spinal cord, kidney and liver and lower in cardiac and skeletal muscle

(Coovert et al., 1997). SMA Type I patients on the other hand have a 100-fold decrease in

SMN protein in the spinal cord (Coovert et al., 1997). In short, disruption of SMN1 gene in patients causes a decrease in gems and SMN protein levels in patients. Unsurprisingly,

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Type I patients have a bigger decrease of SMN and fewer gems than Type III patients

(Coovert et al., 1997; Lefebvre et al., 1997).

1.3.2 Differential splicing of SMN2

The exons of the two genes involved with SMA, SMN1 and SMN2, are remarkably similar and differ by only 2 nucleotides that do not change the amino acid sequence (Lorson et al., 1999; Monani et al., 1999). As mentioned earlier (section 1.1),

SMN2 produced only a small amount of full-length (FL) SMN protein and majority of its transcripts lack exon 7. Secondly, the more the SMN2 copies, the less severe the phenotype of SMA. Taken together, characterizing the mechanism between production of

FL-SMN and SMNΔ7 is critical to understanding SMA. Experiments with SMN2- hybrid constructs showed that the critical difference is a C to T change 6 bp into exon 7 of SMN2 that decreases the efficiency with which exon 7 is incorporated in the final transcript (Lorson et al., 1999). Assessment of transcripts from SMA patient-derived cell lines and tissues show a similar splicing pattern throughout (Lorson et al., 1999). In essence, the splicing defect is due to the CT change and is not restricted to a specific cell-type.

Exon splice enhancers (ESEs) and exon splice silencers (ESSs) are splice modulating sequences in exons that bind specific splicing factors, and thereby either enhance or silence the splicing of the exon respectively. The 5’ end of exon 7 of SMN1 has a heptanucleotide ESE that binds to splicing-related SR (serine-argininine-rich) proteins and ensures its inclusion in the final transcript (Cartegni and Krainer, 2002). The

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CT change, six nucleotides into SMN2’s exon 7, disrupts this ESE and hence decreases binding of the particular SR proteins (Cartegni and Krainer, 2002). Another alternative proposed is that the CT change in SMN2 creates an ESS instead in an otherwise neutral sequence (Kashima and Manley, 2003). The creation of this ESS causes it to bind to a splicing repressor protein, hnRNP A1, preventing exon 7 inclusion in the mRNA

(Kashima and Manley, 2003). It is possible that there occurs a competition between positive and negative splicing regulators, and the end result is exon 7 missing in SMN2’s transcripts. Thus the CT in exon 7 in SMN2 alters an ESE or an ESS resulting in exon

7 being skipped during splicing (Burghes and McGovern, 2010).

The loss of exon 7 in the SMN protein decreases its self-oligomerization efficiency and stability. Therefore, SMN2 produces less functional SMN protein than

SMN1. To top that, SMNΔ7 is more rapidly degraded than FL-SMN (Burnett et al.,

2009). This is illustrated in Figure 1.2 (Burghes and Beattie, 2009). In patients, due to absence of SMN1, the modulation of severity of symptoms thus lies in the FL-SMN produced by the multiple copies of SMN2.

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Figure 1.2. SMN1 and SMN2 genes (Burghes and Beattie, 2009): SMN1 and SMN2 are 99.9% identical in sequence. The critical difference is a CT change in exon 7 of SMN2. This single-nucleotide change affects the splicing of SMN2 such that majority of its transcripts lack exon 7. However, SMN2 produces some full-length SMN which can form functional oligomers. SMN lacking exon 7 is unstable and is degraded rapidly.

1.3.3 Domains of SMN and the SMN complex

The 38 kD ubiquitously expressed SMN protein can oligomerize with itself. The self-association domain lies in exons 6 and 7 (see Figure 1.3). This was proven by co- immunoprecipitation and binding assays. The domain was further defined by a series of mini-SMN constructs bearing only parts of exons 5 through 7 (Lorson et al., 1998). A 15 binding curve of GST-tagged WT SMN to various SMN patient missense mutations shows that the more severe the mutation, the less is its tendency to self-oligomerize

(Lorson et al., 1998). Also, SMNΔ7, the main product of SMN2, has a reduced ability to self-oligomerize (Lorson et al., 1998; Pellizzoni et al., 1999). Thus SMN oligomerization defect correlates to the severity of SMA. In addition, the oligomerization domain of SMN is unique and conserved across species implying a critical function (Lorson et al., 1998).

The oligomerization domain has a tyrosine-glycine-rich region named YG box wherein patient missense mutations are condensed, alluding to its importance in SMN’s function

(Talbot et al., 1997). Exon 2b of SMN is highly conserved across species too and was identified as a site for self-association as well as interaction with Gemin2 (formerly SIP-

1, SMN-interacting protein 1) (Young et al., 2000). Another important domain of SMN is the encoded by exon 3 that binds to the Sm proteins involved in snRNA metabolism (Buhler et al., 1999). In fact the severe patient mutation E134K in the Tudor domain disrupts interaction with Sm proteins (Buhler et al., 1999). As explained in the subsequent section 1.3.4, interaction with Sm proteins and loading them into snRNAs is the main function of SMN, which is hampered in Tudor domain mutations.

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Figure 1.3. Diagram of SMN protein showing exons and domains (Burghes and Beattie, 2009)

SMN oligomerizes and interacts with various proteins to form a large multiprotein complex called the SMN complex. The SMN complex consists of Gemin2, Gemin3,

Gemin4, Gemin5, Gemin7, Gemin8 and UNRIP (UNR-interacting protein) (Figure 1.4)

(Carissimi et al., 2006; Gubitz et al., 2004). SMN, Gemin7 and Gemin8 are at the center of the complex while the other components are bound to them via multiple interactions

(Otter et al., 2007). Amongst these, SMN interacts with itself and Gemins 2, 3 and 8 directly interact with SMN (Otter et al., 2007). Gemin 2 was the first identified protein that colocalizes with SMN in gems and in the cytoplasm (Liu et al., 1997). Experiments of immunoprecipitation, colocalization, yeast two-hybrids and mass spectrometry have thus shown the interaction of the other Gemins in the SMN complex. For instance, Gemin

8 interacts with Gemin6-Gemin7 heterodimer (Carissimi et al., 2006). UNRIP is an atypical part of this macromolecular complex in that it does not colocalize with SMN in gems but binds Gemins 6 and 7 (Carissimi et al., 2005; Grimmler et al., 2005). Gemin5

17 on the other hand is peripherally bound to the SMN complex and specifically binds to the unique Sm site on snRNAs (Battle et al., 2006; Otter et al., 2007). SMN via its Tudor domain directly interacts with the spliceosomal Sm proteins SmB/B’, SmD1 and SmD3 after their methylation (Brahms et al., 2001). Moreover, SMN itself is an oligomer in the complex; the exact stoichiometry is unknown but it ranges from dimer through octamer

(Martin et al., 2012). Even the Gemins exist as oligomers in the SMN complex; their stoichiometry remains to be determined. In Figure 1.4, they are shown as monomers for simplicity. All of these results point towards a role of the SMN complex in spliceosomal snRNP metabolism.

Figure 1.4. A schematic illustration of the known components and interactions within the SMN complex (Carissimi et al., 2006)

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1.3.4 Role of the SMN complex in snRNP assembly

Post-transcriptional metabolism of mRNA requires large macromolecular complexes composed of ribonucleoproteins (RNPs) (Pellizzoni, 2007). Small nuclear

RNPs, abbreviated as , are a complex of proteins and RNA that are a part of the which excises the non-coding out of a pre-mRNA. The major spliceosome consists of five U snRNAs - U1, U2, U4, U5 and U6 while the minor spliceosome has U11, U12, U5, U4atac and U6atac snRNAs (Kambach et al., 1999; Patel and Steitz, 2003). Each snRNP is composed of a U small nuclear RNA (U snRNAs), seven common Sm proteins (B/B’, D1, D2, D3, E, F and G) and a set of specific proteins

(Will and Luhrmann, 2001). The Sm proteins form a heptameric ring around a conserved sequence on the snRNAs called the ‘Sm-site’ (Burghes and Beattie, 2009; Neuenkirchen et al., 2008). Methylation of the arginine-glycine-rich (RG) domains of the Sm proteins

B/B’, D1 and D3 facilitates their binding with the Tudor domain of SMN and conversely

SMN patient mutations within the Tudor domain hamper the interaction with Sm proteins

(Brahms et al., 2001; Buhler et al., 1999; Friesen and Dreyfuss, 2000). The Sm-like protein, LSm4 also possesses a similar RG domain which upon methylation can bind to

SMN’s Tudor domain (Brahms et al., 2001; Friesen and Dreyfuss, 2000).

When antibodies against SMN or Gemin2 were injected in Xenopus oocytes along with radioactive snRNA to test for snRNP assembly on the snRNA, assembly of snRNPs was disrupted (Fischer et al., 1997). Conversely snRNP assembly on U1snRNA could be restored with addition of affinity-purified SMN-Gemin 2 complex (Meister et al., 2001). This gave evidence for a direct role of the SMN complex in snRNP biogenesis. 19

Purified SMN complexes from HeLa cells can assemble the Sm core proteins on in vitro transcribed snRNAs in an ATP-dependent manner (Pellizzoni et al., 2002). It was further shown that the SMN complex imposes an order and specificity to the assembly reaction of Sm ring onto the snRNAs (Pellizzoni et al., 2002). Thus it was proven that the SMN complex is necessary and sufficient for snRNP assembly. Figure 1.5 shows a simplified diagram of SMN-mediated snRNP assembly (Burghes and Beattie, 2009).

Figure 1.5. Simplified illustration of the role of SMN complex in snRNP assembly (modified from (Burghes and Beattie, 2009)): (A) In the cytoplasm, the seven Sm proteins upon methylation bind the SMN complex. (B) The SMN complex consists of an oligomer of SMN (shown in blue), Gemins 2-8 and UNRIP (UNR-interacting protein). (C) snRNA is transcribed in the nucleus and exported to the cytoplasm wherein it binds to Gemin 5. Gemin 5-bound snRNA is brought to the SMN complex. (D) The SMN complex loads the heptameric ring of Sm proteins onto the snRNA. The 7-methyl guanosine cap of snRNA gets hypermethylated, allowing the SMN complex with the snRNA to bind to import proteins which mediate transport of the entire complex into the nucleus. (E) In the nucleus, the SMN complex and snRNPs localize to the and the snRNPs undergo further maturation. 20

The Sm-like LSm proteins, LSm10 and LSm11, replace SmD1 and SmD2 in the heteroheptameric ring around the U7snRNA (Pillai et al., 2003). The U7snRNP is different than the other snRNPs in that it is involved in histone mRNA processing.

Replication-dependent histone mRNAs are distinguished from regular mRNAs by the absence of introns and polyA tails (Schumperli and Pillai, 2004). The U7snRNP is involved in the cleavage of the 3’ end of histone mRNAs which itself is cell cycle regulated (Schumperli and Pillai, 2004). Assembly of the unique Sm/Lsm core around the

U7snRNA is indeed mediated by the SMN complex (Pillai et al., 2003). Incubation of cell extracts with SMN antibodies inhibits U7snRNP assembly, further supporting that the SMN complex is involved (Pillai et al., 2003).

The levels of snRNPs in SMA mice and patients are affected by the low SMN levels. While Gemins 2, 6 and 8 are strongly decreased in SMA mice, Gemin 4 and

UNRIP appear unaffected (Gabanella et al., 2007). Quantification of histone mRNAs in the motor neurons of SMN∆7 SMA mice reveal an increase in unprocessed 3’ ends corresponding to decreased U7snRNA levels (Tisdale et al., 2013). This is corroborated by detection of unprocessed 3’ ends of histone mRNAs in patient spinal cord samples as well (Tisdale et al., 2013). The decrease in SMN affects only a subset of spliceosomal snRNPs though. In the spinal cord of severe SMA mice (Smn-/-; SMN2+/+), a significant decrease in the levels of U1, U2, U11 and U12 snRNA is found (Gabanella et al., 2007).

This is accompanied by a ten-fold decrease in snRNP assembly activity in the spinal cord extracts of severe and SMN∆7 SMA mice (Gabanella et al., 2007).

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Thus there is plenty of evidence indicating that snRNP assembly activity is the main function of SMA. Though other roles have been ascribed to SMN, none of them have definitive proof. One strong hypothesis for the cause of SMA is that the decrease in

SMN causes alteration in snRNP profile and assembly activity which leads to an alteration in splicing of mRNAs important in motor neuron circuitry (Burghes and

Beattie, 2009). U11 and U12 snRNA decrease points towards the minor spliceosome being affected more. The analyses of transcripts of minor introns will shed light on the etiology of SMA. However while exons are conserved across species, the sequence and structure of introns may not be conserved, as in the case of resistin (Ghosh et al., 2003).

So care should be taken while interpreting data from mouse or porcine models in the case of SMA mis-spliced transcripts. Moreover SMA is a disease of degeneration of motor neurons and motor neurons are a small part of the whole spinal cord. Hence analysis of motor neuron tissue exclusively via laser-capture microdissection would give more reliable information.

1.3.5 Other postulated roles of SMN

SMN has been shown to bind to a variety of different proteins. In fact SMN has been called the master ribonucleoprotein assembler (Terns and Terns, 2001). Many RNA- binding proteins have RG/RGG domains which reportedly bind to SMN. The Sm-like,

LSm, proteins are of interest. SMN interacts with LSm4 upon methylation of its RG domain (Brahms et al., 2001; Friesen and Dreyfuss, 2000). LSm4 is a part of the LSm2-8 heptameric ring around the nuclear U6snRNA. However U6snRNA levels are unchanged in SMA and no assay has been designed to study how U6 is affected in SMA (Zhang et 22 al., 2008). LSm4 and LSm6 are also a part of the cytoplasmic LSm1-7 heptameric ring that binds to the 5’ end of mRNA (Tharun, 2009). The LSm1-7 ring is associated with cytoplasmic mRNPs and activates decapping of mRNA and promotes its turnover (He and Parker, 2000; Tharun, 2009). An interesting finding was the detection of SMN in dendritic mRNPs characterized by LSm1 (di Penta et al., 2009). The role of the axonal mRNPs is supposedly mRNA transport for local translation. But defective axonal transport has not been conclusively shown in SMA yet. Whether SMN is involved in the assembly of the LSm1-7 and the LSm2-8 rings remains to be answered.

Studies in SMN∆7 SMA mice unfold a defect in axonal transport and vesicle release at the NMJ (Dale et al., 2011; Kong et al., 2009). SMN has been shown to bind to the nuclear proteins hnRNP-Q and hnRNP-R and form a unique RNP complex that is involved in transport of mRNAs, in particular beta-, down the long axons of motor neurons (Mourelatos et al., 2001; Rossoll et al., 2003; Rossoll et al., 2002). SMN can bind to profilin and Fragile-X mental retardation protein (FMRP) that are part of axonal transport systems (Piazzon et al., 2008; Sharma et al., 2005). The axonal transport hypothesis as the reason for SMA etiology is debatable. SMN is reported to interact with a lot of proteins that are part of RNPs, for example, fibroblast growth factor 2 (FGF-2),

RNA helicase A and co-activator associated methyl transferase 1 (CARM1) to name a few (Cheng et al., 2007; Claus et al., 2004; Pellizzoni et al., 2001). SMN via its interaction with and GAR proteins has been implicated in the assembly of small nucleolar RNAs (snoRNAs) that serve as guides for the methylation of rRNA, tRNA and snRNA in snoRNPs (Jones et al., 2001; Whitehead et al., 2002).

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From the large platter of SMN-interacting proteins, it is crucial to determine which ones are dependent on SMN, are functionally disrupted in SMA and the restoration of which is essential for improving the SMA phenotype.

1.4 Animal models of Spinal Muscular Atrophy (SMA)

Evolutionary research has revealed that the chimpanzees, humans’ closest relatives, are the only other species to possess SMN gene duplication (Rochette et al., 2001). This dates SMN gene duplication to 5 million years ago. But the modification to SMN2 is exclusive to Homo sapiens; SMN2 is present in all extant human populations (Rochette et al., 2001). Thus it can be said that SMA is a disease unique to humans, because no other species has a less-efficient SMN gene copy to produce small amounts of SMN protein.

In all species used to model SMA, complete loss of SMN causes lethality, the extent of survival is dependent on maternal SMN. In addition, SMN and its binding partners are conserved across species too. Starting from lower eukaryotes, an ortholog of SMN gene has not been identified in Saccharomyces cerevisiae. Three independent groups identified yeast SMN in Schizosaccaromyces pombe which was shown to be essential for viability

(Hannus et al., 2000; Owen et al., 2000; Paushkin et al., 2000). Yeast SMN shows high degree of similarity with hSMN at the N- and C-terminals and the conserved YG box.

Also, yeast SMN interacts with a homologous yeast Gemin2/SIP1 protein and human

SMN, SIP1 and Sm proteins. Yeast, S. cerevisiae, due to its lack of SMN, is a useful system to study the interaction of SMN and its partners and the effect of various mutations in the absence of endogenous WT SMN (discussed further in Chapter 5).

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1.4.1 C. elegans and Drosophila models of SMA

The well characterized genomes and nervous systems of Caenorhabditis elegans and

Drosophila melanogaster make them good invertebrate model systems to study SMA and the tissues involved, and SMA with respect to development. C. elegans ortholog of SMN,

CeSMN, is expressed uniformly throughout larval development and upon hatching, in the nuclei of every cell (Miguel-Aliaga et al., 1999). The expression of CeSMN however becomes heterogeneous in adulthood, with higher localization observed in neurons.

Knock-down of CeSMN gave a variety of phenotypes, with decreased embryonic viability to neuronal, muscular and reproductive defects (Miguel-Aliaga et al., 1999).

Various recessive SMN alleles and point-mutations have been studied in Drosophila models of SMA due to the ease of genetic crosses in the fly. Developmental studies show that dSMN is highly expressed in embryogenesis and the levels decrease in adulthood

(Rajendra et al., 2007). Homozygous deletion of Drosophila Smn (dSMN) causes death at various stages of larval development, as the full-length SMN of maternal origin dips

(Chan et al., 2003; Chang et al., 2008; Rajendra et al., 2007). Like SMA patients,

Drosophila larvae with SMN depletion exhibit decrease in motor rhythm, muscle size and locomotion, and aberrant neuromuscular junction transmission (Imlach et al., 2012).

Studies of hypomorphic dSMN mutants reveal a motor dysfunction phenotype, with the flies being incapable of flying or jumping. Defect in motor neuron organization pattern coupled with decreased routing and branching have been observed (Rajendra et al.,

2007). RT-PCR, pull-down and staining assays link SMN’s role to skeletal actin and uncover muscle defects such as decreased thin filaments (Rajendra et al., 2007). However 25 it is difficult to implicate a muscle-exclusive role for SMN from these studies. The hypomorph mentioned above is selective reduction of dSMN in adult thorax only and not ubiquitously. To dissect the relative role of neuron and muscle in SMA, Gal4 tissue- specific drivers were used in fly to express SMN in muscle or neurons (Imlach et al.,

2012). SMN restoration exclusively in muscle or motor neurons did not improve the phenotype or the electrophysiological output. Restoring SMN pan-neuronally fully rescued the muscle defect and electrophysiology (Imlach et al., 2012). Yet no viable adults were produced, a probable reason being complete depletion of SMN in other tissues. Hence a better way to address the question is to maintain low SMN levels in other tissues while replacing SMN levels in muscle or neurons (addressed in Chapters 3 and 4). In our mouse studies, we found improvement in electrophysiology parameters upon SMN restoration in motor neurons too. One reason could be the basic difference in the models – in fly, motor neurons are glutamatergic while interneurons and sensory neurons are cholinergic; the reverse of mouse and humans. An interesting finding by

Imlach and colleagues was that temporal restoration of SMN after embryogenesis and completion of nervous system development completely rescued the larvae suggesting

SMA is not a developmental defect but a defect in maintenance of NMJ (Imlach et al.,

2012). Switching on of inducible SMN expression from 24-48 hours, after larval hatching, led to flies with intermediate phenotypes (Imlach et al., 2012).

Analyses of UsnRNAs in SMN null Drosophila showed a decrease in U1, U5, U11,

U12 and U4atac snRNAs, albeit with no reduction in splicing of minor-introns (Praveen et al., 2012). Expression of WT SMN or mild-mutant SMN in SMA flies rescued the

26 locomotor and viability defects in fly without rescuing the snRNA profile to WT levels, prompting the authors to uncouple snRNP biogenesis and neuromuscular defects

(Praveen et al., 2012). However given the facts that Drosophila has only a handful of minor introns and that no snRNP assembly assays were done on the rescue flies, it is difficult to conclude the above. Nevertheless, creation of human patient-equivalent SMN mutations in Drosophila has shown that human and Drosophila SMN proteins are biochemically similar (Praveen et al., 2014). For example, dSMNM194R and dSMNG206S are defective in self-oligomerisation, consistent with human SMN mutations M263R and

G275S (Clermont et al., 2004; Praveen et al., 2014). Additionally, severe YG-box SMN mutants in Drosophila were rescued by overexpression of WT SMN indicating that for human patients with these point mutations, WT SMN could be an effective therapy

(Praveen et al., 2014). Thus the point-mutation dSMN lines hold the potential to shed more light on SMN and its interacting partners.

1.4.2 Zebrafish model of SMA

Zebrafish (Danio rerio) owing to its well characterized embryonic development, relatively simple neuromuscular system and optical clarity has been exploited to study axonal developmental defects in SMA conditions. Zebrafish smn gene has been knocked down (KD) with morpholino-oligonucleotides (MOs) in embryonic stage to obtain SMA fish (McWhorter et al., 2003). MO-dose dependent defects on motor axon length and branching were observed; the less the SMN, the more the number of truncated axons and the more the branching (McWhorter et al., 2003). High-dose MO KD also led to less percentage of total surviving embryos. Axonal defects were further confirmed by 27 visualization of single motor axons via iontophoresis-mediated KD (McWhorter et al.,

2003). The aforementioned phenotypes confirm that the less the SMN, the more severe is the phenotype. Interestingly, the sensory neurons and interneurons did not show any overt defects indicating that the motor neurons are very sensitive to decreased SMN levels

(McWhorter et al., 2003). Also, development and organization of skeletal muscle remained unaffected upon smn-KD in fish. Our work in mouse model of SMA confirms this finding (discussed in Chapter 3). Additionally, as expected, coinjection of hSMN with the smnMO led to partial rescue of zebrafish embryos. Complete rescue was not obtained likely due to the mosaic nature of the injections.

Missense mutations of SMN have also been studied in the zebrafish model, specifically three mutations at the C-terminal of zebrafish smn (Boon et al., 2009). For all three mutations studied, homozygous fish die in larval stages as the levels of maternal

SMN fall off. Nonetheless, the mutant larvae have been used to investigate the neuromuscular junction in SMA-state. Immunohistochemistry shows that the homozygous mutants have a decreased ratio of presynaptic SV2 protein to postsynaptic acetylcholine receptors (Boon et al., 2009). Yet the presynaptic terminals are present in the mutants, with a decrease in SV2 protein. Motor-neuron exclusive expression of hSMN rescues the SV2-defect, but not the survival, since SMN was not expressed in the other tissues (Boon et al., 2009). Developing a zebrafish model closer to mimicking the situation in humans was done by expressing the human SMN2 gene onto the smn-/- background, thus ensuring low levels of SMN in all tissues (Hao le et al., 2011). Further,

SMN2 expression in the homozygous mutant fish lines rescues the aforementioned SV2-

28 defect at the NMJs (Hao le et al., 2011). The transgenic SMN2 fish not only recapitulates the splicing pattern of SMN2 in humans but also shows an increase in exon 7 inclusion upon injection with ISS-N1 MO, a splice-modulating anti-sense oligonucleotide used in

SMA mice (Hao le et al., 2011; Porensky et al., 2012). These data suggest that the SMN2 zebrafish model could serve as an in vivo system with a fast read-out to study approaches for increasing full-length SMN production from SMN2.

Zebrafish model for SMA has been used to study the temporal requirement of

SMN during motor-neuron development. In zebrafish, maternal SMN from the egg lingers till 6 days post fertilization; this would disallow temporal studies of low SMN on motor axon outgrowth (Hao et al., 2013). Hence maternal:zygotic (mz) smn mutants lacking both maternal and zygotic smn were developed by designing transgenic fish bearing RFP-tagged human SMN1 under an inducible heat shock promoter (Hao et al.,

2013). Minor leakage of the promoter allows very low SMN levels to be maintained during development in the absence of heat shock. This severe reduction in SMN in mz- smn mutants causes less number of axonal branches during development (Hao et al.,

2013). Induction of SMN after motor axon growth did not rescue the defects. Complete rescue was obtained when SMN was induced early before axonal outgrowth indicating the requirement of high SMN levels at that critical stage in development (Hao et al.,

2013). The temporal requirement of SMN in mouse models is discussed in section 1.4.3.

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1.4.3 Mouse models of SMA

With the discovery of SMN1 and SMN2 genes’ involvement in SMA, there arose a need to create a mammalian model to understand the disease. With the identification of the Smn gene in mice (Mus musculus), the first mouse model of SMA was made in 1997

(Schrank et al., 1997). Mouse Smn shares 83% amino acid homology with hSMN, and like its human counterpart is ubiquitously expressed (Schrank et al., 1997). Upon knocking-out of the Smn gene death occurred in early embryonic stages, before blastocyst stage, as the maternal SMN depleted (Schrank et al., 1997). However, in humans the disease is due to decrease in SMN protein, SMN2 being an important part of the etiology.

Thus a mouse with deletion of mouse Smn and retention of SMN2 would be a better model. The severe mouse model for SMA was made by insertion of human SMN2 transgene over the Smn+/- background (Monani et al., 2000). Mice with Smn-/-; SMN2 survive for an average of 4-6 days and exhibit decreased movement, labored breathing and tremor of limbs after the first 48 hours (Monani et al., 2000). The severe SMA mice also show difficulty in righting and a bell-shaped chest likely due to weak intercostal muscles (Monani et al., 2000). A beautiful finding from this study was that mice with high copies of SMN2 (8 copies) do no display any obvious phenotype, appear normal and have normal motor neuron and gem counts. The high copy SMN2 mice suggest an important point – phenotypic severity can be ameliorated by increasing SMN2 copy number, and in turn by increasing SMN protein levels. This data lends weight to the hypothesis that SMN2 gene acts as a modifier of the SMA phenotype (Monani et al.,

2000). Embryonic motor axon development has been studied in the severe SMA mice

30 having GFP expressed under the motor-neuron specific HB9 promoter (McGovern et al.,

2008). The study uncovered no defect in axonal migration or any developmental delay, but defects in occupation and maintenance of synapses (McGovern et al., 2008). Thus the data from zebrafish showing axonal developmental issues should be taken with a grain of salt because the defect in a severe mouse SMA model is at the level of the NMJ. The discrepancy between the two may be that in the latter, SMN2 provides small amounts of

SMN to every cell; while in the fish the only SMN available is of maternal origin.

Another widely used mouse model of SMA has been created by targeted deletion of Smn exon 7, giving an Smn-/- genotype that is embryonic lethal (Hsieh-Li et al., 2000).

Thus the mouse Smn locus in this case is not a null allele because it can produce SMN protein lacking exon 7. Addition of human SMN2 to the above background gives SMA- like mice of varying severity, depending on the copy number of SMN2 (Hsieh-Li et al.,

2000). Smn-/- mice with 2 SMN2 copies have severe SMA and live up to 2 weeks like the

SMNΔ7 SMA mice (Hsieh-Li et al., 2000). SMA mice homozygous for the SMN2 transgene possess 4 SMN2 copies survive normally but show mild SMA phenotype

(Hsieh-Li et al., 2000). These models are often referred to as the Taiwanese SMA mice models.

The majority of transcripts produced by SMN2 are SMNΔ7; and SMNΔ7 protein overexpression was thought to be toxic due to its apparent pro-apoptotic activity in cell culture (Le et al., 2005). The argument was put to rest by introduction of SMNΔ7 cDNA into the severe SMA background. The increase in SMNΔ7 levels is beneficial to the mice and improves survival from 5 days to an average of 13.3 ± 0.3 days (Le et al., 2005). This 31

SMA model with the genotype Smn-/-; SMN2+/+; SMNΔ7+/+, is referred to as the SMNΔ7

SMA mouse model (used for our studies in Chapters 2, 3 and 4). They are weak and show shakiness in hind limbs and are smaller in size (see Figure 1.6). The reason for increased SMNΔ7 helping the phenotype is that it can associate with full-length SMN to form oligomers; and so with sufficient full-length protein, more functional oligomers are formed (Le et al., 2005). In these SMA mice, muscle morphology studies show smaller atrophic myofibers and intact dystrophin complex. Higher proportion of partially innervated or non-innervated NMJs and unoccupied AChR clusters are found in SMNΔ7

SMA mice (Le et al., 2005). Thus the mice possess denervation and atrophy similar to

SMA patients. The increased life span to 2 weeks over the severe mice is helpful for studying etiology and therapies for SMA. Moreover the NMJ denervation pattern has been systematically characterized in the SMNΔ7 SMA model (Ling et al., 2012).

Interestingly, the study shows denervation of clinically relevant muscles and a failure of synapse maintenance; similar to the embryonic study of severe SMA mice mentioned earlier in this section (Ling et al., 2012; McGovern et al., 2008). A final seal on the usefulness of the SMNΔ7 SMA model is the remarkable overlap between the differential vulnerability of muscles in SMA patients and Δ7SMA mice (Iascone et al., 2015). A side note is that the cardiac defects observed in Δ7SMA mice and the necrosis seen upon rescue is not typically found in SMA patients and may be a mouse-specific phenotype

(Bevan et al., 2010; Heier et al., 2010; Porensky et al., 2012; Shababi et al., 2010).

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Figure 1.6. A photograph showing a non-SMA control mouse and two SMNΔ7 SMA mice at PND13 (Le et al., 2005). The SMA mice are considerable smaller in size as compared to the normal control pups and have difficulty in righting themselves.

Mice with SMN missense mutations are very useful to understand the biochemistry of the SMN protein. SMNA2G is a mild missense mutation usually found in

Type III patients. The SMNA2G transgene in Smn-/- background does not rescue the embryonic lethality, but in the presence of two copies of SMN2, a mild SMA model is obtained (Monani et al., 2003). The mild SMA mice, Smn-/-; SMN2+/+; SMNA2G+/-, have a normal lifespan, but are smaller in size with muscle weakness and hindlimb clasping

(Monani et al., 2003). These Type III SMA mice also display motor neuron loss, axonal sprouting, reinnervation and electrophysiological abnormalities, including fibrillation, seen in Type III patients (Monani et al., 2003). Biochemical assays show that the

SMNA2G protein has decreased ability not only to self-associate, but also to associate with SMN-WT protein (Monani et al., 2003). Another mutation SMNA111G, like the above, cannot rescue Smn-/- mice (Workman et al., 2009). In contrast, mice that are Smn-/-

; SMN2+/+; SMNA111G survive for over a year and are comparable to healthy controls in 33 terms of weight and ventral root counts. Chiefly, the snRNP assembly activity in the

SMNA111G mice with SMN2 is comparable to controls (Workman et al., 2009). Thus snRNP assembly may be a critical function of SMN complex. Additionally, the aforementioned study showed that SMNA2G and SMNA111G do not complement each other and hence may be affecting the same domain function of SMN (Workman et al.,

2009). Our laboratory has created more transgenic lines of N- and C-terminal SMN- missense mutations. Complementation experiments between N- and C-terminal mutations and SMN2 show that SMN functions as an oligomer and has distinct domains of function

(personal communication).

SMA mice with a range of SMN2 copy numbers have been generated by incorporating varying number of SMN2 genes at the mouse Smn locus (Osborne et al.,

2012). Using different combinations of the alleles, 0 through 6 and 8 copies of effective

SMN2 dosage have been obtained. Thus, these mice offer a range of severities to study

SMA and test therapies. Also, because they are engineered at a single locus, they can be crossed onto other SMA models for study. Conditional mouse models of SMA with selective deletion and overexpression of SMN in muscle or neurons have also been studied. They are discussed and the study is further extended in Chapters 3 and 4.

Further, two conditional hypomorphic Smn alleles have been generated too (Hammond et al., 2010). Along with being Cre inducible, the alleles mimic human SMN2 splicing.

Hence the splicing is similar to human SMN2, while the promoter is the endogenous mouse Smn promoter. Table 1.3 summarizes the SMA mouse models and allelic series

(Osborne et al., 2012; Park et al., 2010).

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Table 1.3 Summary of the mouse models in SMA

Genotype Mean survival Phenotype Reference (Schrank et Smn-/- Embryonic lethal al., 1997) Smn-/-; SMN2+/+ 5 days Very severe SMA phenotype Smn-/-; SMN2 (Monani et high copy (8 Normal Normal, but have a short, thick tail al., 2000) copies) Most Severe: Max 10 days Range of severe, intermediate and (Hsieh-Li Smn-/-; SMN2 Severe: Max 2-4 mild SMA phenotypes depending on et al., 2000) weeks SMN2 copy number Mild: Normal Smn-/-; SMN2+/+; (Le et al., 13.3 days Severe SMA phenotype SMNΔ7 2005) Smn-/-; SMN2+/+; (Monani et Normal Mild SMA phenotype SMNA2G+/- al., 2003) Smn-/-; SMN2+/+; (Workman Normal Normal SMNA111G et al., 2009) Smn allelic Effective SMN2 dosage; and Mean survival Reference series: phenotype SmnA/A Embryonic lethal 0 SmnA/B Embryonic lethal 1 SmnA/C Embryonic lethal 2 SmnA/D Viable 4; Necrosis onset by 4 weeks of age SmnB/B Embryonic lethal 2 (Osborne et SmnB/C Embryonic lethal 3 al., 2012) SmnB/D Viable 5; Necrosis onset by 4 weeks of age SmnC/C Viable 4; Necrosis onset by PND5 SmnC/D Viable 6; Necrosis onset by 6 weeks of age SmnD/D Viable 8; No necrosis observed

With the availability of inducible models in SMA, a crucial issue is to decipher the spatial and temporal requirement of SMN. The spatial requirement of SMN is the subject of study in Chapters 3 and 4. The studies on temporal requirement of SMN are 35 discussed here. A doxycycline-responsive expression cassette of SMN was designed into the SMNΔ7 SMA background to switch on and off SMN expression as desired (Le et al.,

2011). Induction of SMN expression in the embryonic and neonatal stages resulted in a striking increase in survival with some SMA mice living over 200 days. Earlier the induction, the better the survival; embryonic induction at E13 led to the best rescue.

Ceasing of SMN induction at 28 days of age led to some mice surviving normally while most mice rapidly declined and died within a month (Le et al., 2011). The neuromuscular junction morphology and transmission in all the mice were comparable to healthy controls (Le et al., 2011). Thus it can be concluded that the low amounts of SMN from 2 copies of SMN2 (and SMNΔ7) in the background is sufficient in adulthood, while high levels of SMN are necessary during development and early postnatal period.

Temporal studies on SMN were furthered by conversely depleting SMN down at various time points to that produced by 2 copies of SMN2. The work shows that high

SMN is a must till PND17 in mice, after which NMJ maturation is complete (Kariya et al., 2014). Interestingly, the mice are resistant to SMN depletion in adulthood and exhibit normal NMJs. SMN depletion at PND50 however causes pathology 6 months after removal, i.e. at 230 days of age in mice with 2 copies of SMN2. The aged mice displayed central muscle nuclei indicative of muscle regeneration albeit with no decrease in muscle fiber size. The mice also showed modest NMJ deformities along with a striking loss of gems in MNs. Interestingly, upon nerve-crush, SMN-depleted mice show defective assembly and maturation of NMJ, followed by muscle atrophy (Kariya et al., 2014). An important finding in this work was that in control animals, SMN levels peaked at day 30

36 after nerve-crush during the time of NMJ maturation and declined to baseline after repair

(Kariya et al., 2014). Therefore, SMN seems to be important for regeneration processes after nerve injury. Thus 2 SMN2 copies produce enough protein for a normal phenotype in adulthood after developmental maturation of NMJ, but repair mechanisms of NMJ seem to demand more SMN.

1.4.4 Pig model of SMA

The domestic swine, Sus scrofa, is closer to humans as compared to mice in terms of genomics, lifespan, cardiovascular system, physiology and nervous system (Lorson et al., 2008; Lorson et al., 2011). Moreover pigs have a large litter size which makes it a cost-effective large animal model. With reference to SMA, permeable blood-brain barrier and the presence of tail necrosis and heart defects complicates interpretation of results in rodent models (Duque et al., 2015). Thus study of a large SMA model is necessary to understand SMA and translate an effective therapy.

Pigs, like other animals, possess one copy of SMN. Porcine SMN is 90% identical to humans, compared to 82% of murine Smn and is ubiquitously expressed (Lorson et al.,

2008). An SMA pig model was achieved by shRNA-mediated knock-down of porcine

SMN (Duque et al., 2015). The effective delivery of SMN:shRNA was mediated by self- complimentary AAV9 (scAAV9) injected at post-natal 5 in the cisterna magna (Duque et al., 2015). The virus transduced 78% of motor neurons in the spinal cord that led to a

73% reduction in SMN mRNA levels in the motor neurons. By 3-4 weeks post-injection, the pigs displayed SMA-like weakness, in particular hind-limb weakness, abnormal gait

37 and difficulty standing for prolonged periods (Duque et al., 2015). Electrophysiological tests indicated fibrillations, and decrease in CMAP and MUNE (Duque et al., 2015).

Interestingly, longitudinal studies revealed recovery of CMAP with no improvement in

MUNE, showing that compensation by reinnervation (Duque et al., 2015). The SMA pigs were used to test efficacy and timing of gene therapy and the results are described in section 1.5.3.

1.5 Therapy for Spinal Muscular Atrophy (SMA)

1.5.1 Increasing production of full-length SMN from SMN2

With the discovery that milder SMA patients have more copies of SMN2 and that

SMN2 produces small amount of full-length SMN (FL-SMN), it was postulated that upregulation of SMN2 could be a potential therapy (McAndrew et al., 1997). As mentioned in section 1.3.2, the critical nucleotide change alters an exon splice silencer

(ESS) or an exon splice enhancer (ESE) such that majority of SMN2’s transcripts fail to incorporate exon 7 (Burghes and McGovern, 2010). Apart from the CT change in exon

7 of SMN2, numerous elements in exon 7 and the flanking introns affect splicing. These

ESSs, ESEs, ISSs ( splice silencers) and ISEs (intron splice enhancers) have been extensively studied (Bebee et al., 2010; Burghes and McGovern, 2010; Singh et al.,

2006). A negative element named ISS-N1 was identified in intron 7 whose deletion significantly enhanced incorporation of exon 7 in SMN2 minigenes (Singh et al., 2006).

This was further validated by the use of antisense oligonucleotides to block ISS-N1 in

38 cell culture which led to increased production of full-length SMN protein (Singh et al.,

2006). Anti-sense oligonucleotides (ASOs) with modified backbone chemistry can be used to switch the splicing of an mRNA. Morpholino oligomers (MOs) have the advantage of low toxicity, stability and wide distribution properties (Porensky et al.,

2012). Delivery of the MO via intra-cerebro ventricular (ICV) injection at P0 caused up to 5-fold increase in FL-SMN production in the brain and spinal cord (Porensky et al.,

2012). Peripheral delivery at neonatal stages in mice also causes entry of ASO into the

CNS due to the relative permeability of the blood-brain barrier. A single dose of MO against ISS-N1 element drastically increased the lifespan of the SMNΔ7 SMA mice from

14 days to over 100 days (Porensky et al., 2012). This was in concert with phenotypic improvement of weight, righting ability and grip strength. However, the mice showed necrosis of tail and pinna. This observation is consistent in all rescue SMA mice; mice with increased SMN2 dosage or with increase on only neuronal SMN expression, to name a few. The relevance of necrosis is open to interpretation. Since SMA is a disease of decrease in SMN, one possibility is that necrosis might be due to insufficient upregulation of SMN in the autonomic nervous system or the circulatory system.

Importantly, delayed delivery of MO at PND04 or PND06 led to only a mild improvement in weight and survival (Arnold et al., 2015; Porensky et al., 2012). This indicates the existence of a limited time window and that a therapy would have maximum benefit if administered before it is too late. As promising therapies develop in SMA, there is a need to characterize a clinical biomarker suggesting the progression of disease and the efficacy of treatment. Compound motor action potential (CMAP), motor unit number

39 estimate (MUNE) and electrical impedance myography (EIM) have been explored as potential biomarkers in pre- and post-symptomatic SMN restoration in SMNΔ7 SMA mice. Pre-symptomatic ICV injection of ISS-N1 in SMA mice at PND02 resulted in complete recovery of CMAP, MUNE and EIM parameters while delayed treatment at

PND04 and PND06 did not improve the either of the parameters (Arnold et al., 2015).

Additionally, longitudinal studies with late treatment at PND04 or 06 showed recovery of

CMAP and not MUNE, implying sprouting by the existing rescued motor neurons

(Arnold et al., 2015).

ASOs with 2’-O-methyl chemistry (MOE) targeted against ISS-N1 region also improved the muscle physiology, motor function and survival of SMNΔ7 SMA mice via

ICV route upon administration at P0 (Passini et al., 2010). Yet the improvement in survival was to a median of 26 days and the MOEs decayed to basal levels by 30 days compared to sustained FL-SMN expression at 65 days with MO chemistry. Also, MOE chemistry has toxic effects at high doses while MO seemed safer at even higher doses.

Another study explored peripheral delivery of MOE against ISS-N1 in a different SMA mouse model, the severe Taiwanese model by Hsieh-Li, 2000 with 2 SMN2 copies (Hua et al., 2011). Multiple peripheral MOE doses increased survival from 10 days to a median of 137 days, with increased exon 7 inclusion in CNS and peripheral tissues (Hua et al.,

2011). It is however important to consider that in SMA infants the blood-brain barrier might not be permeable to peripheral ASOs at the time of treatment and this might diminish the effect.

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The aforementioned MOE-ASO targeted against ISS-N1 element, labelled ISIS-

SMNRx, is presently in a double-blind placebo-controlled Phase III clinical trial by ISIS

Pharmaceuticals, in collaboration with Biogen. SMA children of ages 2-15 are being enrolled in the trial and are administered the ASOs intrathecally. [Reference: https://clinicaltrials.gov/ct2/show/NCT02193074]

1.5.2 SMN Gene therapy

The presence of the blood-brain barrier (BBB) prevents crossing-over of drugs making therapy for CNS disorders very difficult. Using viruses as vehicles would be relatively easy provided the virus achieves efficient transduction in the brain and spinal cord. It was found that the serotype scAAV9 can efficiently deliver genes into the CNS

(Duque et al., 2009; Foust et al., 2009). A single intravenous injection of self- complementary AAV9 (scAAV9) in neonatal mice, with a relatively permeable BBB, transduced 56% of lumbar motor neurons (Foust et al., 2009). Along with spinal motor neurons, scAAV9 transduced neurons in the brain, dorsal root ganglion astrocytes in brain and spinal cord with occasional microglia and was sustained for over 5 months

(Duque et al., 2009; Foust et al., 2009). Transduction was observed in cardiac and skeletal muscle as expected. However, in adult mice, an intact BBB restricted the transduction of scAAV9 to mainly astrocytes and less of neurons (Foust et al., 2009).

Interestingly, the potential of scAAV9 to enter and transduce CNS has been validated in domestic cats (Felis catus), pigs (Sus scrofa) as well as non-human primates

(cynomolgus macaques, Macaca fascicularis) (Bevan et al., 2011; Duque et al., 2009;

Duque et al., 2015; Foust et al., 2010). Although the BBB is considered to be fully 41 developed in neonate kittens, 34% of lumbar motor neurons were transduced by scAAV9 upon intravenous delivery; albeit with a decrease in efficiency in adult cats (Duque et al.,

2009). To further test the window of opportunity for injection in non-human primates, scAAV9 was injected in macaques at post-natal days 1, 30, 90 and 3 years of age (Bevan et al., 2011). Similar results were obtained with extensive transduction of neurons and glial cells in brain and spinal cord, in addition to peripheral organs (Bevan et al., 2011).

Additionally it was shown that similar levels of motor neuron-transduction can be achieved at 1/30th the dose if scAAV9 is delivered directly into the CNS as opposed to a vascular injection (Meyer et al., 2015). Intrathecal delivery also decreased the transduction in peripheral organs. The study interestingly showed that the low doses of virus, combined with tilting the animal during transfusion, increases the efficiency of transduction of lumbar motor neurons in adult primates by about 80% (Meyer et al.,

2015).

The discovery of AAV9’s potential opened up avenue for therapy in CNS disorders. Gene therapy for the treatment of SMA was explored by intravenous injection of scAAV9-SMN in SMNΔ7 SMA mice. A one-time injection on post-natal day 1 increased the survival of SMA mice from 14 to over 250 days (Foust et al., 2010). This was accompanied by improved locomotor and righting abilities, neuromuscular transmission and weight gain (Foust et al., 2010). Later studies have shown that to obtain substantial rescue in survival of SMNΔ7 SMA mice about 20-40% MNs should be transduced (Meyer et al., 2015). Delay in gene therapy to SMA mice resulted in modest improvement with no change at all upon injection after P10 (Foust et al., 2010). These

42 results are in line with the oligonucleotide therapy studies, highlighting the existence of a limited time window available for improvement. The efficacy of SMN gene therapy was demonstrated in a large animal model of SMA pigs (Duque et al., 2015). scAAV9-SMN was delivered intracisternally at two time points – presymptomatic and upon onset of symptoms. Administration of therapy within a day of the shRNA-SMN knockdown vector in the pre-symptomatic phase resulted in SMA pigs not developing any weakness

(Duque et al., 2015). Electrophysiological output was preserved too. Treatment upon onset of symptoms (around PND24-24) remarkably ameliorated the SMA phenotype with none of the pigs reaching to a point of severe weakness or paralysis (Duque et al., 2015).

On the front of neuromuscular function, CMAP showed considerable improvement while

MUNE increased partially as compared to SMA pigs (Duque et al., 2015). These data further stress the importance of timely intervention for best therapeutic improvement.

A giant leap in the world of SMA was achieved on the 13th of May, 2014, with the first SMA baby getting intravascular infusion of scAAV9-SMN. The single site, phase

I/II trial for SMA type I babies is being conducted at The Nationwide Childrens Hospital,

Columbus Ohio, licensed to AveXis Inc., Dallas. Two cohorts of SMA patients have been injected with low and intermediate viral dose [Reference: http://avexis.com/latest-news/].

It is unlikely that the succumbed motor neurons can be revived, hence prompt diagnosis and treatment is critical to achieve maximum benefit.

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1.5.3 Additional strategies

Additional strategies to improve the SMA phenotype include small molecules that modulate SMN2 splicing, neuro-protective agents and molecules that help muscle mass.

Some SMN2 splicing modifiers identified from a large screen have been tested in SMA mice and appear promising. One of the compounds, called NVS-SM1, dramatically increased exon 7 inclusion in SMN2 transcripts in brain and spinal cord of mice upon oral dosing (Palacino et al., 2015). Daily oral dosing of NVS-SM1 for a month improved the weight of SMNΔ7 SMA mice and more than half the mice survived normally (Palacino et al., 2015). The splice modulating activity of the compound has been attributed to it stabilizing the U1snRNP−pre-mRNA complex of SMN2’s transcripts and thus ensuring correct splicing (Palacino et al., 2015). Another compound, SMN-C3 could traverse the

BBB in adult mice and increased FL-SMN mRNA in the brain and skeletal muscle

(Naryshkin et al., 2014). Daily oral dosing of SMN-C3 increased FL-SMN protein levels by 50-70% in the mild SmnC/C SMA model (Naryshkin et al., 2014). In SMNΔ7 SMA mouse model, daily intraperitoneal injection of the compounds at high doses extended survival to beyond 150 days (Naryshkin et al., 2014). One of the compounds from the study, RG7800, is being tested on Type I-III SMA patients by PTC and Roche

Pharmaceuticals [Cure SMA Conference 2015]. Novartis is conducting an open-label clinical study in Europe in Type I SMA infants to test the efficacy of LMI070, an orally dosed compound with the potential to increase FL-SMN and improve motor neuron condition in SMA [Cure SMA Conference 2015].

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A neuro-protective approach to SMA treatment is being explored by Roche

Pharmaceuticals with their compound Olesoxime [Cure SMA Conference 2015]. In a double blind placebo-controlled European trial, olesoxime delayed the onset of motor function for 2 years in Type II and Type III SMA patients [Families of SMA Conference

2014]. Other approaches to SMA management include increasing the muscle response to the available neuronal input. A fast-skeletal activator, CK-2127107, has passed

Phase I level of safety in humans. The study by Cytokinetics/Astellas aims to enhance the muscle performance in patients with motor neuron degenerative diseases like SMA [Cure

SMA Conference 2015].

[Reference: http://www.curesma.org/news/updates-from-drug-programs.html]

1.6 Significance of study

Dr. Victor Dubowitz, the pundit on childhood neurological disorders, in his study of SMA muscle histology noticed that in spite of atrophy, the SMA muscles have a normal distribution of fibers responsive to enzymatic histochemistry (Dubowitz, 1966).

Moreover, earlier work shows that to have a normal enzymatic differentiation, the muscle should have reached a gestation of atleast 26 weeks (Dubowitz, 1966). In Dr. Dubowitz’s words, “This suggests that the very small fibers (in SMA) are the result of fully, differentiated mature muscle, and not an arrest in the development of embryonic muscle.”

The implication is that the atrophic process in SMA should then have started after 26 weeks of fetal development.

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With the discovery of the SMN gene and that a ubiquitous decrease of SMN is responsible for causing SMA, the debate arose as to whether the increased requirement of

SMN is in neurons or neurons plus muscle. The muscle itself being developmentally defective has been implicated in SMA. As promising therapies for SMA are moving forward, this is an important question to address. Studies on mouse and pig models of

SMA have established the ‘when’ factor for therapy, i.e. when should therapy be given to obtain maximum benefit – the earlier the better. Once the neurons in SMA are degenerated, they are lost forever. The present dissertation addresses the ‘where’ aspect for SMA therapy – where is the increased requirement for SMN? Because SMA is a disease of low SMN and not complete absence of SMN, a good way to approach the above question is by decreasing and not eliminating SMN in a particular tissue. Previous studies in SMA have either (i) completely eliminated SMN from the neurons or muscle or

(ii) only decreased SMN in a tissue or (iii) only replaced SMN in neurons or muscle. We chose to titer SMN levels down to 2 copies of SMN2 (and SMN∆7) in either muscle or neurons and mainly not just decreased SMN in neurons or muscle but also restored SMN in neurons or muscle. This project is the first instance of performing the study both ways

– decreasing SMN and conversely replacing SMN in neurons or muscle. Performing the experiment both ways is necessary to interpret the results in its entirety. Chapter 4 deals with decreasing and restoring SMN in neuronal tissue and Chapter 3 is the study in skeletal muscle. Muscle atrophy is the first presentation in SMA and various methods have been explored to ameliorate the atrophy or enhance muscle performance. One attractive direction is to block the ubiquitin ligases that degrade muscle proteins and thus

46 block atrophy. In Chapter 2, we deleted the two main muscle-specific ubiquitin ligases in SMA and found no improvement whatsoever. An over-arching conclusion of the entire dissertation is that the motor neurons are most sensitive to SMN decrease and should be targeted for maximum therapeutic benefit. Secondly, the muscle tissue under normal physiological conditions develops and functions well with low levels of SMN.

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CHAPTER 2

Deletion of atrophy enhancing genes fails to ameliorate the phenotype in a mouse

model of spinal muscular atrophy

2.1 INTRODUCTION

Spinal muscular atrophy (SMA) is due to decrease in levels of a ubiquitous protein, Survival Motor Neuron (SMN) (Burghes and Beattie, 2009). Though the decrease of SMN occurs everywhere in SMA, it causes motor neuron degeneration and muscle atrophy. Moreover, in SMA patients, the loss of muscle strength is the first phenotypic presentation of the disease. Thus it has been proposed that blocking the muscle atrophy pathway could ameliorate the symptoms of muscle weakness in SMA patients (Engvall and Wewer, 2003). There have been clinical trials with creatine, phenylbutryate, gabapentin and thyrotropin releasing hormone targeting the muscle for treatment of type II and III SMA patients. An analysis of the efficacy of the aforementioned drugs in clinical trials showed no significant effect on the disease course

(Bosboom et al., 2009). Delivery of follistatin, which targets the negative regulators of muscle growth, to SMA mice improved the muscle mass in several muscle groups with a minimal effect on mean survival and no increase in maximium survival (Rose et al.,

48

2009). Moreover, transgenic over expression of follistatin showed little increase in muscle mass but no improvement in motor function or survival in the SMA mouse model

(Sumner et al., 2009). To date the most effective therapies in mice increase SMN expression in the nervous system by (i) reintroducing SMN with viral vectors or (ii) introducing antisense oligonucleotides that block negative regulators of splicing

(Dominguez et al., 2011; Foust et al., 2010; Passini et al., 2011; Porensky et al., 2012).

Increasing SMN levels in the mice results in rescue of both muscle weakness and survival

(Passini et al., 2011). Previous studies have shown that early introduction of any SMN- inducing therapy is needed for maximum effect on survival and phenotypic improvement in mice (Foust et al., 2010; Le et al., 2011; Lutz et al., 2011).

Muscle Atrophy F-box, MAFbx (also called Atrogin1), and Muscle RING Finger

1, MuRF1 (also called Trim63), are two muscle specific E3 ubiquitin ligases that are required for muscle atrophy (Bodine et al., 2001). The E3 Ubiquitin ligases recognize the target proteins (sarcomeric, contractile, signaling, metabolic and transcriptional proteins in the case of muscle) and transfer the ubiquitin moiety from E2 enzyme onto the target

(Perera et al., 2011). The ubiquitin-tagged proteins get sent to the ubiquitin-proteasome system (UPS). The UPS degrades the muscle proteins, thus maintaining their regular turnover and in turn the muscle mass. Upon receiving a signal for atrophy, the ubiquitin ligases are upregulated causing increased breakdown of muscle proteins, tipping the balance towards decrease in muscle mass (Glass, 2005, 2010; Lecker et al., 1999).

MAFbx also down-regulates protein synthesis in muscles (Glass, 2005). The known substrates of MAFbx are MyoD, calcineurin and translation initiation factor 3-f (eIF3-f)

49

(Lagirand-Cantaloube et al., 2008; Li et al., 2004; Tintignac et al., 2005). A second muscle , MuRF1 targets light-chain, MyLC1 and MyLC2, myosin heavy chain (MyHC), myosin-binding protein-C (MyBP-C) and cardiac

(Cohen et al., 2009; Kedar et al., 2004). MuRF1 might have a role in post-transcriptional modification, titin turn over and metabolic regulation (Hirner et al., 2008; McElhinny et al., 2002). Homozygous deletion of either MAFbx or MuRF1 results in sparing of muscle mass in mice subjected to atrophy by denervation (Bodine et al., 2001). The beneficial effect of deletion of the muscle ubiquitin ligases was reflected in increased muscle weight, and maintenance of mean fiber size and fiber size variability (Bodine et al.,

2001). Thus, deletion of MAFbx or MuRF1 has been shown to protect against muscle atrophy in mice.

We proposed that deletion of the ubiquitin ligases in the SMNΔ7 mouse model of

SMA could result in muscle sparing and the prevention of atrophy. By ameliorating atrophy in the SMA mouse, we predict that both weight and survival would increase.

Using MAFbx-/- or MuRF1-/- transgenic mice we deleted the ubiquitin ligases in the

Δ7SMA mouse. We found that loss of MAFbx in the SMNΔ7 mouse resulted in no improvement in either weight or survival of SMA mice, although there was a minimal increase in muscle fiber size. Furthermore, deletion of MuRF1 in the SMNΔ7 mouse not only failed to improve survival, the mice in fact died earlier in the absence of MuRF1. It has been suggested that HDAC inhibitors act to benefit SMA mice by inhibition of the upregulation of MAFbx and MuRF1 (Bricceno et al., 2012). We thus investigated the expression of MAFbx, MuRF1, and the other muscle-specific ubiquitin ligases with

50 known expression at embryonic and postnatal stages, namely MuRF2 and MuRF3, in both normal and SMA mice. In summary, we found that the expression of MAFbx and

MuRF1 increases at post-natal day 14 in SMA mice, while there is no significant difference in the levels of MuRF2 and MuRF3 between SMA and control mice. It appears unlikely that HDAC inhibitors act by blocking the upregulation of ubiquitin ligases given that deletion of MAFbx or MuRF1 did not improve survival in SMA mice.

2.2 MATERIALS AND METHODS

2.2.1 Mouse strains and breeding

The breeding and maintenance of mice was in accordance to The Institutional Animal

Care and Use Committee (IACUC) regulations of the Ohio State University. The

MAFbx-/- and MuRF1-/- mice used for the experiments have been described previously

(Bodine 2001). Briefly, the genomic DNA of MAFbx and MuRF1 were disrupted with a

LacZ/neomycin cassette through the start codon (Bodine et al., 2001). The MAFbx-/- and

MuRF1-/- mice were phenotypically normal and fertile.

MAFbx-/- and MuRF1-/- breeders were crossed to the SMNΔ7 SMA carrier mice

(Smn+/-, SMN2+/+, SMNΔ7+/+) (Le et al., 2005). F1 mice carrying the Smn KO allele were interbred to homozygosity for SMN2, SMNΔ7 and the MAFbx/MuRF1 KO allele. Mouse genotypes used for the study were as follows. MAFbx-/--SMA: MAFbx-/-, Smn-/-, SMN2+/+,

SMNΔ7+/+; MuRF1-/--SMA: MuRF1-/-, Smn-/-, SMN2+/+, SMNΔ7+/+; SMA: MAFbx+/+,

MuRF1+/+, Smn-/-, SMN2+/+, SMNΔ7+/+; and MAFbx-/-- Smn+/-: MAFbx-/-, Smn+/-, SMN2+/+, 51

SMNΔ7+/+; MuRF1-/-- Smn+/-: MuRF1-/-,Smn+/-, SMN2+/+, SMNΔ7+/+; Smn+/- Control:

MAFbx+/+, MuRF1+/+, Smn+/-, SMN2+/+, SMNΔ7+/+ and Smn+/+ Control: MAFbx+/+,

MuRF1+/+, Smn+/+, SMN2+/+, SMNΔ7+/+ as controls.

2.2.2 Genotyping and weighing

Genomic DNA was isolated for tail clips and PCR amplified using the following primers

- MAFbx KO: 5’CTTCCTCGTGCTTTACGGTATC and

5’AGCACAGATATGGTACCTTCC; MAFbx WT:

5’CTGCAACAAGGAGGTATACAGT and 5’CATGCAGGTGTACATGCAAGTAG;

MuRF1 KO: 5’TGGCTACCCGTGATATTGCTG and

5’CGTTCGAGGGTTAAGAAAGTCTAG; MuRF1 WT:

5’CGTTCGAGGGTTAAGAAAGTCTAG and 5’GCACTCCTGCTTGTAGATGTC.

The PCR conditions were 95 ºC for 5 min, followed by 95 ºC for 1 min, 57 ºC for 1 min,

72 ºC for 1 min, for a total of 35 cycles. The genotyping of SMA mice has been described previously (Le et al., 2005). Pups were weighed from the day of birth (PND01) until weaning (PND21).

2.2.3 Muscle fiber analysis

The gastrocnemius muscle of PND08 pups was flash frozen in liquid-nitrogen cooled isopentane and cryostat sectioned (IEC Minotome Plus, MN). 14 µm thin transverse sections were blotted on Superfrost Plus slides (Fischer Scientific), air dried, and stained with hematoxylin and eosin (H&E) as follows. Sections were fixed sequentially in 50%,

70%, 90% and 100% ethanol, rinsed in tap water (3x) and stained in Harris’ Hematoxylin

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(Sigma) for 30 s. The slides were then washed in tap water (3x), stained in Eosin (Sigma) for 20 s and rinsed in tap water (3x). This was followed by a step-wise dehydration in

50% (4x), 70% (4x), 90% (7x) and 100% (10x) ethanol, and clearing in Histoclear

(Sigma) twice (10x each). The slides were mounted with Permount (Sigma).

The myofiber cross-sections were viewed with a Nikon Eclipse 800 microscope

(Nikon Corporation, Japan) and imaged using a Nikon FDX-35 digital camera. The cross- sectional area was measured using SPOT Advanced software (v3.5.9, Diagnostic

Instruments, Inc., MI).

2.2.4 Statistical Analyses

Survival (Kaplan-Meier curve) and statistical analyses (Mann-Whitney Rank Sum Test,

Shapiro-Wilk Normality Test and Equal variance test) were performed using SigmaPlot v11 (Systat Software Inc., CA).

2.2.5 Droplet digital PCR (ddPCR)

Fresh tissue was flash frozen in liquid nitrogen. RNA was isolated from Trizol

(Invitrogen) homogenized tissue and purified using the RNeasy kit (Qiagen). 2.0 µg of

RNA was used for the RT-PCR reaction performed using AMV reverse transcriptase

(Affymetrix/USB). The quantification of transcripts was done using droplet digital PCR

(Bio-Rad). Approximately ten to fourteen thousand droplets, containing the template with the primers and probe, were generated. The measured fluorescence after PCR amplification was used to calculate the concentration of a transcript using Poisson statistics by the QuantaSoft software (Bio-Rad) (Porensky et al., 2012). Relative levels of

53 a transcript were determined with reference to cyclophilin expression. The sequences of the primers and probes were as follows. MAFbx: 5’TCCTTATGCACACTGGTGCA,

5’CTCAGCCTCTGCATGATGTTC, Probe-5’FAM-CAACATTAACATGTGGGTGT-

MGB; MuRF1: 5’AGCTGAGTAACTGCATCTCCATGC,

5’TTCTGCTCCAGGATGGCGTA, Probe-5’FAM-CGAGTGCAGACGATCA-MGB;

MuRF3: 5’CACTTGGAGGGCTCCTCAAAG, 5’AGAGCCTTGCTCCATGCTCTC,

Probe-5’FAM-TGTCGAAGGTGGAGCTG-MGB; Cyclophilin:

5’GTCAACCCCACCGTGTTCTT, 5’TTGGAACTTTGTCTGCAAACA, Probe-5’VIC-

CTTGGGCCGCGTCT-MGB. For MuRF2 isomers p50A and p60A, a common reverse primer in exon10 was used; 5’AGAAGGGGCCTCAAATCCAATC and the forward primers and probes were p50A 5’GAAAGCTGCAGAGCCCTCTCAG, p60A

5’TAGGGCCTCTGGGCATTGAG; p50A Probe-5’FAM-TCTCCAGAACCGTTTT-

MGB, p60A Probe-5’FAM-CAGTGAGTGGTAAGGAGTC-MGB.

2.3 RESULTS

2.3.1 Generation of the transgenic mouse lines

The SMNΔ7 SMA model mouse has a homozygous deletion of mouse Smn, and carries two copies of human SMN2 transgene and the human cDNA expressing SMN lacking exon 7 (Smn-/-; SMN2+/+; SMNΔ7+/+) (Le et al., 2005). The phenotypically normal

Smn+/- Control contains one copy of the Smn gene and is homozygous for SMN2 and

SMNΔ7. The Smn+/- mice were crossed to the MAFbx and MuRF1 homozygous KO mice

54 to obtain MAFbx-/- and MuRF1-/- in the SMA background. The Smn+/- mice with homozygous KO of MAFbx or MuRF1 (genotype: MAFbx-/- or MuRF1-/-; Smn+/-;

SMN2+/+; SMNΔ7+/+) appear phenotypically normal. They are referred to as MAFbx-/--

Smn+/- or MuRF1-/-- Smn+/- in this study. Thus, deletion of MAFbx and MuRF1 does not have any phenotypic effect in the absence of an atrophy signal. To obtain the affected

SMA mice, the aforementioned mice were interbred to get MAFbx-/-- SMA or MuRF1-/--

SMA (genotype: MAFbx-/- or MuRF1-/-; Smn-/-; SMN2+/+; SMNΔ7+/+). Mice with two copies of wild type Smn, SMN2 and SMNΔ7 transgenes (Smn+/+; SMN2+/+; SMNΔ7+/+) are referred to as Smn+/+ Control.

2.3.2 Weight and survival analyses

To study if deletion of the muscle ubiquitin ligases namely MAFbx and MuRF1 results in muscle sparing in a model of SMA, the body weight was measured daily from the day of birth to weaning at PND21 (post-natal day 21). Upon homozygous deletion of

MAFbx in SMA animals (n=17), the pattern of weight gain and subsequent loss was similar to that seen in SMA pups which retain the WT copies of MAFbx (n=11) (Figure

2.1A). Thus, there was no improvement in body weight in MAFbx-/--SMA mice as compared to SMA mice. Homozygous knock-out of the second muscle ubiquitin ligase,

MuRF1 in SMA, worsened the survival. Only one mouse survived till PND12 (Figure

2.1B).

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Figure 2.1. Weight and survival analyses of MAFbx-/--SMA and MuRF1-/--SMA animals: (A) MAFbx-/--SMA (red) mice showed a pattern of weight gain similar to SMA (green) until PND03. (B) MuRF1-/--SMA (blue) mice showed early death and one mouse survived till PND12. The weights of MAFbx-/--Smn+/- and MuRF1-/--Smn+/- controls (gray) were similar to the Smn+/- control (black). (C) The mean survival time of MuRF1-/- -SMA (blue) was 2.6±0.8 days (n=15), MAFbx-/--SMA (red) was 14.4±0.4 days (n=17) compared to SMA animals (n=11) (green) with 17.0±0.8 days. Thus, deletion of neither MAFbx nor MuRF1 improved the survival in SMA. MAFbx-/--Smn+/- and MuRF1-/-- Smn+/- (gray) and the Smn+/- control (black) survived for >21 days (n=15). (Log-Rank P = <0.001) (error bars = SEM)

Furthermore, the deletion of MAFbx and MuRF1 in SMA background did not improve the survival of the mice. There was no improvement in the mean survival time of

MAFbx-/--SMA (14.4 ± 0.4 days, n=17) compared to SMA animals (17.0 ± 0.8 days, n=11) (Figure 2.1C). Upon the deletion of MuRF1 in SMA, there was a drastic drop in the mean survival time (2.6 ± 0.8 days, n=15) (Figure 2.1C). Only one MuRF1-/--SMA mouse survived till PND12. The controls, MAFbx-/-- Smn+/-, MuRF1-/-- Smn+/- and Smn+/-

Control survived beyond 21 days (n=15 for each). Thus, deletion of MAFbx in the SMA background failed to improve the survival of the mice and deletion of MuRF1 significantly decreased survival of the mice.

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2.3.3 Analyses of muscle morphology

If the muscle ubiquitin ligases ameliorate the SMA phenotype, it should be reflected in improved muscle fiber size. Hence, we studied if muscle fiber size was preserved upon deletion of the ubiquitin ligases in an SMA background. The muscle morphology was examined by H&E staining of the gastrocnemius muscle of MAFbx-/--

SMA (Figure 2.2A) and SMA (Figure 2.2B) mice at PND08. The mean fiber size distribution was measured for each group (n=2070 fibers per group) (Figure 2.2C). The muscle fiber size in MAFbx-/-–SMA mice (mean 315 ± 68 µm2, median 225 µm2) was determined to be greater than the fiber size in SMA mice (mean 211 ± 1 µm2, median 204

µm2). The fiber size distribution in MAFbx-/-–SMA reveals a significant increase in large fibers as compared to SMA (P<0.001, Mann-Whitney Rank Sum Test). MAFbx-/-- Smn+/- and and Smn+/- Control had mean fiber sizes of 323 ± 2 µm2 (median 312 µm2) and 272 ±

2 µm2 (median 258 µm2) respectively. The increase in number of large fibers upon deletion of MAFbx in SMA would be expected as the loss of MAFbx will prevent severe atrophy. It should however be noted that median fiber size of MAFbx-/-–SMA is still less than the median fiber size in Smn+/- Control (204 µm2 vs 258 µm2 respectively). Thus while deletion of MAFbx in the SMA background does not improve total body weight and survival, it does result in an increase in gastrocnemius fiber size in SMA animals.

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Figure 2.2. Gastrocnemius muscle fiber of PND08 (A) MAFbx-/-–SMA (B) SMA pups (at 20x magnification) after H&E staining and (C) their corresponding muscle fiber area frequency distributions. MAFbx-/-–SMA: Mean fiber size - 315±68 µm2, median size - 225 µm2 which was higher than that of SMA (mean fiber size 211±1 µm2,median 204 µm2). The difference in the median values between SMA and MAFbx-/-–SMA was significant (Mann-Whitney Rank Sum Test, P<0.001). The fiber size distribution of both SMA and MAFbx-/-–SMA vary significantly from a normal distribution (Shapiro-Wilk Normality Test, P<0.001). For the other controls: MAFbx-/--Smn+/-: mean fiber size - 323±2 µm2, median size - 312 µm2 while Smn+/- Control: mean fiber size - 272±2 µm2 and median - 258 µm2. (n=2070 for each group) 58

2.3.4 Expression of the muscle ubiquitin ligases in SMA mice

For MAFbx and MuRF1 to improve the muscle morphology in SMA, it is important to investigate if, firstly, they are expressed in neonatal mice and secondly, if they are upregulated in SMA. Previous studies have shown that in the skeletal muscle,

MuRF1 is present embryonically but is upregulated only postnatally, yet deletion of

MAFbx and MuRF1 have been studied only in adult models of induced atrophy (Glass,

2005; Perera et al., 2012). Hence I quantified the expression of MAFbx and MuRF1 in

SMA and control mice. Two other members of the muscle-specific tripartite-motif

(TRIM) family of ubiquitin ligases are MuRF2 and MuRF3. Of the two isoforms of

MuRF2 with known timing of expression, p50A and p60A, p50A dominates in embryonic stages with a switch to the p60A isoform postnatally (Perera et al., 2011;

Perera et al., 2012). MuRF1 and MuRF3 are expressed only postnatally. I performed droplet digital PCR (ddPCR, BioRad) to determine if the ubiquitin ligases are upregulated in the skeletal muscle of severe SMNΔ7 SMA mice from PND02 to PND14.

Finally, because studies in SMA have shown early heart failure, arrhythmia and cardiac defects (Bevan et al., 2010; Heier et al., 2010; Shababi et al., 2010), the levels of the ubiquitin ligases in the cardiac muscle were examined as well.

The expression of MAFbx, MuRF1, MuRF2p50A, MuRF2p60A and MuRF3 were quantified in SMA and Smn+/+ Control (Smn+/+, SMN2+/+, SMNΔ7+/+) mice at PND02,

PND05, PND08 and PND14 by ddPCR (Figures 2.3 and 2.4). Cyclophilin expression was used as an internal control to calculate the relative fluorescence units (RFU). In the skeletal muscle, I found low expression of both MAFbx and MuRF1 levels between SMA

59 and Smn+/+ Control mice at time points before PND14 (Figure 2.3A). At PND14, MAFbx and MuRF1 expression in the SMA background increase dramatically as compared to

Smn+/+ Control: MAFbx increased to 11.8 fold (mean RFU in SMA - 1.04±0.28, mean

RFU in Smn+/+ Control - 0.08±0.02, *P=0.04) and MuRF1 increased to 3.9 fold (mean

RFU in SMA - 1.8±0.18, mean RFU in Smn+/+ Control - 0.46±0.13, **P=0.01). Similarly, the levels of MAFbx and MuRF1 show an increase at PND14 in SMA in the cardiac tissue (Figure 2.3B). MAFbx increased to 2.7 fold and MuRF1 increased to 1.97 fold of

Smn+/+ Control’s expression, however the increase did not reach statistical significance

(MAFbx: mean RFU in SMA - 1.22±0.32, mean RFU in Smn+/+ Control - 0.45±0.16,

MuRF1: mean RFU in SMA - 1.34±0.26, mean RFU in Smn+/+ Control - 0.68±0.12).

Thus, the expression of MAFbx and MuRF1 are increased in the SMA model. Next, I measured the levels of MuRF2 and MuRF3 to determine if they increase postnatally in the SMA mouse. Figure 2.4 shows the expression of MuRF2p50A, MuRF2p60A and

MuRF3 in the skeletal and cardiac tissue in the SMA mouse. I found no significant difference in expression levels of the MuRF2 isoforms (Figure 2.4A and B) or MuRF3

(Figure 2.4C and D) in the SMA mouse in the skeletal and cardiac muscle at the time points examined. Thus MuRF2 and MuRF3 transcripts appear to be expressed normally in the early postnatal period (PND02-14) in the SMNΔ7 SMA mouse model.

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Figure 2.3. Quantification of MAFbx and MuRF1 transcripts (by ddPCR) at PND01, PND 05, PND08 and PND14 of Smn+/+ Control and SMA animals. (n=3 for each group at each time point) An increase in the mean relative fluorescence units (RFU) for MAFbx and MuRF1 was observed at PND14 in SMA. (A) MAFbx and MuRF1 transcripts in skeletal muscle: MAFbx in Smn+/+ Control and SMA were 0.08±0.02 and 1.04±0.28 respectively at PND14, while for MuRF1 transcripts the mean RFU for Smn+/+ Control was 0.46±0.13 SMA while that for SMA was 1.80±0.18 at PND14. (*P=0.04, **P=0.01) (B) MAFbx and MuRF1 transcripts in heart: For MAFbx, the mean RFU for Smn+/+ Control was 0.45±0.16 v/s 1.22±0.32 for SMA at PND14. For MuRF1, the mean RFU for Smn+/+ Control and SMA were respectively 0.68±0.12 and 1.34±0.26 at PND14. (error bars = SEM)

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Figure 2.4. Quantification of MuRF2p50A, MuRF2p60A and MuRF3 transcripts (by ddPCR) at PND01, PND 05, PND08 and PND14 of Smn+/+ Control and SMA animals. (n=3 for each group at each time point) There is no significant difference in MuRF2 and MuRF3 transcripts between control and SMA at any time point. MuRF2 transcripts in (A) skeletal muscle and (B) heart. MuRF3 transcripts in (C) skeletal muscle and (D) heart. (error bars = SEM)

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2.4 DISCUSSION

The genes MAFbx (Atrogin 1) and MuRF1 (Trim63) are upregulated upon a signal for atrophy of muscle (Bodine et al., 2001). Indeed, their upregulation has been observed in 13 distinct models of skeletal muscle injury (denervation, immobilization, hindlimb suspension, lipopolysaccharide injection, sepsis, glucocorticoid dexamethasone, cachectic cytokine interleukin-1 (IL-1) or nutritional deprivation) (Glass, 2005). While

KO of MAFbx spares the muscle against denervation and immobilization, the KO of

MuRF1 protects against denervation, unloading, glucocorticoids, acute lung injury as well as aging (Baehr et al., 2011; Baehr et al., 2014; Bodine et al., 2001; Files et al.,

2012; Gomes et al., 2012; Hwee et al., 2014; Labeit et al., 2010). In all the studies, adult mice as opposed to neonatal mice have been examined. Removal of either MAFbx

(Atrogin 1) or MuRF1 significantly blocks atrophy by denervation, as these E3 ligases are responsible for mediating muscle protein breakdown through the ubiquitin proteasome system (UPS). As the genes (MAFbx or MuRF1) have been shown to play a major role in atrophy, we investigated whether removal of these genes in spinal muscular atrophy reduced the atrophy of muscle. In the present study we found that deletion of

MuRF1 in SMA mice significantly decreased the survival whereas deletion of MAFbx had no impact on weight or survival. The loss of MAFbx in SMA significantly altered the fiber size distribution leading to an increase in number of large fibers. Furthermore, the fiber size decreases much before the marked increase in MAFbx and MuRF1 expression.

This indicates that an alternative pathway is used in SMA to obtain small fibers. While

MAFbx and MuRF1 play a role at a later stage (PND14), their role in early stages of

63 atrophy is unclear. To analyze the expression pattern of the muscle ubiquitin ligases in a neonatal SMA mouse, we quantified the transcripts of MAFbx, MuRF1, MuRF2 and

MuRF3 in skeletal and cardiac muscle. Our findings indicate an 11.8 fold increase in the levels of MAFbx and a 3.9 fold increase in the levels of MuRF1 in SMA in skeletal muscle at PND14. The levels of MuRF2 and MuRF3 are comparable in control and SMA animals. Thus MAFbx and MuRF1 are upregulated at PND14 yet we find no increase in weight or survival when these genes are deleted in the SMA mice. It could be that the increased expression is too late in the lifespan of the SMA mouse to have any effect. It is also possible that other muscle ubiquitin ligases, unknown as of now, might be responsible for atrophy in SMA.

Previous studies in the Δ7SMA mouse model have shown that the histone deacetylase (HDAC) inhibitor trichostatin A (TSA) improves body weight and survival of the animals (Avila et al., 2007; Bricceno et al., 2012). Furthermore the authors found a significant increase in myofiber number and size (Avila et al., 2007). TSA does increase

SMN expression but as an HDAC inhibitor, it will block HDAC4 activity which can change the expression of genes important for the development of atrophy. Elevation of

MuRF1 and MAFbx1 in both SMA mice and human muscle tissue from SMA patients was also reported (Bricceno et al., 2012). The authors thus proposed that TSA improves

SMA muscle pathology by inhibiting the muscle atrophy pathway via down regulation of the muscle ubiquitin ligases, and hence the suggestion that blocking the atrophy pathway may improve SMA. Interestingly the activation of MAFbx and MuRF1 occur relatively late (PND11) and well after the effective window for TSA to have a benefit. The first

64 dose of TSA was administered at PND05 (Bricceno et al., 2012). Moreover, there is a marked reduction in muscle fiber size in SMA animals prior to the high expression of

MAFbx and MuRF1 indicating that at least the early fiber size reduction is independent of these two genes. In the current paper we show that loss of either MAFbx or MuRF1 in the severe SMA mouse model does not improve the survival. The fiber size distribution in SMA mice with MAFbx deletion tends towards larger fibers, but there is no increase in total body weight or survival. Based on the present work, TSA’s beneficial role in SMA is due to a pathway other than that involving down regulation of MAFbx and MuRF1.

It has been reported that MAFbx-/- mice though phenotypically normal initially, develop cardiomyopathy later on and do not survive beyond 16-18 months of age (Zaglia et al., 2014). The death of MAFbx-/- mice was attributed to dysregulation of the autophagy pathway due to lack of turnover of key proteins by MAFbx, leading to cardiomyopathy and eventually heart failure (Zaglia et al., 2014). In our murine colonies however, we did not observe early death. Our MAFbx-/- breeders, with SMN2 and SMNΔ7 in the background, lived beyond 2 years of age and did not show any apparent phenotype. Our

Δ7SMA mice in FVB background were bred to MAFbx-/- in C57BL background to obtain desired breeders. The lack of cardiac phenotype in the MAFbx-/-; Smn+/-; SMN2+/+;

SMNΔ7+/+ mice in late adulthood may be due to mixing of backgrounds or increase in dosage of full-length SMN protein. Additionally, the drastic decrease in survival of

MuRF1-/--SMA mice remains to be investigated. Yeast two-hybrid studies have identified transcriptional regulators and translation factors, in addition to mitochondrial and sarcomeric proteins as targets of MuRF1 (Witt et al., 2008). Overexpression of MuRF1 in

65 the muscle has thrown light on its role in carbohydrate metabolism, causing increased insulin secretion and depletion of liver’s glycogen stores (Hirner et al., 2008). Also,

MuRF1 protects against multiple types of atrophy signals as described earlier. Thus, it has been suggested that MuRF1 might target a variety of regulatory proteins involved in key pathways in the muscle and heart cell (Baehr et al., 2014). Hence, it is probable that in addition to low levels of SMN protein, complete absence of MuRF1, in the MuRF1-/--

SMA mice may be affecting a critical pathway in development leading to death of the mice in the neonatal stage.

In summary, though the molecular mechanisms of atrophy in the SMA mouse seem to involve MAFbx and MuRF1 at late stages (PND14), the genetic deletion of these ubiquitin ligases did not improve the weight or survival of the mice. Yet to date, unlike treatments targeted to the nervous system, no therapy directed solely towards blocking atrophy or increasing muscle function has significantly ameliorated the phenotype of the

Δ7SMA mouse.

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CHAPTER 3

Low levels of Survival Motor Neuron protein are sufficient for normal muscle

function in the SMN∆7 mouse model of SMA

3.1 INTRODUCTION

SMN protein functions in the assembly of snRNPs, by loading the Sm proteins onto the snRNA (Burghes and Beattie, 2009; Meister et al., 2001; Pellizzoni et al., 2002).

It has also been suggested to act as a master ribonucleoprotein assembler, assembling the

Sm-Lsm10/11 ring on the U7snRNA as well as other protein complexes on other RNAs

(Paushkin et al., 2002; Pillai et al., 2001; Schumperli and Pillai, 2004; Terns and Terns,

2001; Tisdale et al., 2013). Apart from the canonical snRNP assembly, SMN has only been demonstrated to be active in U7snRNP assembly while the other reactions are currently only suggested (Li et al., 2014). Since SMN is also found in the axons of neurons, it has been proposed that SMN could play a role in assembling the transport granules of axons (Gu et al., 2002; Rossoll et al., 2003; Rossoll et al., 2002). Although

SMN is expressed in all tissues, SMA particularly affects motor neurons and results in

67 atrophy of muscle. Our laboratory has previously shown that high expression of SMN in neurons with low levels in other tissues gives substantial rescue of severe SMA mice

(Gavrilina et al., 2008). In addition, it was shown that expression of very high levels of

SMN in skeletal muscle with no leakage into other organs had no impact on the SMA phenotype. However, with high expression of SMN in skeletal and cardiac muscle and a low level of SMN (above that produced by SMN2) in all tissues, the mice survived to 160 days (Gavrilina et al., 2008). If SMN is removed from muscle but retained in other tissues, then a severe muscular dystrophy results (Cifuentes-Diaz et al., 2001). However complete loss of SMN is not what occurs in SMA. A minimum amount of SMN is required in all cell types; complete removal of SMN leads to cell death (Cifuentes-Diaz et al., 2001; Frugier et al., 2000; Vitte et al., 2004). Since atrophy of muscle is a symptom of SMA, treatment of the muscle tissue has been explored. Transgenic expression of follistatin showed a minimal improvement in muscle mass with no improvement of motor function or survival (Sumner et al., 2009). Administration of follistatin, which blocks myostatin, a negative regulator of muscle mass, did not improve the survival of SMA mice (Rose et al., 2009); neither did transgenic inactivation of murine myostatin in SMA mice (Rindt et al., 2012). Moreover, homozygous knock out of muscle ubiquitin ligases,

MAFbx and MuRF1, did not improve the survival or muscle mass of SMA mice (Iyer et al., 2014).

A question that has not been addressed is how muscle would perform when completely reliant on two copies of SMN2 for its SMN protein requirement. Does SMN reduction, not removal, affect the ability of muscle to produce force? In the current

68 chapter, we address this question directly. To get a complete picture of the role of the muscle in SMA, we not only deleted Smn specifically in the muscle, but also replaced

Smn in the muscle. We chose to use the Myf5-Cre driver so as to remove mouse Smn from both myoblasts and myotubes in mice containing two copies of SMN2 and SMNΔ7 transgenes (Le et al., 2005; Tallquist et al., 2000). We found that decreasing SMN in muscle to SMA-levels does not hamper the muscle’s force production or fiber size or morphology. Moreover, the total body weight and survival of mice with decreased SMN in muscle remain unchanged. Conversely, replacement of SMN in muscle tissue of SMA mice with SMN depleted to SMA-levels elsewhere resulted in no improvement in survival or body weight of the SMA mice. Thus, we conclude that the muscle tissue per se can function in a completely normal manner when SMN is at reduced levels and that the muscle does not play a crucial role in SMA pathogenesis.

3.2 MATERIALS AND METHODS

3.2.1 Mouse breeding

The Cre lines and the floxed Smn lines were crossed to the SMNΔ7 SMA mouse model

(SmnWT/KO; SMN2+/+; SMNΔ7+/+ - Jackson No. #005025, FVB.Cg-Tg(SMN2)89Ahmb

Smn1tm1MsdTg(SMN2*delta7)4299Ahmb/J) (Le et al., 2005). Throughout the experiments, the Cre lines and floxed lines were maintained separately to eliminate germline recombination and crossed to obtain the affected mice: Cre+; SmnWT/KO;

SMN2+/+; SMNΔ7+/+ x SmnF7/F7; SMN2+/+; SMNΔ7+/+  Cre+; SmnD7/KO; SMN2+/+;

SMNΔ7+/+, where SmnD7 allele is deletion of Smn exon 7 post recombination, and Cre+;

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SmnWT/KO; SMN2+/+; SMNΔ7+/+ x SmnINV/WT; SMN2+/+; SMNΔ7+/+  Cre+; SmnRe/KO;

SMN2+/+; SMNΔ7+/+, where SmnRe allele is replacement of Smn exon 7 post recombination (Figure 3.1B). Littermates with Cre+; SmnD7/WT and Cre+; SmnRe/WT were used as controls. The mouse handling and procedures were in strict compliance with the

IACUC protocols and The Ohio State protocol #2008A0089. The Cre and floxed lines used in the study are described in Table 3.1 and 3.2 respectively. SmnINV and SmnRe alleles were sequenced (Figure 3.1). To visualize the motor neurons by immunohistochemistry, mice with GFP expression under the motor neuron specific HB9 promoter (Jackson No. 005029) were used (Arber et al., 1999). To identify Cre recombinant tissues, the reporter line tdTomato RFP (Jackson No. 007914) was used

(Madisen et al., 2010).

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Table 3.1 Cre drivers

Jax Strain Abbreviation Site of Cre Description References No. name used expression

Tg(Sox2- Expression of Cre 008- (Hayashi et Cre)1Amc Sox2-Cre Ubiquitous under mouse Sox2 454 al., 2002) /J promoter at e6.5 Expression of Cre (Tallquist 007- Myf5tm3(cre) Muscle under endogenous Myf5-Cre et al., 893 Sor/J precursors mouse Myf5 2000) promoter at e8

Table 3.2 Floxed alleles

Jax Recombination Result upon Cre Strain name References No. Before After recombination

006- Deletion of Smn (Frugier et Smn1tm1Jme/J SmnF7 SmnD7 138 exon 7 al., 2000)

007- Smn1tm3(SMN2/ Replacement of (Lutz et al., SmnINV SmnRe 249 Smn1)Mrph/J Smn exon 7 2011) Gt(ROSA)26 (Madisen 007- Sortm14(CAG- tdTom RFP expression et al., 914 tdTom tdTomato)Hze/J 2010)

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Figure 3.1. Sequence of SmnINV and SmnRe alleles. (A) SmnINV and (B) SmnRe alleles were PCR-amplified with primers designed around the 5’ and 3’ junctions and cloned into pCRTM 4-TOPO® vector (Invitrogen), transformed in E. coli DH5α strain, and sequenced.

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3.2.2 Genotyping, weighing and phenotypic assessment of mice

Genomic DNA was isolated from tail clips for genotyping. The primers used for PCR are described in Table 3.3. The PCR conditions and genotyping of SmnKO, SMN2 and

SMNΔ7 was as described in Le et al., 2005 (Le et al., 2005). The mice were weighed daily from day of birth, PND01, to weaning at PND21, and weekly thereafter. Phenotype was observed every day till weaning, and then weekly. Mice were sacrificed according to our IACUC approved protocol. Approximately equal number of males and females were studied in each genotypic grouping. The number of animals required for analysis was determined as described previously (Butchbach et al., 2007).

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Table 3.3 Primer sequences

Transgene/ Forward primer Reverse primer Allele

F7 AGA AGG AAA GTG CTC ACA TAC TGT CTA TAA TCC TCA TGC TAT Smn AAA TT GGA G

INV ACT TCT TAA TTT GTA TGT GAG CGC TTC ACA TTC CAG ATC TGT Smn CAC T CTG

Re ATT TAA GGA ATG TGA GCA CCT CGC TTC ACA TTC CAG ATC TGT Smn TCC CTG

Cre CCT GTT TTG CAC GTT CAC CG ATG CTT CTG TCC GTT TGC CG

TAG ACG CCT GAA GAA GGT CAA GAC TAT GGT AAA AGC GAG GCT Myf5-Cre C TAC

HB9:GFP GTC GAG CTG GAC GGC GAC GT CTG CAC GCT GCC GTC CTC GA

ACG CTG ATC TAC AAG GTG AAG CAT TAA AGC AGC GTA TCC ACA tdTomato ATG C TAG CG

Rosa26 AAG GGA GCT GCA GTG GAG TA CCG AAA ATC TGT GGG AAG TC

3.2.3 Hematoxylin and Eosin staining for fiber size distribution

Hematoxylin and Eosin (H&E) staining was performed on 14 µM muscle sections as described (Le et al., 2005). Briefly, slides were fixed by dipping in 50%, 70%, 90% and finally 100% ethanol, followed by 3 washes in tap water and Hematoxylin (30s, Sigma).

After 3 washes in tap water, they were stained in Eosin (Sigma) for 20s and washes thrice in tap water and dehydrated in 50% (4x), 70% (4x), 90% (7x), 100% (10x) and Histoclear

(2 round of 10x each, Sigma). Mounting was done with Cytoseal-280 (Richard-Allen

Scientific). The slides were imaged on Nikon Eclipse 800 microscope (Nikon 74

Corporation, Japan) with Nikon FDX-35 digital camera. Morphometric measurements were done using SPOT Advanced (v3.5.9) software (Diagnostic Instruments, Inc., MI), after binning the frequency of fiber size area on Microsoft Excel 2007. All investigators were blinded to genotype during phenotypic assessment.

3.2.4 Immunohistochemistry

PND12 mouse pups were perfused with PBS followed by 4% paraformaldehyde (pH 7).

The tissues post-harvesting were infused with 30% sucrose and flash frozen in isopentane cooled to -150 ºC in liquid nitrogen. The frozen tissue was embedded in Tissue-Tek OCT

(Fisher Scientific) and cryostat (IEC Minotome Plus) sectioned. 14 µM thin sections were blocked with goat serum (30 min), and stained with Chicken anti-red RFP (1:2500,

Millipore AB3528) and Rabbit anti-GFP (1:1000, Molecular Probes A11122) for 2 hours.

Secondary antibody incubation (Alexa-488, Alexa-594 at 1:1000 each) was done for 30 min. Washes with PBS (1.0% Tween) (6x, 30 min) were done before and after application of secondary antibody. Flouromount-G (Southern Biotech) was used for mounting the tissues. For visualizing gems, SMN was similarly stained with mouse anti-

SMN (1:500, BD Biosciences 610647; Alexa-488 secondary) and DAPI was used to visualize the nucleus. All confocal imaging was performed using Leica DM IRE2 with

Leica TCS SL point scanning laser confocal system with photomultiplier tube detection.

Subsequent image processing was done using Adobe Photoshop CS2.

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3.2.5 Muscle physiology tests

In both male and female mice, at age of 8 weeks, muscle function test were performed, with the investigator being blinded to the genotype. First, electrocardiograms were obtained in unrestrained, unanaesthetized mice using the ECGenie system. Briefly, the mice were allowed to sit on a platform that registered ECG-signals through a set of three electrodes located on the platform. From the obtained recordings, heart rate, heart rate variability, and QT interval, an important indicator for cardiac muscle dysfunction, were assessed. Next, after cervical dislocation, the extensor digitorum longus (EDL) muscle was dissected, and mounted into a muscle physiology set-up, and functional assessment of contractions were assessed as previously described (Martin et al., 2009). Briefly, after stretching the muscle to optimal length, using twitch-contraction force as a read-out, a maximal tetanic force was achieved by applying a 250 ms duration 180 Hz stimulation train (1 ms per pulse) to the EDL muscle. Thereafter, a series of 10 eccentric contractions were performed, in which the muscle was stimulated for 700 ms total duration (180 Hz pulses of 1 ms duration), while in the last 200 ms the muscle length was linearly increased by a servo-controlled motor, for a total stretch of 3% of the muscle length. At t=700 ms, the stimulation was halted and the muscle was returned within 200 ms to its original optimal length. The muscle was allowed to rest for 2 minutes in between contractions. After assessment of contractions, the muscle was removed from the set-up, blotted dry in between 2 kimwipes using a 10 gram weight for 10 seconds, and muscle weight was assessed. All obtained forces were normalized to the cross-sectional area of the muscle to calculate specific force of the muscle.

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3.2.6 H&E staining for LCM

Slides for LCM (Zeiss Membrane Slide 1.0 PEN, 415190-9041-000) were treated with

UV radiation for 30 min just before blotting 14 µm gastrocnemius sections on to them.

For H&E staining, the slides were fixed in 70% ethanol (1 min), followed by 50% ethanol (1 min) and H2O (3x over 1 min). Muscle sections were stained with Weigert’s iron Hematoxylin (Sigma) for 1 hour, rinsed in dH2O, destained in 0.3% acid alcohol (5-7 dips) and rinsed in tap water. Next, the sections were stained with Eosin for 5 sec (Sigma) and dehydrated with 70%, 90% and 100% ethanol, air-dried shortly (10 min) and stored at 4 ºC.

3.2.7 Laser-capture Microdissection (LCM)

Laser capture microdissection (LCM) was performed with the Palm Microbeam (Carl

Zeiss MicroImaging) under 10x magnification. Approximately 150,000-200,000 µm2 of muscle tissue was collected from H&E stained slides, avoiding the extra-sarcomeric tissue. The tissue was collected in 18 µl of cell-lysis solution containing 20mM Tris-HCl pH 8, 0.02% Tween and 1 mM EDTA and frozen on dry ice. Before performing ddPCR,

2 µl of 2 mg/mL proteinase K (freshly prepared 1:10 dilution of 20 mg/mL of proteinase

K (Invitrogen) in a solution of 20mM Tris-HCl pH 8, 0.1 mM EDTA) was added to the sample. The sample was heated at 55 ºC for 2 hours, followed by inactivation of proteinase K at 90 ºC for 10 minutes. Siliconized tubes and low-bind tips (Eppendorf) were used throughout the experiment to prevent the loss of low amounts of DNA via adsorption onto plastic. Siliconization was performed as described in Sambrook’s

Laboratory Manual (Sambrook, 1987).

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3.2.8 Droplet digital PCR (ddPCR)

LCM-collected tissue was divided into two parts and multiplex ddPCR was thus performed in duplicates with each sample. After addition of primers, probes and ddPCR

SuperMix (BioRad), the sample was partitioned into approximately ten to sixteen thousand droplets and PCR was performed. The fluorescence signal was measured and calculated according to Poisson statistics and analyzed using QuantaSoft (BioRad) software (Porensky et al., 2012). The sequence of the primers and probes are as follows:

Smn exon 7 – FP 5’ AAGATGACTTTGAACTCCGGGTCCT, RP 5’

TGTGAGTGAACAATTCAAGCCC, Probe 5’ FAM-

CTGCCCATGCATCACCAAGCTTGG CAT-MGB; Smn intron 1 – FP 5’

CTGTGTGACTGTGAGGGGATGTG, RP 5’

CCTGTGAACATCTTCATCCTGACCTAA, Probe – 5’ VIC-

AGGCTGGCTGAAGCAAGG CAACCAGATA-MGB. Eppendorf LoBind tips were used throughout to prevent loss of DNA.

3.2.9 ELISA on whole muscle sections

Gastrocnemius muscle of PND10 mice was flash frozen in isopentane cooled to -150 °C in liquid nitrogen and cryostat (IEC Minotome Plus) sectioned. 2-4 sections of 14 µm thickness were collected in 100 µl of ELISA-lysis solution (PharmaOptima, MI). 40 µl of whole muscle section lysates were diluted 1:2 with sample buffer to a final volume of 80

µl. 25 µl of the lysate was used in the SMN ECL immunoassay. The assay is a quantitative sandwich immunoassay, where a mouse monoclonal antibody (2B1, Liu and

Dreyfuss 1996) functions as the capture antibody and a rabbit polyclonal anti-SMN

78 antibody (Protein Tech, Cat. No. 11708-1-AP) labeled with a SULFO-TAG™ is used for detection. SMN levels are determined from a standard curve using recombinant SMN protein (Enzo Life Sciences, Cat. No. ADI-NBP-201-050) as calibrator. The dynamic range of the assay is 10 pg/ml (lower limit of quantitation) to 20,000 pg/ml (upper limit of quantitation). Assay plates were read using a Meso Scale 6000 sector imager. 5 µl of non-diluted lysate was used for Pierce BCA protein assay. SMN levels were then normalized to total soluble protein.

3.2.10 Western blot analysis

Western blots were performed as previously described (Gavrilina et al., 2008; Le et al.,

2005; Porensky et al., 2012). Briefly, 50 µg of protein (skeletal muscle, PND10 pups) was loaded per lane and SMN was detected with mouse anti-SMN (1:1500, BD

Biosciences, 610647) while tubulin was detected with mouse anti-tubulin (1:25,000,

Sigma, T8203). Mouse-anti HRP (1:10,000, Jackson Immuno Research) was used as the secondary antibody and the blot was visualized using the ECL system, as described by the manufacturer (GE Healthcare Life Sciences). Blots where scanned and quantified as described (http://lukemiller.org/index. php/2010/11/analyzing-gels-and-western-blots- with-image-j/) and the area under each peak determined with ImageJ software.

3.2.11 Statistical analyses

Survival (Kaplan–Meier curve) and statistical analyses (Mann–Whitney Rank Sum Test,

Shapiro–Wilk Normality Test, t-test) were done using SigmaPlot v12.0 (Systat Software

Inc., CA). Weight curve analysis was performed with Statmod (Baldwin et al., 2007; Elso et al., 2004).

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3.3 RESULTS

3.3.1 Testing the deletion and replacement Smn alleles

To selectively reduce SMN in a tissue, the murine line SmnF7 with floxed exon 7 was used (Frugier et al., 2000) as one allele, with the other allele being the exon 2A disruption, SmnKO allele, which results in the production of no functional mouse SMN

(Schrank et al., 1997). The floxed Smn lines and the Cre lines were each first crossed on to the SMNΔ7 SMA background and were made homozygous for the SMN2 and SMNΔ7 transgenes and heterozygous for SmnKO and Cre respectively. Thus, SMN2 and SMNΔ7 in the background provide low SMN essential for viability in all cells. Breeders positive for Cre and the SmnKO allele were bred to the floxed Smn mouse lines (Le et al., 2005;

Monani et al., 2000; Schrank et al., 1997). As shown in Figure 3.2A, upon action of Cre, exon 7 gets deleted, resulting in SmnD7 allele (for deletion of Smn), creating an SMA- situation in the places Cre is expressed. For replacement experiments, the rescue allele

SmnINV (also referred to as SmnRes in other works) was used (Lutz et al., 2011). SmnINV allele is a hybrid allele with human SMN exon 7 joined to inverted mouse exon 7, flanked by lox66 and lox71 sites, which enable reversion of the cassette upon Cre recombination (Oberdoerffer et al., 2003). Thus the floxed inverted mouse exon 7 is reverted to the correct orientation upon Cre recombination, called SmnRe (for reversion of exon 7), and expression of mouse Smn is restored (Figure 3.2). I sequenced the SmnINV and SmnRe alleles (Figure 3.1).

In previous experiments with Cre drivers, the floxed lines, SmnF7 and SmnINV, have been used with both Smn alleles bearing the floxed allele (Cifuentes-Diaz et al.,

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2001; Frugier et al., 2000; Lee et al., 2012; Lutz et al., 2011; Martinez et al., 2012; Paez-

Colasante et al., 2013; Vitte et al., 2004). We chose to have one allele that could either be deleted or reverted; the design of crosses is shown in Figure 3.2B. This was for two reasons: firstly the Cre driver can be separated from the floxed exon 7 allele, thus eliminating any chance of germline conversion, and second, the Cre only has to act on one allele, maximizing its chance of action on the floxed allele. To first demonstrate that the floxed Smn alleles, SmnF7 and SmnINV, for deletion and replacement respectively are fully functional, I crossed the floxed Smn alleles to a ubiquitous Cre line, Sox2-Cre. The

Sox2-Cre transgenic line has expression of Cre under the Sox2 promoter element and efficient Cre recombination is seen in all epiblast-derived cells by e6.5 (Tajbakhsh et al.,

1996). Sox2-Cre mice were crossed onto the SMA background and made homozygous for SMN2 and SMNΔ7. Mice with Sox2-Cre; SmnD7/KO have Smn-deletion in all tissues and thus all tissues are dependent on SMN produced by SMN2 and SMNΔ7. Sox2-Cre;

SmnD7/KO mice had a phenotype and weight (Figure 3.2D, n=9, blue inverted triangle) similar to SMNΔ7 SMA mice and a mean survival of 13.8 ± 1.3 days (Figure 3.2C, n=9, blue line). Upon crossing the Sox2-Cre; SmnWT/KO mice to the replacement allele, SmnINV,

I obtained complete rescue of survival (Figure 3.2C, n=10, red line) and weight (Figure

3.2D, n=10, red diamond). The rescue mice, Sox2-Cre; SmnRe/KO appeared normal in phenotype, with no necrosis or decrease in weight.

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Figure 3.2. Smn deletion and replacement alleles used in the study. (A) Diagram of mouse Smn alleles used for the Cre recombination experiments: SmnF7 allele contains floxed exon 7 wherein exon 7 is deleted upon Cre recombination, giving the allele, SmnD7. SmnINV allele has mouse Smn exons 1-6 fused to human SMN exon 7 with an inverted mouse Smn exon 7. The lox66 and lox71 sites flanking the fused human and mouse exon 7 construct get reverted upon Cre recombination. With mouse Smn exon 7 in the right orientation, SmnRe produces full-length SMN protein. Red arrows indicate the lox sites. (B) Diagrammatic representation of strategy of mouse breeding: To avoid germline recombination and ensuring that Cre acts on only one floxed allele, we chose to maintain the floxed lines and Cre line separately. The Cre lines bore the SmnKO allele. These were crossed to either SmnF7 allele maintained in a homozygous state or SmnINV that was maintained over an SmnWT allele. The affected mice with Smn-deletion thus had SmnD7/KO and the Smn-replacement mice had SmnRe/KO. (C) Testing the floxed alleles with the ubiquitous Sox2-Cre driver: Deletion (blue line): Sox2-Cre; SmnD7/KO mice (n=9) had a mean survival of 13.8 ± 1.3 days which is similar to SMA mice (14.3 ± 0.7 days, n=16, green line). Replacement (red line): Sox2-Cre; SmnRe/KO mice (n=10) were phenotypically normal and survived beyond 36 weeks. (D) The total body weights of Sox2-Cre; SmnD7/KO mice (n=9, blue inverted triangle) are similar to SMA mice (n=16, green circle) while the weights of Sox2-Cre; SmnRe/KO mice (n=10, red diamond) are similar to controls (Sox2-Cre; SmnRe/WT). Thus, the SmnF7 and SmnINV alleles are functional. (Error bars = sem)

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Figure 3.2

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3.3.2 Expression of Myf5-Cre driver

We chose Myf5-Cre line to drive Cre in the muscle because Myf5 is the first skeletal muscle marker expressed by e8.5 in immature somites before terminal myogenesis (Ott et al., 1991; Tajbakhsh et al., 1996). Thus Smn will be deleted or replaced in the early stages of skeletal muscle development. Myf5-Cre line was made homozygous for SMN2 and SMNΔ7. To test the expression of Myf5-Cre, I crossed the

Cre line to an RFP reporter line, ROSA26-loxP-STOP-loxP-tdTomato (Madisen et al.,

2010). The reporter line has a floxed STOP codon upstream of tdTomato, a modified RFP gene. Upon excision of the STOP codon by Cre, RFP is expressed in that tissue (Madisen et al., 2010). To visualize motor neurons, mice with HB9:GFP which expresses GFP in the motor neurons under the murine HB9 promoter at e9.5 (Arber et al., 1999), were crossed to the tdTomato-RFP mice. Figure 3.3 shows immunostained spinal cord and skeletal muscle sections of PND12 mouse pups bearing Myf5-Cre, tdTomato and

HB9:GFP. Robust RFP expression was observed in the skeletal muscle (Figure 3.3D-E).

However, rare motor neurons in the spinal cord also showed RFP expression (Figure

3.3A-C). Out of 4 Myf5-Cre; tdTomato; HB9:GFP animals examined, leaky expression was found in some cells in the heart, lung, liver and pancreas in half of the animals examined. Only 1 animal showed RFP expression in kidney and adrenal gland (Figure

3.4). Scattered blood vessels and connective tissue cells were also positive for Myf5-Cre expression in all organs examined above. In all cases the leaky expression was in a few cells in the entire section of the tissue. Also, it should be noted that not all animals showed leaky expression in every tissue and to the same degree. Myf5-Cre is thus a

84 chimeric driver with ectopic expression in a small percentage of cells in other tissues.

Unlike the tissues mentioned above, the skeletal muscle consistently showed uniform strong expression of RFP. We thus used the Myf5-Cre line to determine the importance of

SMN levels above that produced by two copies of SMN2 (and SMNΔ7) to the normal function of muscle.

Figure 3.3. Immunohistochemistry of Myf5-Cre; tdTomato; HB9:GFP mice (PND12): (A and D) indicate expression of RFP, (B and E) show GFP channel indicating motor neurons, and (C and F) show the overlaid images. (A-C) Lumbar spinal cord sections reveal Myf5-Cre expression (red) in a few motor neurons (green) indicated by white arrows. (D-F) Gastrocnemius muscle sections show robust RFP expression indicating high expression of Myf5-Cre in the muscle. Representative images from n=5 mice. (Scale bar = 100 µm)

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Figure 3.4. Immunohistochemistry of Myf5-Cre; tdTomato; HB9:GFP mice (PND12) indicating ectopic expression of the Myf5-Cre driver: Some blood vessels and connective tissue showed RFP expression in all the organs mentioned below. (A-C) 2/4 Heart sections showed a patch of cells expressing RFP. (D-F) Lung sections in 2/4 samples showed scattered RFP expression. (G-I) Scattered groups of bright red cells indicating Cre expression were observed in 3/4 liver samples. (J-L) Pancreas-staining revealed groups of acinar cells and islet cells (white arrow) expressing RFP in 2/4 samples. (M-O) Only 1/4 Kidney samples had scattered nephrons and tubules expressing RFP. (P-R) Adrenal gland cells in the medulla and cortex in 1/4 samples showed RFP expression. (Scale bar = 100 µm)

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Figure 3.4

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3.3.3 Deletion and replacement of Smn in muscle tissue in the SMNΔ7 SMA mouse model

SMN levels in the muscle were selectively reduced using the Myf5-Cre driver

(Tajbakhsh et al., 1996). The Myf5-Cre; SmnD7/KO; SMN2+/+; SMNΔ7+/+ mice had deletion of the exon 7 of the floxed Smn allele in the skeletal muscle. Thus the muscle becomes reliant on the SMN produced by 2 copies of SMN2 (and SMNΔ7). Myf5-Cre;

SmnD7/KO mice appeared phenotypically normal, showing no signs of weakness or necrosis. They had normal survival as shown in (Figure 3.5, blue line) and weight

(Figure 3.5B, blue inverted triangle, p>0.05 vs. control, ns) (n=15). The Smn-deletion pups showed no righting defects either (n=4, righting time < 10s). Furthermore, gastrocnemius muscle sections of Myf5-Cre; Smn-deletion mice were stained for SMN.

We observed no gems of SMN in the muscle nuclei of Myf5-Cre; Smn-deletion mice, which was similar to SMA, indicating that there was a decrease of SMN in the muscle

(Figure 3.6B and C). Mice with a WT copy of Smn on the contrary showed the presence of gems in the nucleus (Figure 3.6A, white arrows).

Lastly we examined the effect of SMN restoration in the muscle in mice where all other tissues are in an SMA-like state, with SMN only from SMN2 and SMNΔ7. When

Smn is replaced in the muscle of SMA mice (Myf5-Cre; SmnRe/KO), the affected mice had a mean survival of 14.1 ± 2.4 days (n=11) which is identical to the survival of the

SMNΔ7 SMA mice (14.3 ± 0.7 days, n=16) (Figure 3.5A, red line). There was no marked difference between the mice with SMN restoration in muscle and SMA mice with respect to weight (Figure 3.5B, red diamond) and phenotype. Analysis of the weight

88 curve of Myf5 Smn-replacement mice vs. SMA at days 10 and 14 revealed no significant difference (p=0.69 and 0.24 respectively). The results obtained with the replacement experiments are consistent with 2 copies of SMN2 (and SMNΔ7) providing the required amount of SMN for normal function of muscle.

Figure 3.5. Survival and weight analyses of muscle driver Myf5-Cre: (A) Deletion (blue line): Affected mice, Myf5-Cre; SmnD7/KO (n=15), had a normal survival. Replacement (red line): Mean survival of affected mice, Myf5-Cre; SmnRe/KO mice (n=11) was 14.1 ± 2.4 days, similar to SMA mice (14.3 ± 0.7 days, n=16, green line). (B) In Myf5-Cre; SmnD7/KO mice, the weight curve was similar to controls (blue inverted triangle). Mice with Smn replacement in the muscle, Myf5-Cre; SmnRe/KO (n=11, red diamond) were similar to SMA mice (n=16, green circle) in body weight. (Error bars = sem)

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Figure 3.6. Immunohistochemical localization of SMN in gastrocnemius muscle sections (PND10): (A) SMN staining (green, indicated by white arrows) is observed in the nucleus (stained blue with DAPI) of muscle cells of SmnWT/KO mice without Cre. (B) SMA and (C) Myf5-Cre; Smn-deletion mice show no SMN staining in the nucleus. (Scale bar = 10 µm)

3.3.4 Efficiency of Cre recombination in the muscle

To determine the efficiency of Cre excision with a more direct read-out, we quantified the percentage of Cre recombination by performing droplet digital PCR

(ddPCR, BioRad) on the SmnF7 allele in the muscle tissue. Importantly, we eliminated the influence of connective tissue by laser capture microdissection (LCM) of gastrocnemius muscle cells, leaving out the nuclei of extra-sarcomeric tissue (n=3 per genotypic group). ddPCR of SmnF7 was normalized to Smn intron 1 (2 copies per genome) in a multiplex reaction. ddPCR of LCM-collected Sox2-Cre; SmnD7/KO gastrocnemius muscle (PND9-

15) showed 100 ± 5.5% recombination while No Cre; SmnF7/KO had no recombination

(0.6 ± 14.7%). Myf5-Cre; SmnD7/KO gastrocnemius (PND10) tissue collected via LCM showed 80.8 ± 4.2% recombination (Figure 3.7A), thus proving that Myf5-Cre efficiently

90 acts on the floxed alleles in the skeletal muscle. The efficiency of Cre recombination by

Myf5-Cre is less than the expected cent percent for two reasons as follows. Myf5-Cre is not expressed in the subsynaptic muscle nuclei (Dr. Jill Rafael-Fortney; personal communication). In our immunohistochemistry studies, we have observed specks of

HB9:GFP staining indicative of the presence of motor neuron in muscle sections (data not shown); implying that the muscle cell adjoining the motor neuron would be a subsynaptic muscle cell not expressing Myf5-Cre. Secondly because the technique of

LCM aids in the collection of a muscle nuclei-enriched sample, we cannot rule out the possibility that our samples would have a small percentage of connective tissue that do not express Myf5-Cre. Thus the presence of nuclei of subsynaptic muscle cells and connective tissue would cause the efficiency of Myf5-Cre recombination to be reported as less than 100 percent in our experiment. Hence an efficiency of 80% Myf5-Cre recombination is sufficient to imply complete recombination of floxed Smn alleles in the muscle cells.

3.3.5 Determination of SMN protein levels in muscle

SMN protein was quantified via ELISA immunoassay (proprietary,

PharmaOptima, MI, USA) in cryostat-sectioned whole muscle of PND10 pups. Myf5-

Cre; SmnD7/KO mice, as shown in Figure 3.7B, displayed a decrease in SMN levels. The

SMN level in the Myf5-Cre mice was higher than the SMA sample, which is not unexpected due to SMN contribution from non-muscle cells such as connective tissue and blood vessels. There was no statistical difference between SMN levels in SMA and

Myf5-Cre; SmnD7/KO mice as the ELISA assay showed relatively high variation. We thus

91 also analyzed SMN protein levels in skeletal muscle via western blotting. Figure 3.7C and D respectively show representative bands of SMN and tubulin with quantitation of

No Cre; SmnF7/KO and Myf5-Cre; SmnD7/KO mice (PND10, n=3 in each group). Upon deletion of Smn with Myf5-Cre, there was a significant decrease of SMN protein in skeletal muscle to 31.3% of control No-Cre levels (2.08 ± 0.13 vs 0.65 ± 0.26, p = 0.009).

We believe an approximate drop of 70% in SMN levels is sufficient to imply good recombination by Myf5-Cre in muscle. In addition to small amounts of SMN protein produced by SMN2 and SMNΔ7, there would be contribution of non-muscle cells of connective tissue and post-synaptic muscle as described above in the whole muscle lysate. Martinez et al. reported a robust increase of SMN transcript and protein upon replacement of Smn using Myf5-Cre (Martinez et al., 2012). Other groups have shown efficient recombination by Myf5-Cre in muscle and an 80% drop in RNA and protein levels of the floxed allele (Klover and Hennighausen, 2007). Beedle et al. have shown that deletion of Fukutin, a gene required for dystroglycan processing, by Myf5-Cre induced a severe dystrophic phenotype, and a near-absence of functionally glycosylated dystroglycan (Beedle et al., 2012). Lack of endogenous Myf5 expression in fibroblasts, adipocytes, neurons, blood vessels and 10% of Pax7-positive satellite cells included in whole muscle may account for the apparent lack of recombination (Beedle et al., 2012;

Kuang et al., 2007; Waddell et al., 2010). Thus the total SMN detected in whole muscle is consistent with an 80% deletion rate indicated by the DNA recombination test.

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Figure 3.7. Analysis of Cre activity upon deletion of Smn in muscle. (A) Quantification of percentage of Cre recombination on the SmnF7 allele: ddPCR of muscle nuclei collected via LCM of H&E stained gastrocnemius muscle of PND9-15 mice show complete recombination by the Sox2-Cre driver (100 ± 5.5%). The Myf5-Cre driver specific for skeletal muscle showed 80.8 ± 4.2% recombination of SmnF7 allele. (B) ELISA-estimation of SMN protein levels in whole muscle sections (PND10): Smn- deletion with Myf5-Cre leads to a drop in muscle SMN levels to 1510.9 ± 529.8 pg SMN/mg soluble protein from 4013.8 ± 1721.9 pg in No Cre; SmnF7/WT mice. SMA mice had a fall in SMN levels to 160.8 ± 33.8 pg/mg soluble protein. (C) Representative western blot bands of SMN and tubulin from No Cre; SmnF7/KO and Myf5-Cre; SmnD7/KO mice. (D) SMN protein levels determined by western blot relative to tubulin (PND10): A 68.7% decrease in SMN was observed in whole muscle of Myf5-Cre; SmnD7/KO mice. SMN to tubulin ratios at 2.08 ± 0.13 in No Cre; SmnF7/KO control were significantly different from 0.65 ± 0.26 in affected mice (P = 0.009) (n=3 in each group, error bars = sem)

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3.3.6 Functional analyses of muscle force upon Smn-deletion

To investigate the integrity and function of muscle that was dependent on SMN from SMN2 (and SMNΔ7), physiological tests of twitch force (Figure 3.8D), tetanic force (Figure 3.8E) and eccentric contraction-induced loss of force were performed

(Figure 3.8F) on the extensor digitorum longus (EDL) muscle at 8 weeks postnatally.

The investigators were blinded to the genotype of the mice. The muscle depleted for mouse Smn and dependent on SMN2 for full-length SMN showed no impairment of force production in any of the measures performed and was essentially identical to control muscle (n=8 control males, n=6 control females, n=5 test males and n=5 test females).

Furthermore, Myf5-Cre; SmnD7/KO mice did not show any difference in muscle length

(Figure 3.8A), diameter (Figure 3.8B) or weight (Figure 3.8C), as compared to the controls, Myf5-Cre; SmnD7/WT. I also plotted the fiber size distribution of the vastus lateralis muscle at 8 weeks of age. The fiber size distributions of control (Myf5-Cre;

SmnD7/WT, Figure 3.8I) and test Myf5-Cre (Myf5-Cre; SmnD7/KO, Figure 3.8J) mice do not differ (Mann-Whitney Rank Sum Test P=0.926, n=1350 fibers/group from 3 animals in each group). The heart rate was also investigated, although Myf5-Cre is not efficiently expressed in cardiac muscle. At 8 weeks of age, there was no significant difference in the unanaesthetized heart rate, heart rate variability and QT interval (Figure 3.9) of the control and Myf5-Cre test mice (n=5 in each group). Thus we conclude that 2 copies of

SMN2 (and SMNΔ7) provide sufficient full-length SMN for normal morphology and function of skeletal muscle.

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Figure 3.8. Muscle physiology tests on mice with SMN reduction in the muscle, Myf5- Cre; SmnD7/KO, at 8 weeks: There was no significant difference in the (A) Length (B) Diameter (C) Weight (D) Twitch force (E) Maximal Tetanic force and (F) Eccentric- contraction-induced loss of force of the EDL (extensor digitorum longus) muscle of control (Myf5-Cre; SmnD7/WT) and affected (Myf5-Cre; SmnD7/KO) mice (Control: n=8 males and n=6 females, Test: n=5 males and n=5 females). Representative H&E stained Vastus lateralis muscle of (G) control and (H) affected mice at 8 weeks of age and the muscle fiber size distribution of (I) control and (J) affected mice show no significant difference in morphology. (a total of n=1350 fibers/group from 3 animals in each group, Mann-Whitney Rank Sum Test, P=0.926, Error bars = sem)

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Figure 3.8

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Figure 3.9. Cardiac physiology tests on unanaesthetized mice with SMN reduction in the muscle, Myf5-Cre; SmnD7/KO, at 8 weeks: There was no significant difference in the (A) Heart rate (B) QT Interval and (C) Heart rate variability of control (Myf5-Cre; SmnD7/WT) and affected (Myf5-Cre; SmnD7/KO) mice. (n=5 animals in each group, Error bars = sem)

3.4 DISCUSSION

SMA is caused by insufficient levels of the SMN protein. The reduced levels of

SMN do not occur only in motor neurons but also in other tissues such as muscle. A number of studies have indicated that muscle could play a role in SMA. Muscle extracts from SMA Type III patients have been shown to inhibit neonatal chicken neurite outgrowth, albeit with no cytotoxicity (Henderson et al., 1987). In severe SMA mice there is no abnormality of neurite outgrowth in vivo (McGovern et al., 2008).

Furthermore, muscle biopsy from all SMA Type I patients and most Type II, but not

Type III, when co-cultured with normal rat motor neurons showed normal connection of the nerve to the muscle followed by sarcomeric disorganization and muscle degeneration

(Braun et al., 1995). Guittier-Sigrist et al. showed apoptotic myonuclei and increased release of microparticles in SMA muscle co-cultured with rat motor neurons (Guettier-

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Sigrist et al., 2002). The results would indicate an intrinsic defect in the muscle that only becomes apparent with connection to a nerve even though the nerve is normal. Further experiments revealed that when Type I SMA myoblast cells were mixed with 50% normal myoblasts cells, the hetero-myotubes corrected the degeneration phenotype seen in the co-cultures (Guettier-Sigrist et al., 1998). The aforementioned results suggest that treatment of muscle could have a major influence on the SMA phenotype. However in the current chapter we show that reduction of SMN to SMA-levels in muscle but not nerve does not result in any marked abnormality of muscle; in particular the contraction and force generating properties are normal. Studies in severe Smn-/-; SMN2+/+ SMA mice have shown that the number of satellite cells is comparable to wild-type controls

(Hayhurst et al., 2012). But when satellite cells of the severe SMA mice are cultured, they differentiate rapidly and fail to form multinucleated myotubes efficiently (Hayhurst et al., 2012). In the mice in the current study, the development, function and structure of muscle is normal upon decreasing SMN in the muscle. Myf5-Cre will be expressed in myoblasts at embryonic day 8 and thus the aforementioned defects maybe present in myoblasts and satellite cells in the Myf5-Cre; SmnD7/KO mice reported here. However they do not lead to morphological or functional defects. This is also consistent with the recent report of Kariya et al. which showed a marked reduction of SMN to levels produced by two copies of SMN2 at p50 in all tissues including muscle (Kariya et al., 2014). This

SMN-reduction did not result in a dramatic phenotype and grip strength was not reduced at 120 days. Some pathology of muscle did occur at 230 days with increase in central nuclei, which indicates muscle regeneration from satellite cells. It can be noted that

98 central nuclei is not a prominent feature of SMA muscle biopsies particularly in the milder forms of the disease (Dubowitz, 1985). One factor that we have not examined is the ability of the SMN-deficient muscle to perform repair. Indeed MyoD-/- mice with a homozygous deletion of an early myogenic regulatory factor, MyoD, appear phenotypically normal. MyoD-/- mice do not show a marked phenotype of deficiency in repair unless they are put under stress; in the case here by crossing with mdx mice which lack dystrophin (Megeney et al., 1996).

SMA mice and SMA muscle has been examined previously in the situation where

SMN is deficient everywhere. Cell culture studies of SMA muscle have shown abnormalities in myoblast fusion and malformed myotubes along with delayed expression of myogenic proteins (Boyer et al., 2014; Bricceno et al., 2014; Shafey et al., 2005). The results have been supported by in vivo studies showing defects in myogenic program and apoptosis in muscle in SMA mouse models (Boyer et al., 2014; Fayzullina and Martin,

2014). Furthermore, skeletal muscle from SMA Type I fetuses exhibit decreased myofiber diameter and fiber size distribution, and abnormalities in early markers of muscle development, leading to the hypothesis of delayed growth and maturation of skeletal muscle in SMA (Martinez-Hernandez et al., 2014; Martinez-Hernandez et al.,

2009). 54% of SMA Type I, II and III cases sampled in a study showed DNA fragmentation and immaturity of muscle fibers (Stathas et al., 2008). In the studies mentioned above, muscle appears to show a lack of maturity suggesting a delay in myogenesis. In addition, muscle in this state shows a reduced ability to produce force

(Boyer et al., 2013). However there are two difficulties in interpreting these results: first,

99 under conditions of denervation, the muscle atrophies. The fibers are smaller and likely not as developed therefore can produce less force. Indeed atrophied muscle due to disuse shows a number of changes and reduced capacity to produce force (Brooks and Myburgh,

2014). This effect of decreased force production is likely independent of SMN’s requirement in muscle. Second, no experiments were done to restore SMN in the muscle in vivo; for instance by using AAV-SMN to determine whether these alterations were reversible. Lastly Cifuentes-Diaz et al. reported that complete removal of Smn from muscle resulted in a dystrophic phenotype (Cifuentes-Diaz et al., 2001). However in

SMA, the SMN2 gene produces some full-length SMN and the critical question is whether this low amount of SMN is sufficient for normal function of muscle. Also, we have not observed loss of dystrophin staining in SMNΔ7 SMA mice (Le et al., 2005).

In the current chapter we found no apparent change in the phenotype of mice upon SMN-reduction in muscle, with the total body weight, fiber size distribution and survival comparable to healthy controls. Furthermore, at 8 weeks there was no change in twitch and tetanic force or eccentric-contraction-induced loss of force of EDL upon Smn- deletion in the muscle. The mice with Smn-replacement in the muscle were phenotypically similar to SMA mice, having a median survival of 16 days as compared to

14 days for SMA mice. Martinez et al. reported a median survival of 21 days in Myf5-

Cre; Smn-replacement mice (Martinez et al., 2012). An increase in muscle area and myofiber diameter was also reported in the Smn-replacement mice in this study. The slightly increased survival obtained in this study most likely relates to the SMA mice having two of the replacement alleles, i.e. Myf5-Cre; SmnRe/Re. The SMA mice without

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Cre, SmnINV/INV, had a median survival of 15 days (Martinez et al., 2012). Our laboratory has previously reported that extremely high levels of SMN in muscle specifically, with no expression in other tissues, did not improve survival, weight or fiber size (Gavrilina et al.,

2008). Both in the current study and previous study we looked at the complete distribution of muscle fibers and found no difference with SMN overexpression in muscle. An HSA-SMN line with leaky expression in the nervous system significantly increased the mean survival of SMA mice to 160 days (Gavrilina et al., 2008). Complete deletion of SMN in the muscle using the HSA-Cre driver resulted in severe muscle dystrophy and a drop in survival to a mean of 33 days but loss of SMN in all tissues tested to date results in severe problems (Cifuentes-Diaz et al., 2001). One important consideration with the use of HSA-Cre is whether there is any significant ectopic expression of the driver, as this has not been fully tested for HSA-Cre driver and we have previously shown that HSA can, in some transgenes, have ectopic expression (Gavrilina et al., 2008). In the latter case weak but clear expression could be observed throughout the nervous system. The specificity and strength of the Cre driver is an important consideration. Here we showed strong tdTomato-RFP expression in all muscle tissue using a loxP-stop-loxP-tdTomato line as well as efficient deletion of SMN exon7 in muscle using laser microdissection and droplet digital PCR. There is occasional and rare expression of Myf5-Cre in motor neurons but this is in only certain animals and only a few motor neurons in a section; so unlikely to have a major impact as the remaining motor neurons will sprout and compensate for the loss of only a few motor neurons.

Expression in a patch of cells was observed for this Myf5-Cre line in the heart, lung,

101 liver, pancreas and kidney in some, but not all animals examined. It is unlikely for a patch of Cre expressing cells in a tissue to significantly contribute to the overall phenotype. Another important aspect of muscle is that it is a syncytia with essentially a tube containing multiple nuclei. Given the volume of a muscle fiber (5nl for a mouse limb fiber and five orders of magnitude larger that of a small uninucleated cell, the presence of one nucleus at the postsynaptic plate is unlikely to make major contribution over the whole muscle (Bruusgaard et al., 2003). It is also well known that proteins in muscle can have a relatively restricted nuclear domain with the product produced by that nucleus restricted to a relatively tight region (Brooks and Myburgh, 2014; Bruusgaard et al., 2003; Folker and Baylies, 2013; Gundersen and Bruusgaard, 2008; Hall and Ralston,

1989; Kong and Anderson, 2001; Pavlath et al., 1989). In the current experiments we have shown that at least 80% of the muscle nuclei have loss of the mouse Smn exon7 and thus become reliant on SMN2. This also resulted in a drop of SMN protein in muscle.

Given the large volume of a muscle cell and the nuclear domain, this should result in a myopathic phenotype being revealed and an abnormal force of contraction being present.

However this is not the case and thus we conclude that two copies of SMN2 provide sufficient SMN for normal function of muscle.

We are of the opinion that it is important to perform both replacement and removal of SMN from a tissue to fully understand its role. Thus in the current chapter it is clear that reduction of SMN to SMA-levels does not have a major impact on muscle and indicates a minimal role for muscle in SMA.

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Many studies have targeted muscle for therapy and amelioration of atrophy.

Follistatin injections which block myostatin thus enhancing muscle mass, resulted in weight increase and a mild increase in survival (Rose et al., 2009). Transgenic overexpression of follistatin or inactivation of myostatin did not ameliorate the disease in

SMA mice (Rindt et al., 2012; Sumner et al., 2009). In Chapter 2, we show that deletion of muscle ubiquitin ligases, MAFbx and MuRF1, which target the muscle proteins for degradation, did not alter the disease course in SMA mice (Iyer et al., 2014). Treatment of Smn2B/- SMA mice with Fasudil, an inhibitor of RhoA/Rho kinase pathway, improved the muscle fiber size but did not improve the performance (Bowerman et al., 2012b).

Thus even therapies that target various aspects of muscle have so far had minimal impact.

Various therapeutic strategies targeted towards the central nervous system have been explored in SMA. Gene therapy with delivery of SMN in an scAAV9 viral vector increased the survival of SMA mice beyond 250 days (Dominguez et al., 2011; Foust et al., 2010; Valori et al., 2010). Secondly, anti-sense oligonucleotides that trick the SMN2 gene into splicing like SMN1 and thus increase the production of full-length SMN have been successful in increasing the life span and improving the phenotype in SMA mice

(Hua et al., 2011; Passini et al., 2011; Porensky et al., 2012). Thirdly, from a wide-array of compounds, specific compounds that modulate the splicing of SMN2 gene and promote the incorporation of exon 7 were screened for (Naryshkin et al., 2014). Daily oral dosing of three promising compounds in SMA mice showed a marked increase of full-length SMN in brain, spinal cord and muscle along with an increase in survival and body weight (Naryshkin et al., 2014). Increased SMN expression in the neurons is an

103 important factor for improvement of the SMA phenotype in the above-mentioned therapies. Recently, the importance of high SMN in the peripheral tissues has been indicated (Hua et al., 2015). We have also examined neuronal SMN and we discuss the role of the neurons and the periphery in the following chapter (McGovern et al., 2015).

Given our results in the present work where deletion of Smn in the muscle did not decrease the muscle fiber size or contractile function, and replacing Smn solely in the muscle did not improve the survival or weight of SMA mice, we conclude that low SMN levels is sufficient for normal muscle function.

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CHAPTER 4

SMN expression is required in motor neurons to rescue electrophysiological deficits

in the SMNΔ7 mouse model of SMA

4.1 INTRODUCTION

The motor neuron disorder proximal 5q SMA is the most frequent hereditary cause of death in infants (Roberts et al., 1970). The loss of motor neuron function results in paralysis of muscles and respiratory failure (Crawford and Pardo, 1996). SMA is caused by the loss of function of SMN1 and retention of SMN2 (Burghes, 1997; Lefebvre et al., 1995). Both genes encode the SMN protein, however the SMN2 gene produces only a small amount of full-length transcript. The truncated SMN protein lacking the amino acids encoded by exon 7 does not oligomerize efficiently and is rapidly degraded

(Burnett et al., 2009; Lorson and Androphy, 2000; Lorson et al., 1998). The ubiquitously expressed SMN protein plays an essential role in snRNP assembly and splicing in all cell types. Yet reduced levels of SMN protein result in the selective loss of motor neurons and impairment of the neuromuscular junction (NMJ) formation and maturation (Kariya et al., 2014; McGovern et al., 2008; Murray et al., 2008). Over expression of SMN has been shown to completely ameliorate the SMA phenotype in mice (Gavrilina et al., 2008; 105

Monani et al., 2000). Several promising therapies that increase the amount of full-length

SMN protein including gene therapy, oligonucleotide therapy and small molecules have been developed (Dominguez et al., 2011; Duque et al., 2009; Foust et al., 2010; Hua et al., 2011; Naryshkin et al., 2014; Passini et al., 2011; Porensky et al., 2012; Valori et al.,

2010). Yet any therapy if administered at the wrong time or in the wrong tissue will not be effective. Previously, others have studied the temporal requirement for SMN expression in the mouse (Le et al., 2011; Lutz et al., 2011). It is critical for the advancement of therapies in SMA to understand the spatial requirement of SMN for proper function of the motor neuron. Previously our laboratory has reported that over expression of SMN in neurons completely ameliorates the SMA phenotype while over expression in muscle had no effect (Gavrilina et al., 2008). Other studies have suggested the importance of high levels of SMN in neurons by either deleting or replacing Smn and then examining neuromuscular junction (NMJ) physiology and morphology (Gogliotti et al., 2012; Lee et al., 2012; Martinez et al., 2012; Paez-Colasante et al., 2013; Park et al.,

2010; Taylor et al., 2013). While NMJ morphology was improved there was no substantial increase in survival of the SMA mouse in these studies.

Here we present a comprehensive study of the requirement of SMN in the motor neuron, neurons and glia with both elimination of Smn using a floxed exon 7 allele

(SmnF7) and replacement of Smn using a hybrid Smn exon 7 allele (SmnINV) in the

SMNΔ7 mouse model of SMA (Frugier et al., 2000; Lutz et al., 2011). We assayed the function of the motor unit using clinically relevant electrophysiological measurements. In these experiments SMN2 and the SMNΔ7 transgene provide the ubiquitous low level of

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SMN protein required for survival of all cells and tissues (Dominguez et al., 2011;

Monani et al., 2000). We found that the reduction of SMN from all neurons and glia resulted in loss of function of the motor unit and death of the mouse. Conversely, replacement of SMN in neurons and glia corrected the functional output of the motor unit and survival of the mouse. Interestingly, replacement of Smn in just the motor neuron did not alter the survival of the SMA mouse yet the functional deficit of the motor unit was fully restored. Furthermore replacement of Smn in neurons and glial cells was sufficient to rescue both survival of the mouse and the function of the motor unit in the SMNΔ7 mouse model.

4.2 MATERIALS AND METHODS

4.2.1 Mouse breeding

This study was carried out in strict accordance with the recommendations of the

Institutional Animal Care and Use Committee and University Laboratory Animal

Resources at The Ohio State University under protocol number 2008A-0089. Floxed Smn alleles used in this study are listed in Table 4.1 and described here. Mice containing the floxed Smn exon 7: SmnF7, Smn1tm1Jme/J (JAX #006138) or the conditional inversion

Smn allele: SmnINV, Smn1tm3(SMN2/Smn1)Mrph/J (JAX007249) were crossed onto the

SMNΔ7 SMA mouse model: (FVB.Cg-Tg(SMN2)89Ahmb Smn1tm1Msd

Tg(SMN2*delta7)4299Ahmb/J (JAX #005025). Cre drivers used are listed in Table 4.2.

Cre drivers used include: Nestin-Cre, Tg(Nes-Cre) 1Kln/J (JAX #003771) (Tronche et al., 1999), ChAT-Cre, ChATtm1(cre)Low1/J (JAX #006410) (Lowell BB; Olson D; Yu J. 107

2006 direct submission to JAX); SYN1-iCre, Tg(SYN1-icre/mRFP1)9934Rdav/J (JAX

#012687) (Davis R. 2009 direct submission to JAX), rSyn1-Cre, Tg(Syn1-cre)671Jxm/J

(JAX #003966) (Zhu et al., 2001) and Sox2-Cre, Tg(Sox2-Cre)1Amc/J (JAX #008454)

(Hayashi et al., 2002). The HB9:GFP, Tg(Hlxb9-GFP)1Tmj/J (JAX#005029), line was used to identify motor neurons and the reporter line tdTomato, Gt(ROSA)26Sortm14

(CAG-tdTomato)Hze/J (JAX #007914), was used to identify recombinant tissues. All Cre drivers were also crossed onto the SMNΔ7 SMA mouse model background. To generate experimental animals the Cre drivers in the SMNΔ7 SMA background were crossed to either the SmnF7 line or the SmnINV line. After Cre activation the SmnF7 allele is referred to as SmnD7 (for deletion of exon 7) and the SmnINV line is referred to as SmnRe (for reversion of exon 7). All experimental animals contain the SmnKO allele in addition to the

SmnF7 or SmnINV alleles (i.e. Cre driver; SmnD7/KO; SMN2+/+; SMNΔ7+/+ or Cre driver;

SmnRe/KO; SMN2+/+; SMNΔ7+/+). To eliminate the possibility of germline recombination, known to occur in the Nestin-Cre line, only experimental offspring contained both floxed alleles and Cre drivers (Zhang et al., 2013a). Floxed allele and Cre driver breeder mice were maintained separately. Controls used in this study contained the Cre driver and a wild-type Smn allele (i.e. Cre driver; SmnD7/WT; SMN2+/+; SMNΔ7+/+ or Cre driver;

SmnRe/WT; SMN2+/+; SMNΔ7+/+).

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Table 4.1 Cre drivers

Jax Strain Abbreviation Site of Cre Description References No. name used expression

Tg(Sox2- Expression of Cre 008- (Hayashi et Cre)1Amc Sox2-Cre Ubiquitous under mouse Sox2 454 al., 2002) /J promoter at e6.5 B6.Cg-Tg 003- (Nes- All neurons e10.5, rat Nestin (Tronche et Nes-Cre 771 Cre)1Kln/ and glia promoter al., 1999) J (Ivanova et B6;129S6- e10.5, endogenous 006- Cholinergic al., 2010; ChATtm1(cr ChAT-Cre mouse ChAT 410 neurons Tallini et e)Low1/J promoter al., 2006) B6(129S4 )- Tg(SYN1- 012- Human Synapsin I (Thiel et icre/mRFP SYN1-iCre All neurons 687 promoter al., 1991) 1) 9934Rdav /J B6.Cg-Tg 003- e12.5, rat Synapsin I (Zhu et al., (Syn1-cre) rSyn1-Cre All neurons 966 promoter 2001) 671Jxm/J Gad2/GAD (Katarova 65 (glutamic e10.5, endogenous et al., 2000; 010- Gad2tm2(cre acid Gad2-Cre mouse Gad2 Taniguchi 802 )Zjh/J decarboxyla promoter et al., se) positive 2011) neurons

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Table 4.2 Floxed alleles

Jax Recombination Result upon Cre Strain name References No. Before After recombination

006- Deletion of Smn (Frugier et Smn1tm1Jme/J SmnF7 SmnD7 138 exon 7 al., 2000)

007- Smn1tm3(SMN2/ Replacement of (Lutz et al., SmnINV SmnRe 249 Smn1)Mrph/J Smn exon 7 2011) Gt(ROSA)26 (Madisen 007- Sortm14(CAG- tdTom RFP expression et al., 914 tdTom tdTomato)Hze/J 2010)

4.2.2 Genotyping

Neonatal mice were tattooed and tail snips were obtained for PCR. The primers used to detect Cre drivers and Smn alleles are listed in Table 4.3.

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Table 4.3 Primer sequences

Transgene/ Forward primer Reverse primer Allele

F7 AGA AGG AAA GTG CTC ACA TAC TGT CTA TAA TCC TCA TGC TAT Smn AAA TT GGA G

INV ACT TCT TAA TTT GTA TGT GAG CGC TTC ACA TTC CAG ATC TGT Smn CAC T CTG

Re ATT TAA GGA ATG TGA GCA CCT CGC TTC ACA TTC CAG ATC TGT Smn TCC CTG

Cre CCT GTT TTG CAC GTT CAC CG ATG CTT CTG TCC GTT TGC CG

TCT CAA GAT TCT CTC AGA AAA GTT GCT GGA TAG TTT TTA CTG Nes-Cre TCA CC CCA G

CTC ATC TGT GGA GTT TGC AGA GAA AGA CCC CTA GGA ATG CTC ChAT-Cre AGC

SYN1-iCre CAG GGC CTT CTC CAC ACC AGC CTG GCT GTG AAG ACC ATC

HB9:GFP GTC GAG CTG GAC GGC GAC GT CTG CAC GCT GCC GTC CTC GA

ACG CTG ATC TAC AAG GTG AAG CAT TAA AGC AGC GTA TCC ACA tdTomato ATG C TAG CG

Rosa26 AAG GGA GCT GCA GTG GAG TA CCG AAA ATC TGT GGG AAG TC

4.2.3 Phenotypic assessment of mice

Mice were weighed daily starting at P1. At 21 days after birth pups were weaned and weighed weekly. Observation of weakness and necrosis was made every day, and then every week during weighing. Mice were sacrificed according to our IACUC approved protocol before they lost 30% of their maximum body weight. An equal number of males 111 and females were studied in each genotypic grouping. All investigators were blinded to genotype during phenotypic assessment. The number of animals required for analysis was determined as described previously (Butchbach et al., 2007).

4.2.4 Immunohistochemistry

Neonatal mice were transcardially perfused with PBS followed by 4% paraformaldehyde at P8. Tissues were cryopreserved in 30% sucrose overnight, embedded in Tissue-Tek

OCT (Fisher Scientific) and flash frozen in liquid nitrogen-cooled isopentane. Cryostat section (14 μm) were blocked with 4% goat serum in PBS and stained with antibodies including chicken anti-RFP (1:2500, Millipore AB3528), rabbit anti-GFP (1:1000,

Molecular Probes A11122), goat anti-ChAT (1:50, Millipore, AB144P) or mouse anti-

NeuN 1:50, Millipore, MAB377) for 2 h, followed by 1 h of secondary antibody incubation with Alexa-488 (1:1000, Molecular Probes A11008 and Alexa-594 (1:1000,

Molecular Probes A11042) and mounted in Flouromount-G (Southern Biotech). Confocal imaging was performed using Leica DM IRE2 with Leica TCS SL point scanning laser confocal system with photomultiplier tube detection. Image acquisition, overlays and scale bars were produced with Leica Confocal Software v2.61 and subsequential image processing was performed with Adobe Photoshop CS2. For the whole-body sagittal section, P2 mouse pups, following perfusion with PBS and 4% paraformaldehyde, were injected with molten 2% agarose in their body cavities. The perfused pups were then subjected to a gradient of sterile 10, 20 and 30% sucrose over a few days prior to freezing in liquid nitrogen-cooled isopentane and cryosectioning at 42 μm. Primary antibody was incubated overnight and rest of the immunostaining is as described above. The whole- 112 body sections were imaged with a Leica MZ-16FA stereomicroscope with PhotoFluor

LM-75 light source and Leica DFC-300 Fx camera. All scale bars (except whole-body) are in micrometers (μm).

4.2.5 Muscle fiber size analysis

Gastrocnemius muscle was cryosectioned at 14 μm and stained with H&E (for H&E staining, please refer to section 3.2.3). Muscle fibers were visualized with Nikon 1600 microscope and measured with SPOT Advanced (v3.5.9) software (Diagnostic

Instruments, Inc., MI, USA). The frequency of fiber size area was binned with Microsoft

Excel 2007. An equal number of males and females were studied in each genotypic grouping. All investigators were blinded to genotype during phenotypic assessment.

4.2.6 Laser capture microdissection

Lumbar spinal cord tissue was isolated at P10-P17 and flash frozen in liquid nitrogen- cooled isopentane. Spinal cordswere cryosectioned (14 μm) and adhered to PEN membrane slides (Zeiss). Tissue sections were dried at room temp and then fixed in methanol for 30 s, Nissl stained in 1% cresyl violate acetate for 1 min, washed in methanol 3 times and air-dried. Sections were stored at 4°. Laser capture microdissection

(LCM) was performed with the Palm Microbeam (Carl Zeiss MicroImaging) under 10x magnification. Motor neurons were identified by size and position in the ventral horn of the spinal cord. For DNA percent recombination by ddPCR: Approximately 200 motor neurons were collected per sample and processed as described previously in section 3.2.6.

Briefly, the LCM sample was collected directly into 18 μl of cell-lysis solution (20 mM

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Tris–HCl pH 8, 0.02% Tween, 1 mM EDTA) in siliconized tubes (Sambrook, 1987) and frozen on dry ice. Two microliters of 2 mg/ml proteinase K solution (Invitrogen) were added to the cell-lysis solution; followed by incubation at 55°C for 2 h and inactivation at

90°C for 10 min. For ELISA: Approximately 500–600 motor neurons were collected per sample into ELISA lysis buffer (PharmOptima, Portage MI) and frozen on dry ice.

4.2.7 Droplet digital PCR

Each digested LCM motor neuron sample was divided in half and run in duplicate in ddPCR for an approximate 100 motor neurons per reaction (QX200, BioRad). ddPCR reactions were prepared and amplified according to the manufacture’s recommendations

(BioRad). LoBind tips, plates and tubes (Eppendorf) were used to minimize DNA loss.

Primers and probe used to detect the SmnF7 allele: FP

5′AAGATGACTTTGAACTCCGGGTCCT, RP 5′TGTGAGTGAACAATTCAAGCCC, probe FAM-CTGCCCATGCATCACCAAGCTTGGCAT-MGB. Primers and probe used to detect total Smn DNA are located in intron 1: FP

5′CTGTGTGACTGTGAGGGGATGTG, RP

5′CCTGTGAACATCTTCATCCTGACCTAA, probe VIC-

AGGCTGGCTGAAGCAAGGCAACCAGATA-MGB. The percent of motor neurons displaying recombination was normalized to the total number of Smn alleles present as detected by the Smn intron 1 primers and probe.

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4.2.8 ELISA on LCM collected tissue

SMN protein was measured at PharmOptima (Portage, MI) using the company’s proprietary electrochemiluminescence (ECL) immunoassay based on Meso Scale

Discovery technology. The assay is a quantitative sandwich immunoassay, where a mouse monoclonal antibody (2B1, Liu and Dreyfuss 1996) functions as the capture antibody and a rabbit polyclonal anti-SMN antibody (Protein Tech, Cat. No. 11708-1-

AP) labeled with a SULFO-TAG™ is used for detection. SMN levels are determined from a standard curve using recombinant SMN protein (Enzo Life Sciences, Cat. No.

ADI-NBP-201-050) as calibrator. The dynamic range of the assay is 10 pg/ml (lower limit of quantitation) to 20,000 pg/ml (upper limit of quantitation). Assay plates were read using a Meso Scale 6000 sector imager. Samples from LCM were diluted with lysis buffer to a total volume of 40 μl. Twenty-five microliters of the lysate was used in the

SMN ECL immunoassay. SMN levels were then normalized with the number of motor neurons collected by LCM per sample.

4.2.9 Western blot analysis

Western blots were performed as previously described (Gavrilina et al., 2008; Porensky et al., 2012). Briefly, 50 μg of protein were loaded per lane. SMN was detected with mouse anti-SMN (1:1000, BD Biosciences, 610647); tubulin was detected with mouse anti-tubulin (1:10,000 Sigma, T8203). Mouse-anti HRP (1:10 000 Jackson Immuno

Research) and the ECL system was used to visualize the SMN protein as described by the manufacturer (GE Healthcare Life Sciences). Blots where scanned and quantified as

115 described (http://lukemiller.org/index. php/2010/11/analyzing-gels-and-western-blots- with-image-j/) and the area under each peak determined with ImageJ software.

Quantification of tubulin was performed with LI-COR 800 secondary antibody and the

LI-COR Quantitative Fluorescent Imaging Systems as previously described (Belteki et al., 2005).

4.2.10 Gem counts

SMN was detected with mouse anti-SMN (1:500, BD Biosciences, 610647) in 14 μm spinal cord cryo sections from P10 SMA, control, Nestin-Cre deletion and Nestin-Cre +

ChAT-Cre deletion mice (n = 3 for each genotype). The nucleus was stained with DAPI and mounted in Fluoromount G (Southern Biotech). Motor neurons were identified by their morphology and location in the spinal cord. The number of gems was counted in

100 nuclei for each sample under 60x magnification using a Nikon E800 microscope.

4.2.11 Electrophysiological studies of sciatic CMAP and MUNE

Electrophysiological techniques were performed as previously described (Arnold et al.,

2014). Briefly, electromyography (EMG), compound muscle action potential amplitude

(CMAP) and motor unit number estimates (MUNE) were recorded from sciatic- innervated hindlimb muscles at P21 in mice with deletion of Smn exon 7 with Nestin-Cre

+ ChAT-Cre drivers and compared with control mice. Similarly CMAP and MUNE were performed at P12 in mice with restoration of Smn exon 7 with Nestin-Cre + ChAT-Cre drivers and compared with SMNΔ7 SMA mice and control mice. An equal number of

116 males and females were studied in each genotypic grouping. All investigators were blinded to genotype during phenotypic assessment.

4.2.12 Statistical analyses

Statistical analyses were performed as previously described for MUNE and CMAP

(Arnold et al., 2014). Weight curve analysis was completed with Statmod (Baldwin et al.,

2007; Elso et al., 2004). Kaplan–Meier survival curves (log-rank), Mann-Whitney Rank

Sum Test and Shapiro-Wilk Normality Test were performed with SigmaPlot v12 (Systat

Software Inc., CA, USA).

4.3 RESULTS

4.3.1 Specific deletion and replacement of SMN upon Cre-mediated recombination in neural tissue

In order to study the requirement of SMN in the nervous system we selectively eliminated and replaced SMN expression using the Cre/lox system. These experiments were performed with multiple Cre drivers and floxed Smn alleles in the SMNΔ7 SMA mouse. The SMNΔ7 model contains two copies of the human SMN2 transgene, is homozygous for the SMNΔ7 transgene and has a disruption in the mouse Smn allele

(Smn−/−; SMN2+/+; SMNΔ7+/+) (Le et al., 2005). These experiments were conducted in the presence of low levels of SMN expression from the SMN2 and SMNΔ7 transgenes because the complete absence of SMN from any tissue is lethal (Cifuentes-Diaz et al.,

2001; Frugier et al., 2000; Vitte et al., 2004). SMN expression is required in all cell types 117 for survival due to SMN’s ubiquitous essential role in snRNP assembly. Thus we were able to specifically examine the effect of decreasing or increasing SMN levels in the nervous system while SMN2 (and SMNΔ7) provided the required low level of SMN protein necessary for general cellular development and viability.

To eliminate Smn expression the previously characterized Smn allele containing a floxed Smn exon 7 (SmnF7) was used (Frugier et al., 2000). Conversely, to replace Smn expression an Smn allele (SmnINV) containing a hybrid genomic cassette consisting of an inverted Smn exon 7 fused to human SMN exons 7–8 and flanked by lox66 and lox71 sites was used (Lutz et al., 2011; Oberdoerffer et al., 2003). Upon Cre-mediated recombination the entire cassette is inverted so that SMN is expressed from a mouse Smn exon 1–7/human SMN2 exon 8 hybrid allele (SmnRe) (Lutz et al., 2011). These Smn alleles are diagrammed in Figure 4.1A. The sequence of the SmnINV genomic cassette also contains a portion of adjacent intronic regions of the mouse Smn and human SMN alleles (Figure 3.2) (Iyer et al., 2015).

For full interpretation of the SMN requirement in nerve and muscle we chose to perform both deletion and replacement of Smn exon 7 with several Cre drivers. To eliminate the possibility of germ line recombination all floxed alleles were maintained separately from mice containing Cre drivers. The progeny of floxed alleles crossed to Cre drivers were only used for study and never for subsequent breeding. The mouse crosses used in this study to generate deletion of Smn (SmnD7) and replacement of Smn (SmnRe) are diagramed in Figure 4.1B. Littermates with a WT copy of Smn and the Cre driver

(Cre+; SmnD7/WT or Cre+; SmnRe/WT) were used as corresponding controls.

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Before selectively eliminating or replacing Smn in neuronal tissue we first determined the integrity of each Smn allele with the ubiquitous Sox2-Cre driver. Upon

Sox2-Cre-mediated deletion of Smn exon 7 (SmnD7), the mice developed an SMA-like phenotype and survived on average 13.8 ± 1.3 days (n = 9, Figure 3.1C and D) (Iyer et al., 2015). Conversely, as expected, ubiquitous Sox2-Cre recombination of the SmnINV allele resulted in mice that lived beyond one year of age with no phenotype (n = 10,

Figure 3.1C and D) (Iyer et al., 2015). Once it was established that the Smn alleles were fully functional I crossed the SmnF7 and the SmnINV alleles to five different Cre drivers

(Nestin-Cre, ChAT-Cre, Gad2-Cre, rSyn1-Cre and SYN1-iCre) listed in Table 4.2. The primers used to genotype the Smn alleles and the Cre lines are listed in Table 4.3.

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Figure 4.1. Diagram of Floxed Smn alleles and crosses used in this study. (A) The disrupted Smn allele is referred to as SmnKO. The Floxed exon 7 Smn allele is referred to as SmnF7 before recombination. After recombination when exon 7 is deleted the allele is called SmnD7. For the replacement of Smn, the allele termed SmnINV, contains human Smn exon 7 with flanking regions of intron followed by an inverted copy of mouse Smn exon 7 with flanking regions of intron. The two copies of exon 7 were cloned in between lox66 and a lox71 sites. Upon activation of recombination by a Cre driver the area between the lox sites is inverted resulting in a functional Smn exon 7 followed by an inverted SMN exon 7. After recombination this allele is referred to as SmnRe. In all crosses the SmnINV or SmnD7 alleles were crossed to mice that that were heterozygous for the SmnKO allele and contained a Cre driver. Thus all affected animals reported here are SmnD7/KO or SmnRe/KO after recombination. (B) Mice containing a Cre driver and heterozygous for the Smn knockout allele (SmnKO) were crossed to mice homozygous for the floxed exon7 SmnF7 allele. Upon recombination the floxed exon7 Smn allele is referred to as SmnD7. Similarly, mice that contain a Cre driver and are heterozygous for the Smn knockout allele (SmnKO) were crossed to mice heterozygous for the floxed exon7 SmnINV allele. Mice homozygous for the SmnINV allele are not viable beyond 21 days of age. Upon recombination the floxed exon7 Smn allele is referred to as SmnRe. Note that the progeny of these crosses only contain one deletion or replacement Smn allele over the SmnKO knockout allele. This is to ensure maximal efficiency of the Cre driver on just one target allele. Moreover, the progeny of these crosses were only used for data analysis and not for breeding to eliminate the possibility of germ line recombination.

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Figure 4.1

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4.3.2 Characterization of Cre driver using tdTomato-RFP expression

While in these experiments SMN2 provides a low level of SMN in all tissues, any additional deletion or replacement of Smn due to ectopic expression of the Cre driver could contribute to the overall weight and survival of the mouse. Thus it is important to ascertain the expression of any Cre line outside of the tissue of interest. To fully characterize the expression of each Cre driver, the tdTomato-RFP reporter line was used

(Madisen et al., 2010). When a Cre line is crossed to tdTomato-RFP reporter line the floxed stop cassette preceding RFP is removed allowing RFP expression irreversibly in the cells in which the Cre driver is expressed. Thus, the tissues in which recombination has occurred can be identified by the presence of RFP immunofluorescence in those cells.

The mice also contained the HB9:GFP transgene to identify the motor neurons (Arber et al., 1999). Expression of ChAT-Cre was isolated to the large motor neurons of the ventral horn (Figure 4.2A) and colocalized with expression of HB9:GFP (Figure 4.2B and C).

As expected certain motor neurons did not express HB9:GFP but did express ChAT-Cre

(Figure 4.2C) (Hinckley et al., 2005; Shneider et al., 2009). No additional RFP expression was found in any other tissue including skeletal muscle, heart, lung, pancreas, kidney, adrenal gland, liver and spleen. Nestin-Cre expression was found throughout the white matter of the spinal cord; however surprisingly, minimal RFP expression was detected in the motor neurons themselves (Figure 4.2D–I). The absence of Nestin-Cre expression is demonstrated by the lack of colocalization of tdTomato-RFP in the motor neuron upon staining with choline acetyltransferase (ChAT) antibody (Figure 4.3).

Additionally I examined the organs of ten different Nestin-Cre mice and identified

122 limited expression in other tissues (Figure 4.4). In all 10 mice, patches of RFP positive blood vessels and connective tissue were found. In skeletal muscle, four of the ten mice studied displayed small patches of weak RFP expression (Figure 4.4A–C). The expression in muscle was very limited in area with only a few fibers showing expression, and this expression was far less intense than expression driven by the Myf5-Cre driver

(Figure 4.4A–C) (Iyer et al., 2015). In six of the ten mice examined skeletal muscle was completely negative for RFP. One of ten mice displayed RFP expression in the heart

(Figure 4.4D–F). Lung was weakly positive for RFP in more than half of the mice examined, but bronchioles were always negative (Figure 4.4G–I). Pancreas expression was found in most animals in the acinar cells, however the islet cells responsible for the endocrine activity of pancreas and the pancreatic duct were always negative in all samples (Figure 4.4J–L). Most kidney sections were positive for RFP in the nephrons, tubules and glomeruli (Figure 4.4M–O). More than half of the mice studied had scattered expression in the medulla of the adrenal gland (Figure 4.4P–R). No RFP expression was detected in the liver or spleen. In summary, Nestin-Cre expression was consistently identified in the spinal cord, blood vessels, acinar cells of the pancreas and kidney nephrons. Most of the ten mice studied had no expression in skeletal muscle, heart, lung, adrenal gland, liver and spleen.

To give an overall representation of the expression pattern of Nestin-Cre and

ChAT-Cre together I visualized tdTomato-RFP at P2 in a whole-body sagittal section

(Figure 4.5). The brain and spinal cord reveal the most intense staining while the connective tissue found at the periphery of the gut and the lungs was weakly positive.

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Most notably however is that the skeletal muscle is completely negative when tdTomato is driven by Nestin-Cre and ChAT-Cre.

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Figure 4.2. Nestin-Cre and ChAT-Cre expression in the spinal cord: Mice containing ChAT-Cre (A-C) or Nestin-Cre (D-I) were crossed to tdTomato RFP reporter line where mice contained HB9:GFP transgene to label the motor neurons (B,E and H). (A-C) ChAT-Cre is heavily expressed in all motor neurons (arrow). Scale bar: 100 µm (D-I) Nestin-Cre is expressed throughout the white matter but very weakly in motor neurons (arrow). (G-I) Higher magnification inset image showin in the white box in images D-F. Representative images from ChAT-Cre mice (n=3) and Nestin-Cre mice (n=10) at P12. Scale bars = 10 µm.

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Figure 4.2

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Figure 4.3. The expression of the Nestin-Cre driver in the spinal cord: (A) Nestin-Cre pattern of expression was determined by crossing mice to the tdTomato-RFP reporter line (red). (B) ChAT staining marks the motor neurons and (C) NeuN staining identifies the nuclei in spinal cord sections of P10 mice. (D) The merged image reveals a lack of Nestin-Cre expression in some motor neurons. The white arrow indicates a hole where tdTomato-RFP is not present. (E-H) Insets from the white boxes in A-D clearly show the absence of RFP in some motor neurons. Scale bars = 100 µm.

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Figure 4.4. Limited expression of Nestin-Cre in several tissues: The Nestin-Cre driver was crossed to tdTomato reporter line to visualize expression of the driver in all tissues. For each tissue RFP expression is show in red (A,D,G,J,M,P), followed by the complementary DIC image (B,E,H,K,N,Q) and the overlaid images (C,F,I,L,O,R). (A- C) Very weak muscle expression was found in 4 of 10 mice. Scattered blood vessels (inset) and some connective tissue are positive for RFP. (D-F) Only one animal had expression in the heart. (G-I) Lung was weakly positive for RFP in more than half of the mice examined, but bronchioles were always negative. (J-L) Pancreas expression was found in most animals in the acinar cells, however the islets and pancreatic duct were always negative in all samples (inset). (M-O) Most kidney sections were positive for RFP in the nephrons, tubules and glomeruli. (P-R) More than half of the mice studied has scattered expression in the medulla of the adrenal gland. No RFP expression was detected in the liver or spleen (data not shown). These images are representative of 10 different P12 mice. Scale bars: 200µm.

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Figure 4.4

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Figure 4.5. Whole body sagittal section of Nestin-Cre + ChAT-Cre tdTomato RFP expression at P2: (A) RFP expression is most intense in the brain (Br) and spinal cord (SC). Additionally, connective tissue (CT) in the gut as well as cells in the periphery of the intestines (Int), lung (Lu) and trachea (Tr) can be identified. (B) A grayscale overlay is shown for orientation. Of note is the complete lack of staining in the skeletal muscle in these mice. These images are representative of 3 animals examined. Scale bar: 1mm.

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4.3.3 Survival and weight of mice are improved upon replacement of Smn in neural tissue

Both survival and weight in the mice were monitored to study the impact of deletion or replacement of Smn. Upon deletion of Smn from the motor neurons we expected to generate an SMA-like phenotype where the mice survive approximately 14 days. Instead it was found that upon deletion of Smn with ChAT-Cre, 100% of the mice survived past 250 days and displayed no apparent SMA-like weakness (n = 7, Figure

4.6A). ChAT-Cre; Smn-deletion mice exhibited a very mild tremor in their hindlimbs when held by the tails. This peculiar feature might not be SMA-specific, and may be an unrelated cerebellar phenotype due to Smn-deletion in all motor neurons throughout the

CNS. Deletion of Smn in most neurons and glia with Nestin-Cre resulted in 79% of mice living longer than 100 days (n=18 total mice) with a tremor of the forepaws and clasp of the hindlimbs at the time of weaning (∼25 days) (Figure 4.6A). Noting that Nestin-Cre had weak expression in the motor neurons we chose to delete Smn with both Nestin-Cre +

ChAT-Cre, to ensure that recombination occurred in all neurons and glia. This resulted in only 50% of the mice surviving past 100 days of age (n = 23, Figure 4.6A). All mice displayed severe tremor and weakness illustrated by a full hindlimb clasp. The mice that did survive past 100 days weighed half as much as control mice, similar to the Nestin-Cre deletion mice (Figure 4.6B). Indeed the phenotype of the mice with deletion of Smn with both Nestin-Cre + ChAT-Cre was clearly severe with significant weight loss and weakness.

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To study the opposite effect of replacing Smn in motor neurons mice containing the SmnINV replacement allele were crossed to ChAT-Cre mice. We found that high expression of SMN in motor neurons alone only increased survival of the mouse from 14 days to a mean of 17.3 ± 2.7 days (n = 9, Figure 4.6C). Replacement of Smn with Nestin-

Cre increased survival to a mean of 30.5 ± 7.0 days (n = 14, Figure 4.6C). The mice weighed 40% less than controls and displayed mild tail necrosis (Figure 4.6D). Finally replacement of Smn with both Nestin-Cre + ChAT-Cre increased survival to a mean of

80.4 ± 21.6 days (n = 16) and now 38% of the mice lived longer than 100 days (Figure

4.6C). Interestingly these mice displayed no weakness or clasping of the hind limbs, and thus it appeared that the primary neuromuscular defects were corrected. Distal limb swelling, slight tail necrosis and occasionally ear necrosis were observed only in mice that survived past 100 days. The weights of the Nestin-Cre + ChAT-Cre mice were also improved over that of the Nestin-Cre Smn-replacement mice, although they were still approximately 50% smaller than the control mice (Figure 4.6D). Thus replacement of

Smn in motor neurons and glia was sufficient to rescue both the SMA phenotype and survival of the mouse.

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Figure 4.6. Survival and weight of SMA mice is improved upon replacement of Smn with neuron specific Cre drivers. (A) Upon deletion of Smn from the motor neurons with ChAT-Cre 100% of the mice survived past 250 days with no SMA like behavioral phenotype (mean survival >250 days, n=7 total mice, p<0.001 vs. SMA). Deletion of Smn in all neurons and glia with Nestin-Cre resulted in 79% of mice living longer than 100 days (mean survival 205.4±22.7 days, n=18 total mice, p<0.001 vs. SMA). Deleting Smn with both Nestin-Cre+ChAT-Cre resulted in only 50% of the mice surviving past 100 days of age (mean survival 140.1±20.5 days, n=23 total mice, p<0.001 vs. SMA). SMA mice lived 14.3±0.7 days, n=16 mice. (B) At 36 weeks of age there was no significant different between the weight of the ChAT-Cre deletion mice and controls (30.5±1.8g vs. 33.3 ±3.7g). Nestin-Cre deletion mice were 19.6±0.5g and Nestin- Cre+ChAT-Cre deletion mice were 16.4±0.7g, which is approximately 50% smaller than control mice. (C) Replacement of Smn with the ChAT-Cre driver resulted in mice that lived 17.3±2.7 days (n=9, p=0.3 vs. SMA). Replacement of Smn with Nestin-Cre further increased survival to a mean of 30.5±7.0 days (n=14, p=0.02 vs. SMA). Finally replacement of Smn with both Nestin-Cre+ChAT-Cre increased survival so that 30% of the mice lived longer than 100 days (mean survival 80.4±21.6 days, n=16 total mice, p<0.0001 vs. SMA). (D) ChAT-Cre replacement mice weighed 5.4±1.0g at P16 while Nestin-Cre replacement mice weighted 4.9 ±5.0g. SMA mice at P16 weighed on average 2.7±0.4g. Nestin-Cre+ChAT-Cre replacement mice that survived past 100 days weighted approximately 50% less than controls at 36 weeks of age.

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Figure 4.6

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4.3.4 Muscle fiber size is decreased upon deletion of Smn in neural tissue

The fiber size of the gastrocnemius muscle in Nestin-Cre and Nestin-Cre + ChAT-

Cre deletion mice was examined at P8 (Figure 4.7A–D) and ChAT-Cre, Nestin-Cre and

Nestin-Cre + ChAT-Cre deletion mice at P21 (Figure 4.7E–H). I found that Nestin-Cre +

ChAT-Cre; Smn-deletion mice possess a fiber size distribution that is very similar to the fiber sizes found in SMNΔ7 SMA mice at the same time point (Figure 4.7B and D). The mean fiber size in Nestin-Cre + ChAT-Cre; Smn-deletion mice was 257.4 ± 2.5 μm2 compared with 211 ± 1 μm2 for SMNΔ7 SMA mice (n = 1500 fibers per group from n =

3 total mice, thus approx. 500 fibers per mouse). At P21 the decreased fiber size is quite dramatic as Nestin-Cre + ChAT-Cre; Smn-deletion mice had a mean fiber size of 302.7 ±

3.2 μm2 as compared with a mean of 835.9 ± 8.8 μm2 for controls (n = 1500 fibers per group) (Figure 4.7H and E). The decrease in fiber size in Nestin-Cre; Smn-deletion mice is not as large (mean 477.6 ± 4.8 μm2, median 447.0 μm2) and ChAT-Cre Smn-deletion mice (mean 559.6 ± 6.1 μm2, median 530.5 μm2) were most similar to control mice at

P21. There is no comparison to SMNΔ7 SMA mice at this time point as SMNΔ7 SMA mice only live 14 days. The decrease in muscle fiber size illustrates that loss of Smn in the neurons and motor neurons results in denervation and subsequent muscle atrophy.

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Figure 4.7. Muscle fiber size upon deletion with Nestin-Cre and ChAT-Cre: The gastrocnemius muscle from (A) control, (B) SMA and (C) Nestin-Cre; Smn-deletion mice and (D) Nestin-Cre+ChAT-Cre; Smn-deletion mice were measured at P8. Myofiber size in control mice at P8 (mean 387.8 ± 3.3 µm2; median 379.5 µm2) is significantly different from that of Nestin-Cre; Smn-deletion mice (mean 277.5 ± 2.3 µm2; median 277 µm2), Nestin-Cre+ChAT-Cre; Smn-deletion mice (mean 257.4 ± 2.5 µm2; median 247.0 µm2, p<0.001). The Nestin-Cre+ChAT-Cre; Smn-deletion mice however have myofiber areas similar to that of SMN∆7 SMA mice at P8 (Mean 210.6 ± 1.9 µm2; median 205.1 µm2). (E-H) The difference in myofiber area is more dramatic at P21 where (E) control mice (mean 835.9 ± 8.8 µm2; median 757.0 µm2) and (F) ChAT-Cre; Smn-deletion mice (mean 559.9 ± 6.1 µm2; median 530.5 µm2) display larger fibers as compared with (G) Nestin- Cre deletion mice (mean 477.6 ± 4.8 µm2; median 447 µm2) and (H) Nestin-Cre+ChAT- Cre Smn deletion mice (mean 302.7±3.2 µm2; median 281.0 µm2, p<0.001). There is no comparison to SMN∆7 SMA mice at this time point because the SMN∆7 SMA mice only live for 14 days. For each group, a total of 1500 fibers were measured from 3 mice (approx. 500 fibers per mouse).

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Figure 4.7

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4.3.5 Percent recombination events in LCM isolated motor neuron as determined by ddPCR

We chose to perform droplet digital PCR (ddPCR, BioRad) to determine the percentage of recombination occurring specifically in motor neurons (Figure 4.8A). Only a small fraction (less than 10%) of total spinal cord extract is comprised of motor neurons. The number of recombination events that occurs for each driver line is of utmost importance to proper interpretation of results because Cre drivers often have mosaic or incomplete expression. We isolated approximately 200 motor neurons with laser capture microdissection (LCM) at P10-P12 (n = 3 per genotype) and performed ddPCR to identify the recombination of the Smn-deletion allele. The number of recombination events was normalized to the total number of motor neurons per reaction as determined by Smn intron 1 primers. I found no recombination in mice that did not contain a Cre driver (0.6 ± 7.4%, n = 3) and complete recombination using the Sox2-Cre ubiquitous driver (104.0 ± 1.1%, n = 3) (Figure 4.8A) (Iyer et al., 2015). Recombination in Nestin-

Cre + ChAT-Cre; Smn-deletion motor neurons was also complete (101.3 ± 1.7%, n = 3) and we determined that half of all motor neurons underwent recombination with just the

Nestin-Cre driver (51.3 ± 8.1%, n = 3, P = 0.004 versus Nestin-Cre + ChAT-Cre). The

ChAT-Cre driver alone resulted in 88.7 ± 5.8%, n = 3, P = 0.004 versus Nestin-Cre. The difference in recombination efficiencies between ChAT-Cre alone and Nestin-Cre +

ChAT-Cre was not statistically significant (P = 0.274), illustrating that the recombination occurring in the motor neuron is dependent on the ChAT-Cre driver.

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4.3.6 Determination of SMN protein levels in total spinal cord and motor neurons

The depletion and replacement of SMN in the spinal cord was analyzed using the

Nestin-Cre + ChAT-Cre drivers. In Nestin-Cre + ChAT-Cre mice that possessed the floxed Smn exon 7 allele, along with an Smn null allele, the SMN levels (4.8 ± 1.1) were significantly reduced from no-Cre controls (17.0 ± 1.0) to SMA levels (6.2 ± 1.1) (Figure

4.8B). The difference between Nestin-Cre + ChAT-Cre replacement of Smn with one reversion allele resulted in complete recovery of SMN levels (22.6 ± 3.9) back to no-Cre control levels (P < 0.01, ANOVA). Representative western blot bands are shown in

Figure 4.9B. In addition the SMN protein levels in just motor neurons, collected by

LCM, were examined using ELISA (Figure 4.9A). We found that Nestin-Cre (922.8 ±

168.9 pg SMN/MN) was not as efficient at removal of Smn exon 7 as was deletion with the Nestin-Cre + ChAT-Cre (689.7 ± 274.8 pg SMN/MN) or Sox2-Cre (469.9 ± 23.2 pg

SMN/MN) drivers. Due to the large variability in this small sample size the changes did not rise to statistical significance.

Finally, the number of gems present in the motor neuron nuclei was measured to assess the amount of SMN protein present in our Smn-deletion lines. We counted the number of gems present in 100 nuclei for each sample (n = 3 in each group) (Figure

4.8C). We found the control with one wild-type Smn allele to have the most gems (237.4

± 13.2 gems/100 nuclei) while SMA sample had the least (2.7 ± 0.9 gems/100 nuclei, P <

0.001 compared to control). The Nestin-Cre; SmnD7/KO line was again not as efficient at deletion of SMN from the motor neurons (67.1 ± 1.4 gems/100 nuclei, P < 0.001) while in the Nestin-Cre + ChAT-Cre; SmnD7/KO sample (4.0 ± 0.6 gems/100 nuclei) we

139 identified a similar number of gems per nuclei as found in the SMA control mice. There was no statistical difference between Nestin-Cre + ChAT-Cre; SmnD7/KO and SMA gem counts. Representative spinal cord images of SMN gems are shown in Figure 4.8D and

E. Thus protein measurements by western blot, ELISA on LCM isolated motor neuron tissue and gem counts all show that the Nestin-Cre driver is not efficient at recombination

(either by deletion or replacement of Smn alleles) in the motor neuron.

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Figure 4.8. Determination of Cre activity to deplete or replace Smn: (A) Percent recombination of SmnF7 in motor neurons as determined by ddPCR. Motor neurons were isolated with LCM from P10-P12 mice. There was no recombination in mice that did not contain a Cre driver (0.6 ± 7.4%, n = 3). The Sox2-Cre ubiquitous driver resulted in recombination in all motor neurons (104.0 ± 1.1%, n = 3) (38). Nestin-Cre + ChAT- Cre drivers also gave complete recombination in all motor neurons (101.3 ± 1.7%, n = 3). The ChAT-Cre driver alone resulted in 88.7 ± 5.8% (n = 3, P = 0.004 versus Nestin-Cre). On average only half of all motor neurons underwent recombination with just the Nestin- Cre driver (51.3 ± 8.1%, n = 3, P = 0.004). The difference in recombination efficiencies between ChAT-Cre alone and Nestin-Cre + ChAT-Cre was not statistically significant (P = 0.274). All values are normalized to two copies of Smn intron 1. Percent recombination values were determined by multiplex ddPCR. (B) SMN protein expression as determined by western blot analysis. The alleles used are depicted on the x axis and amount of SMN relative to tubulin on the y. Note that the amount of SMN protein found upon deletion of Smn with Nestin-Cre + ChAT-Cre is not statistically different from that of the SMA sample (P = 0.782). Conversely the replacement of Smn with Nestin- Cre+ ChAT-Cre is not statistically different from that of the control sample (P = 0.322) (C) The number of SMN positive gems in motor neurons. All groups are statistically different by one-way ANOVA except for SMA versus Nestin-Cre + ChAT-Cre which are not different. Thus gem counts indicate equivalent levels of SMN protein in SMA- and Smn-deletion by Nestin-Cre + ChAT-Cre. (D) Smn+/− spinal cord section revealing many motor neurons with gems. (E) Smn−/− (SMA) sample showing lack of gems in motor neurons. (F) Nestin-Cre Smn-deletion sample shows a reduced number of gems in motor neurons. (G) Nestin-Cre + ChAT-Cre, Smn-deletion motor neurons reveal very few gems, similar to the SMA sample (n = 3 for each group, scale bar = 20 µm).

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Figure 4.8

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Figure 4.9. Motor neuron and spinal cord SMN levels in Cre modified SMN∆7 mice. (A) Mice with one Smn wild type allele and the other allele as a floxed exon7 allele containing SMN2 and SMN∆7 were analyzed. The Cre driver is indicated below each bar. 500-600 motor neurons were isolated by laser capture microdissection and analyzed by ELISA for SMN levels. Note that Nestin-Cre does not reduce SMN levels (922.8±168.9 pg SMN/MN) to the same extent as either Sox2-Cre (469.9±23.2 pg SMN/MN) or Nestin-Cre+ChAT-Cre (689.7±274.8 pg SMN/MN). (B) Western blot of total spinal cord tissues from mice that have either depletion or replacement of SMN. The genotype of mice is as indicated in the image. The protein loading control Tubulin is shown below. Notice the decrease to SMA levels in the Nestin-Cre+ChAT-Cre (4.8±1.1) spinal cord samples and the marked increase of SMN protein when using the Smn reversion allele (22.6±3.9). No-Cre control: 17.0±1.0, SMA: 6.2±1.1 for relative protein levels (p<0.01, ANOVA)

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4.3.7 Electrophysiological studies of the functional output of the motor neuron upon deletion and replacement of Smn

Electrophysiological studies including compound muscle action potential

(CMAP), motor unit number estimation (MUNE) and electromyography (EMG) were recorded from the sciatic-innervated hind limb muscles in mice with targeted Smn- deletion and in SMA mice with targeted SMN restoration. In mice with Smn-deletion, electrophysiological studies were performed at P21 to allow assessment of the impact of reduced SMN levels on motor unit function after completion of pruning of polyneuronal innervation at approximately 2 weeks of age (Arnold et al., 2014; Tapia et al., 2012). The goal was to assess loss of motor unit function in comparison with control animals with a

WT copy of Smn. The findings of these studies demonstrate loss of motor unit function in the form of reduced CMAP amplitudes and reduced MUNE when SMN is reduced. Loss of motor unit function was most prominent in animals with deletion of Smn in motor neurons and glia (ChAT-Cre + Nestin-Cre) (Figure 4.10A and B, Figure 4.11).

Furthermore electromyographic (EMG) findings of fibrillations, which results from repetitive depolarization of single muscle fibers, indicating active denervation of muscle fibers was observed in Nestin-Cre + ChAT-Cre animals. EMG analysis revealed fibrillations in hindlimb muscles of mice with deletion of Smn with Nestin-Cre + ChAT-

Cre and ChAT-Cre but not Nestin-Cre, further supporting the requirement of SMN in the motor neuron (Figure 4.10C) (Willmott et al., 2012). Interestingly, in mice with Nestin-

Cre deletion of Smn, MUNE and CMAP were slightly reduced. This is somewhat surprising as the motor neuron is the driving force behind these measures and

144 recombination in motor neurons upon Nestin-Cre deletion was low (Figure 4.10A). We suggest that a possible reason behind these observations is the role of glia, in particular astrocytes, in supporting the health and function of the motor neuron.

CMAP and MUNE were also performed at P12 in SMA mice with targeted Smn restoration. The earlier time point was chosen to allow comparison to control SMA mice that only live 14 days. Upon replacement of Smn, we found improved CMAP amplitudes and MUNE values with Nestin-Cre + ChAT-Cre and ChAT-Cre replacement of Smn but not with Nestin-Cre (Figure 4.10D and E). These findings are consistent with electrophysiological measurements in mice following the deletion of Smn. Thus CMAP and MUNE demonstrate that elimination of Smn reduces the functional output of the motor unit while replacement of Smn restores the functional output.

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Figure 4.10. Electrophysiological measures of motor unit function following targeted Smn-deletion and restoration. (A) Compound muscle action potential (CMAP) at P21 in mice with Smn-deletion. Compound muscle action potential (CMAP) amplitudes at postnatal day 21 (P21) were reduced in ChAT-Cre Smn-deletion (n = 9, 20.4 ± 1.9, P < 0.05) and Nestin-Cre + ChAT-Cre Smn-deletion (n = 13, 26.1 ± 2.1, P = <0.05) but not Nestin-Cre Smn-deletion mice (n = 14, 35.7 ± 3.7) compared with controls (n = 13, 44.0 ± 2.9 mV). CMAP amplitudes in ChAT-Cre Smn-deletion mice were reduced compared with Nestin-Cre Smn-deletion mice (P < 0.05). (B) Motor unit number estimation (MUNE) at P21 in mice with Smn-deletion. Motor unit number estimates (MUNE) at P21 were reduced in ChAT-Cre Smn-deletion (n = 9, 142 ± 22), Nestin- Cre + ChAT-Cre Smn-deletion (n = 13, 117 ± 15), and Nestin-Cre Smn-deletion mice (n = 14, 179 ± 24) compared with controls (n = 13, 283 ± 15) (P < 0.001). (C) Electromyography of limb muscles of mice with Smn deletion. Fibrillations were noted in both ChAT-Cre Smn-deletion (four of five mice) and Nestin-Cre + ChAT-Cre Smn- deletion mice (six of seven mice) but are absent in Nestin-Cre Smn-deletion mice (zero of six) and control mice (zero of five) (P < 0.05). (D) CMAP amplitudes at P12 in mice with Smn-restoration. CMAP amplitudes at P12 in mice with Nestin-Cre + ChAT-Cre Smn-restoration (n = 9, 23.2 ± 3.3) were similar to control mice (n = 13, 24.8 ± 1.2 mV). Conversely, compared with control mice, CMAP amplitudes were reduced in mice with Nestin-Cre-restoration of Smn (n = 8, 17.3 ± 1.1 mV) and in SMA mice with no Cre (i.e. no Smn-restoration) (n = 12, 17.1 ± 0.8 mV) (P< 0.05). CMAP amplitudes were reduced though does not reach statistical significance in SMA mice with ChAT-Cre- restoration of Smn (n = 14, 18.8 ± 1.2 mV). (E) MUNE at P12 in mice with Smn- restoration. MUNE at P12 in mice with ChAT-Cre Smn-restoration (n = 14, 187 ± 16) and Nestin-Cre + ChAT-Cre Smn-restoration (n = 9, 201 ± 22) are similar to controls (n = 13, 196 ± 16, P = 0.962), but MUNE are reduced in mice with Nestin-Cre-restoration of Smn (n = 8, 118 ± 16) and SMA mice with no Cre (i.e. no Smn-restoration) (119 ± 9) (P < 0.05). Compared with mice with Nestin-Cre-restoration of Smn, MUNE at P12 in mice with ChAT-Cre Smn-restoration (P = 0.042) and Nestin-Cre + ChAT-Cre Smn- restoration (P = 0.024) are increased. Similarly, compared with SMA mice with no Cre, MUNE in mice with ChAT-Cre Smn-restoration (P = 0.031) andNestin-Cre + ChAT-Cre Smn-restoration (P = 0.017) are increased. (*P ≤ 0.05, ***P ≤ 0.001).

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Figure 4.10

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Figure 4.11. Electrophysiological Waveforms: Maximal Compound muscle action potential (CMAP) and 10 superimposed incremental single motor unit action potential (SMUP) responses from each inversion and deletion group. CMAP responses are shown at 10mV per division sensitivity and SMUPs are shown at 0.5 mV per division sensitivity. Sweep speed is shown at 1 ms per division for both CMAP and SMUP.

4.3.8 Other neural Cre drivers used in this study

Additionally, the neural drivers SYN1-iCre, rSyn1-Cre and Gad2-Cre (Table 4.1) were used to eliminate and replace Smn. I found significantly improved survival of SMA mice when Smn was replaced with human Synapsin 1 - iCre (SYN1-iCre). More than half of the mice studied (6 of 11 mice) survived for greater than one year of age with only an

18% decrease in weight (Figure 4.12A and B). However, when I examined the expression pattern of the SYN1-iCre line I found nearly ubiquitous RFP staining

148 throughout the spinal cord, muscle, heart lung, liver, pancreas, spleen, kidney and adrenal gland as well as blood vessels, connective tissue and epithelial cells (n = 3 mice) (Figure

4.13). Thus SYN1-iCre is essentially a ubiquitous driver and we would expect results similar to eliminating Smn with our ubiquitous Sox2-Cre control (Chapter 3) (Iyer et al.,

2015).

Figure 4.12. Survival and weight of mice upon deletion and replacement of Smn with SYN1-iCre. (A) Survival is increased in SMA mice upon replacement of Smn with SYN1-iCre with more than half the mice living past 250 days (mean survival 192.7±28.0 days, n=11 SYN1-iCre, SmnRe/KO, p<0.0001). Elimination of Smn with SYN1-iCre resulted in mice that lived on average 26.3±2.7 days (n=14, SYN1-iCre, SmnD7/KO, p<0.0001). SMA mice lived on average 14.3±0.7 days (n=16). (B) Mice were weighed daily until weaning. At 4 weeks of age mice were then weighted weekly. Replacement of Smn with SYN1-iCre resulted in mice that were 18% smaller than control animals at 36 weeks (24.0 ± 1.5g vs. 29.2 ± 1.3g, p=0.03). Deletion of Smn with SYN1-iCre gave mice that were twice the weight of the SMA control mice at P16 (6.0 ± 0.4g vs. 2.7 ± 0.4g, p<0.0001).

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Figure 4.13. Ubiquitous RFP expression of SYN1-iCre: The SYN1-iCre driver was crossed to the tdTomato RFP reporter line to visualize Cre expression. Mice also contained the HB9:GFP transgene to visualize motor neurons. (A-C) Lumbar spinal cord sections reveal expression in the motor neurons (arrow) and throughout the white matter. (D-K) RFP expression is found in all tissues examined including (D) muscle, (E) heart, (F) lung, (G) liver, (H) pancreas, (I) spleen, (J) kidney, and (K) adrenal gland. Blood vessels and connective tissue were positive in all organs. (L) A whole body sagittal section at P2 illustrates ubiquitous RFP expression throughout the mouse. These images are representative of 3 animals examined. A-C, Scale bars: 10 µm, D-K Scale bars: 50µm, L, Scale bar: 1mm.

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Figure 4.13

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When I examined the expression pattern of rat Synapsin1-Cre (rSyn1-Cre), another neuronal specific Cre driver, with tdTomato-RFP I found less than 50% of the mice that were positive for the rSyn1-Cre transgenic insertion by genotyping had any

RFP staining. It appears that some mice carrying the rSyn1-Cre transgene, confirmed with multiple sets of genotyping primers from both the rSyn1 promoter and Cre, were not expressing the driver. Additionally, when rSyn1-Cre was used to eliminate Smn (SmnD7), only four of eight mice displayed reduced survival and an SMA-like phenotype with a mean survival of 25.5 ± 12.5 days (Set1, light blue, n=4, Figure 4.14A and B). The other four mice (Set2, dark blue, n=4, Figure 4.14A and B) displayed no phenotype and no change in survival. Upon crossing rSyn1-Cre with SmnINV, none of the ten affected mice showed rescue in phenotype and survived on average 14.6 ± 1.8 days, similar to SMA mice (n=10, red, Figure 4.14A and B). Furthermore, I crossed the ChAT-Cre line into the rSyn1-Cre to obtain ChAT+rSyn1–Cre. ChAT+rSyn1 – Cre; SmnD7/KO mice appeared similar to ChAT-Cre; SmnD7/KO mice with no decrease in survival (n = 4, Figure 4.14A).

ChAT+rSyn1 – Cre; Smn-replacement mice showed no improvement and were similar to

SMA mice (Figure 4.14A, orange line). Thus the rSyn1-Cre driver was not pursued. It is possible that some of the rSyn1-Cre we were testing had undergone some sort of recombination and no longer expressed Cre. Thus it is paramount to test each Cre line with a reporter gene to ensure accurate interpretation of results. Finally, the glutamic acid decarboxylase 2 - Cre driver (Gad2-Cre) was investigated. When Smn expression was eliminated with Gad2-Cre no phenotype was generated (n = 7, Figure 4.14C and D, blue). Conversely, when replacing Smn expression with Gad2-Cre the mice developed an

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SMA-like phenotype and died on average at 14.5 ± 2.9 days (n = 4, Figure 4.14C and D, red) exactly like SMNΔ7 SMA mice. Hence the Gad2-Cre line was not pursued either.

Figure 4.14. Survival and weight of mice upon deletion and replacement of Smn with (A and B) rSyn1-Cre and (C and D) Gad2-Cre. (A and B) Upon deletion of Smn with rSyn1-Cre, two sets of data were obtained, Set1 (light blue, n = 4) which had a functional Cre, and Set2 (dark blue, n = 4) which apparently had a non-functional Cre. rSyn1-Cre is thus a chimeric line. Set1-deletion mice decrease in weight and show a drop in survival to a mean of 25.5 days while Set2 mice were normal. rSyn1-Cre; Smn-replacement mice (red, n = 10) were no different from SMA mice, surviving to a mean of 14.6 days. ChAT- Cre + rSyn1-Cre did not help the situation – the deletion mice (purple) appeared normal and the replacement mice (orange) were SMA-like. (C and D) Gad2-Cre; Smn-deletion (blue, n = 7) mice showed no weakness whatsoever and had a normal survival and weight. Gad2-Cre with the replacement allele failed to rescue the SMA-phenotype (red, n = 4). 153

4.4 DISCUSSION

While it is clear that SMN reduction causes SMA and that the amount of SMN is critical in determining phenotypic severity, SMN levels are predicted to be decreased in all tissues. It remains unclear where high levels of SMN (above that produced by two copies of SMN2) are required, and whether this varies in different severities of SMA. The current study is the first to use Cre recombinase to both delete and replace Smn using the same Cre driver to study the effects of reduced or restored levels of SMN in glia, motor neurons, autonomic neurons and other neurons in the motor circuit. Furthermore, we have performed a complete characterization of the expression patterns of the Cre drivers used to understand the specificity of drivers, or lack thereof, to allow full interpretation of the effects on weight, survival and motor unit function in the SMNΔ7 mouse.

Prior studies used behavioral analysis, physiological NMJ patch clamp recordings of single synapses ex vivo or morphological assessment of NMJs and motor neurons to indirectly draw conclusions about motor unit function (Lee et al., 2012; Martinez et al.,

2012; Taylor et al., 2013). In contrast, the present study uses electrophysiological techniques (CMAP, MUNE and EMG) to assess motor unit number and function in vivo, similar to techniques that have shown abnormalities in clinical studies of patients with

SMA (Hausmanowa-Petrusewicz and Karwanska, 1986; Swoboda et al., 2005). NMJ patch clamp recordings in the SMNΔ7 mouse have revealed transmission abnormalities of reduced evoked endplate current amplitudes (Kong et al., 2009). These patch clamp

NMJ abnormalities are corrected with both systemic SMN restoration and with SMN restoration in the motor neuron but not with restoration in muscle (Foust et al., 2010;

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Martinez et al., 2012; Paez-Colasante et al., 2013). Though motor deficits have been noted in mice with selectively reduced levels of SMN in motor neurons, NMJ patch clamp recordings have not been performed (Park et al., 2010). For obvious reasons these measures have not been performed in human SMA. However abnormalities on CMAP,

MUNE and EMG are key features in patients with SMA (Hausmanowa-Petrusewicz and

Karwanska, 1986; Swoboda et al., 2005). Prior to the availability of molecular diagnostic testing, EMG was an important diagnostic tool as the presence of fibrillations is a marker of denervation and a typical feature of type 1 SMA, and today it still has utility in atypical cases (Arnold and Burghes, 2013; Arnold et al., 2014). CMAP and MUNE are particularly useful to quantify motor neuron or axon dysfunction, and these measures have been shown to correlate with disease severity and function in several cohorts of patients with SMA (Hausmanowa-Petrusewicz and Karwanska, 1986; Kang et al., 2014;

Kaufmann et al., 2012; Lewelt et al., 2010; Swoboda et al., 2005).

Our laboratory developed CMAP, MUNE and EMG methods for recording even in neonatal mice (Arnold et al., 2014). We used CMAP, MUNE and EMG in the present studies due to the clinical relevance and for the key ability to analyze the functional output and connectivity of the motor neuron in vivo. We have previously shown that these measures are responsive to restoration of SMN protein via intracerebroventricular injection of antisense oligonucleotide to ISS-N1 and scAAV9-SMN in the SMNΔ7 mouse and scAAV9-SMN in a pig model of SMA (Arnold et al., 2014; Duque et al.,

2015). Importantly, our results reveal that decreasing and restoring the level of SMN in the motor neuron is sufficient to both cause and prevent the losses of motor unit function

155 that are characteristic of human SMA. Our results support that SMN level in the motor neuron is central to the pathogenesis of motor neuron loss and that restoration of SMN levels in the motor neuron will be critical for therapeutic implementation in patients with

SMA. While expression of high levels of SMN in just motor neurons does not rescue survival of the mouse it does rescue the function of the motor neuron. In human SMA improving the function of the motor neuron is likely to be critical in effective therapy.

The mouse has other affected organs such as the heart upon low SMN levels, however additional organ involvement does not appear to be a prevalent feature of SMA in man

(Bevan et al., 2010; Heier et al., 2010; Iascone et al., 2015; Shababi et al., 2010).

The clinical and pathological features of human SMA are remarkably restricted to the neuromuscular system (Arnold et al., 2014; Crawford and Pardo, 1996; Harding et al.,

2015; Monani, 2005). Yet, some studies have indicated the potential importance of SMN in the periphery (Hamilton and Gillingwater, 2013; Shababi et al., 2014). In mouse models of SMA, various non-motor features have been noted, namely distal limb necrosis, endocrine abnormalities, dysautonomia, cardiac defects and gastrointestinal abnormalities (Bevan et al., 2010; Hsieh-Li et al., 2000; Porensky et al., 2012; Schreml et al., 2013). It has been suggested that the autonomic nervous system may play a role in some of these phenotypic features (Gombash et al., 2015; Le et al., 2011; Porensky et al.,

2012). For instance, do alterations in the vascular system and the heart in mice relate to the autonomic nervous system? Though some similar atypical features in patients with

SMA have very rarely been presented in the literature, these single case reports are restricted to most severe form of SMA, type 0 and 1a, with onset at or before birth (Arai

156 et al., 2005; Arnold et al., 2014; Bowerman et al., 2012a; Distefano et al., 1994; Rudnik-

Schoneborn et al., 2010; Rudnik-Schöneborn et al., 2008). Thus whether these reports relate to the more typical type 1 SMA situation remains questionable.

In a recent study by Hua et al., the importance of peripheral SMN restoration was proposed (Hua et al., 2015). In this study, a different model, the Taiwanese SMA mouse, was used which has a deletion of mouse Smn exon7 and either 2 copies of SMN2 (for a severe phenotype) or 4 copies of SMN2 (for a milder phenotype) (Hsieh-Li et al., 2000;

Hua et al., 2015). The mouse locus in this model can still produce SMN transcript lacking exon7. Splice-correcting ASO was administered via subcutaneous injection in severe

Taiwanese mice dramatically extending survival similar to prior studies (Hua et al., 2015;

Hua et al., 2011). Importantly in the neonatal period, ASO can cross the blood brain barrier and exert partial correction of splicing to SMN2. Therefore to limit the induction of exon 7 the authors included a blocking ASO that was administered to the CNS via intracerebral ventricular injection (Hua et al., 2015). Yet, despite delivery of the blocking

ASO some increase in SMN exon7 inclusion occurred in the spinal cord. Indeed we have previously shown in the SMNΔ7 mouse model that even a small increase in leaky expression from the human skeletal actin muscle promoter HSA-SMN transgene elicited a major impact on phenotype (Gavrilina et al., 2008). Therefore it is important to consider how the phenotypes of the SMNΔ7 and the severe Taiwanese mice (with two copies of SMN2) compare. While the survival of the SMNΔ7 and severe Taiwanese models are similar (2 weeks), the variability between the two models regarding the involvement of the motor system compared with other organ systems is significant

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(Bevan et al., 2011; Hsieh-Li et al., 2000; Le et al., 2005; Porensky et al., 2012; Schreml et al., 2013). Indeed Hua et al. reported that the longissimus capitis muscle in the severe

Taiwanese did not show any completely denervated NMJs (Hua et al., 2015). In contrast, the same muscle in the SMNΔ7 mouse shows partial-to-complete denervation of 80% of synapses and complete denervation in 28% of synapses (Ling et al., 2012). Given the weaker denervation phenotype in the severe Taiwanese mice, a small increase of SMN in the CNS may be sufficient to overcome the CNS requirement. Furthermore, it is difficult to interpret the survival outcome of the Taiwanese mouse given the severity of dysfunction in several organ systems (Bevan et al., 2011; Hsieh-Li et al., 2000; Le et al.,

2005; Porensky et al., 2012; Schreml et al., 2013). Iascone et al. have commented that using denervation as a quantitative readout for SMNΔ7 mice and comparing it to post- mortem samples from type 1 patients on a muscle-by-muscle basis they have shown a remarkable overlap between the mouse and human neuromuscular phenotype (Iascone et al., 2015). This indicates that the SMNΔ7 mouse model recapitulates the neuromuscular phenotype of patients. The SMNΔ7 SMA mouse does have defects in the heart, yet this defect does not appear to match human patient studies (Bevan et al., 2010; Heier et al.,

2010; Iascone et al., 2015; Shababi et al., 2010).

It is critical to consider whether key phenotypic features that are relevant to human SMA, such as motor neuron loss and denervation, are rescued when assessing where high levels of SMN are required. Our study shows that the effects of SMN level on weight and survival were decoupled from the effects on electrophysiological motor unit function. Despite significant impact on motor unit function following deletion or

158 restoration of SMN levels in the motor neuron, the reduction or improvement of survival and weight was not dramatic unless SMN was reduced or restored in motor neurons plus neurons and glia (with ChAT-Cre + Nestin-Cre). Though both weight and survival are well established and important preclinical endpoints and readouts of sufficiency of SMN restoration, these parameters do not appear to be directly related to the functional status of the motor unit. First, we have found that replacement of SMN in just motor neurons does not correct all phenotypic features in the SMNΔ7 mouse, but it does completely correct the electrophysiological output of the motor neuron. Conversely, upon decreasing

SMN in motor neurons solely there is no impact on weight or survival, but there is a significant decrease in CMAP and MUNE. Second, expression in neurons and glia (with

Nestin-Cre alone) has a considerable impact on survival but less so on motor unit function as indicated by the CMAP and MUNE read out. When SMN was reduced to

SMA levels with Nestin-Cre, which is expressed in all (most) neurons and glia but not efficiently in motor neurons, no fibrillations were detected on EMG whereas with ChAT-

Cre, clear fibrillations are observed indicating denervation. The combination of ChAT-

Cre + Nestin-Cre results in animals that either had the most severe phenotype or the most complete rescue. In fact, upon rescue some animals showed a normal life span indicating that the tissues where SMN was expressed at high levels was sufficient for normal function. In this condition (ChAT-Cre + Nestin-Cre) SMN is expressed in the autonomic nervous system. It has been previously shown that the SMNΔ7 mice have a marked cardiac phenotype, which is partially corrected by delivery of SMN postnatally with scAAV9-SMN (Bevan et al., 2010; Heier et al., 2010; Porensky et al., 2012; Shababi et

159 al., 2010). This results in transduction of the autonomic neurons that innervate the heart.

In the current study Nestin-Cre is likely to result in the restoration or depletion of SMN from most neurons including the autonomic neurons. The cardiac phenotype is important in the mice and certainly one possible reason for improved survival when using ChAT-

Cre + Nestin-Cre for rescue of SMA mice. However, as indicated by Iascone et al., the cardiac defects are not a common feature of SMA in patients and as such the mouse model does not completely mimic the human situation (Iascone et al., 2015). Indeed they concluded that multi-organ dysfunction, including cardiac and vascular defects, is not a general feature of SMA. In addition the motor neuron circuit has been found to be defective in SMNΔ7 SMA mice and this will also be corrected by the Nestin-Cre driver.

Lorson et al. reported that expression of high levels of SMN in astrocytes benefited SMA mice (Lorson et al., 2010; Mentis et al., 2011). The Nestin-Cre driver is very well expressed in astrocytes (Tronche et al., 1999). Thus the changes in the mouse with the

ChAT-Cre + Nestin-Cre drivers can result from expression of SMN in glia and all neurons as well as motor neurons (Tronche et al., 1999).

While we have shown some patchy expression of Nestin-Cre in the periphery, we feel this limited expression is unlikely to be enough to correct the function of these tissues. Instead, others have shown in human SMA-derived iPS cells differentiated into astrocytes that Glial Fibrillary Protein (GFAP) is increased and astrocyte morphology is altered (McGivern et al., 2013). It is quite likely that the extent of glial involvement in the SMA phenotype is still not fully understood. However it is possible that a secreted protein from one of these tissues could also impact the phenotype.

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SMN has a well-established canonical function in the assembly of RNP complexes. SMN is required for the assembly of Sm proteins onto snRNA and assembly of the Lsm10-11-Sm protein ring onto U7 snRNA. SMN has also been suggested to have a role in the assembly of other RNA complexes such as those that function in mRNA transport (Hubers et al., 2011; Li et al., 2014). Since SMN is known to function in splicing, it is quite possible that the alteration of splicing of particular genes could be affected by reduction of SMN in the motor neuron. While abnormal mRNA spliced forms have been reported it is clear that the majority of genes are spliced normally in SMA mice (Baumer et al., 2009; Zhang et al., 2013b). Given the other organ involvement in

SMA mice in addition to the replication of the neuromuscular phenotype, one wonders if the genes affected by SMN deficiency have the same sensitivity to depletion of SMN due to variance in the intron sequence and structure between species (Wang et al., 2015; Xie,

2014). This has been demonstrated in the case of resistin (Ghosh et al., 2003). Thus in humans, some introns may be more sensitive to SMN deficiency than others. It is possible that in the mouse SMN must be reduced to a greater extent to uncover the motor phenotype, resulting in the dysfunction of additional genes that give rise to other organ involvement. If this is the case in humans remains to be determined. The benefit from using both ChAT-Cre and Nestin-Cre together could be due to correction of these other genes with a less pronounced effect on a gene that is sensitive to SMA deficiency. The newly developed pig model of SMA will be important in discerning intron sensitivity upon reduced SMN levels among species (Duque et al., 2015).

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CHAPTER 5

CONCLUSIONS AND FUTURE DIRECTIONS

In this dissertation, we investigated the spatial requirement of the SMN protein in the neurons and muscle in SMA. Upon deletion of muscle ubiquitin ligases, MAFbx and

MuRF1, in SMA mice we found no beneficial effect on the phenotype. The improvement in SMA phenotype with histone deacetylase (HDAC) inhibitors has been attributed to the decrease in MAFbx and MuRF1. In Chapter 2, we rule out that possibility entirely.

Though MAFbx deletion improves the fiber size distribution in SMA mice, it does not alter the weight or survival. Interestingly we uncovered a detrimental phenotype in SMA mice lacking MuRF1. Thus it suffices to say that the HDAC inhibitors like Trichostatin A do not function by downregulation of the muscle ubiquitin ligases in SMA. The systematic deletion and restoration of SMN in muscle, with SMN2 and SMN∆7 in the background, in Chapter 3 shows that low SMN levels do not seem to affect the muscle force production capacity per se. Hence it is unlikely that administering SMN-increasing therapies to muscle alone would be of considerable benefit to SMA patients.

In Chapter 4, we worked with many neuronal Cre drivers; of the ones we worked with, none were pan-neuronal and non-leaky. Nestin-Cre mice have been widely used since their creation and the Cre driver has been called a pan-neuronal one. However we

162 found that Nestin-Cre has poor expression in the spinal cord motor neurons. We thoroughly characterized the Nestin-Cre transgenic line, not just with respect to the spinal cord but in all tissues and found Nestin-Cre to be very chimeric, with varied expression within the same litter. It is possible that the line lost its robustness over years. We found the rat Synapsin1-Cre line to be chimeric too. Additionally, the human Synapsin1-iCre line which is supposedly pan-neuronal turned out to be ubiquitous. Our studies show that is it very important to rigorously characterize Cre lines, especially transgenic Cre lines, before reaching to conclusions of a study. It is our opinion that a reliable and foolproof strategy to create a Cre line is to engineer the Cre with an IRES (internal ribosome entry site) after the endogenous driver gene, as in the case of ChAT-Cre. Moreover an improved Cre (iCre) that has an improved codon usage for eukaryotic systems which is more efficient than the regular prokaryotic Cre (e.g. human Synapsin1-iCre) could be used. Towards the end of our studies, Jackson Laboratories reported the creation of a neuronal Cre line, SNAP25-IRES2-Cre-D, which is targeted insertion of Cre following an IRES sequence under the endogenous synaptosomal associated protein 25, SNAP25

[Jax Stock No: 023525; Reference: https://www.jax.org/strain/023525]. SNAP25 is expressed pan-neuronally, and hence SNAP25-IRES-Cre should be a good pan-neuronal

Cre driver.

In aiming to look at the Cre reporter RFP expression in the whole body, we devised a novel method for whole body cryostat sectioning of neonatal mouse pups

(Methods, Chapter 4). The trick was infusing the body cavities of the PBS-perfused pups

163 with molten agarose solution; this provided the needed physical support for the internal tissues facilitating easy sectioning.

Another novel protocol of the dissertation is the development of an effective cell lysis method for laser-capture microdissected (LCM) tissue, with the aim of performing droplet digital PCR (ddPCR) (Methods, Chapters 3 and 4). I had two hurdles here; with

LCM we could obtain only small quantities of starting material and secondly to perform ddPCR we could not use usual concentrations of detergent for lysis. Higher concentrations of detergent lyse the nanodroplets in ddPCR. After many trials, I obtained an optimum cocktail of cell lysis solution that effectively released DNA from the small amounts of LCM material and did not disrupt the droplet formation in ddPCR.

Additionally I optimized hematoxylin and eosin staining for PEN-membrane coated LCM slides (Methods, Chapter 3). With the cell lysis solution, we can exploit the advantages the techniques of LCM and ddPCR have to offer.

Study with Nestin-Cre and ChAT-Cre drivers in Chapter 4 illuminate the importance of the motor neurons in SMA. Restoration of SMN levels in motor neurons

(ChAT-Cre) rescued the electrophysiological output of the motor unit as indicated by

CMAP (compound motor action potential) and MUNE (motor unit number estimate).

CMAP and MUNE are non-invasive techniques routinely performed on patients that give a read out on the functional status and number of the motor units in a muscle. However,

Nestin-Cre together with ChAT-Cre had the maximum benefit – rescue of survival and electrophysiology. One possibility could be that Nestin-Cre effectively targets the autonomic nervous system (ANS) and the glial cells, the relevance of both in SMA 164 remains to be proven. Another possibility is that 2 copies of the SMN2 transgene in the mouse model do not provide sufficient low levels of SMN for the ANS in the mouse; thus uncovering other phenotypes like cardiac defects and tail necrosis in SMA mice that are rescued. In conclusion, an SMN-enhancing therapy should aim to increase SMN levels in the motor neurons in particular, and not the skeletal muscle, for maximum improvement of phenotype.

The spatial requirement of SMN thus established I am working on assembling the

SMN complex components in yeast, Saccharomyces cerevisiae. S. cerevisiae is the only known eukaryotic model organism that lacks SMN. Therefore it serves as an excellent model to assemble the entire SMN complex, without influence of any endogenous SMN. snRNP assembly assays can be performed without any endogenous SMN interfering in the cell extracts. The project aims to clone SMN, Gemins2-8 and UNRIP under yeast promoters into the yeast genome by homologous recombination. Firstly, no ATP- dependent assembly of the SMN complex has not been demonstrated in vitro.

Assembling the SMN complex thus in yeast can potentially be used to show ATP- dependence of the snRNP assembly reaction. Secondly, missense SMN mutations and synthetic mutations in gemins can be done in the aforementioned yeast system without the need for WT SMN. This mutated SMN complex can be analysed for its capacity to assemble Sm proteins onto snRNA. Synthetic mutations in Sm proteins can be done to study the complementation of function of the mutated SMN or Gemin proteins. These assays will offer invaluable information on the functional domains and interactions of the

SMN complex components.

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Studies so far have established that early intervention is likely to produce the maximum recovery of motor function in an SMA patient. Unless a family has a history of

SMA, prenatal diagnosis is unlikely. If the current clinical trials in SMA show successful outcomes, prenatal screening may be on the horizon. Till then the fact is majority patients arrive at the clinic after onset of symptoms, suggesting considerable loss of motor function has occurred. This holds true for the SMA children enrolled in the ongoing clinical trials. Therefore as clinical trials in SMA advance, a crucial point to consider is how much of the lost motor function can be restored or stalled with additional SMN. A good idea would be to increase the functional output of the existing motor units. Most therapies targeted towards the muscle focus on delaying atrophy or maintaining the muscle mass. An alternative worth exploring is increasing the functional capacity of the existing muscle fibers. Elaborating, making the existing muscle fibers produce more output with the same nerve input is an attractive idea as an adjunct therapy in SMA.

Another strategy could be enhancing the function of the existing motor neurons. The existing motor neurons sprout and reinnervate the muscle fibers that have lost their motor neurons. Administering drugs that increase the sprouting of the existing motor neurons, along with increasing FL-SMN, could increase the overall functional output of the motor unit. For an SMA patient, even a modest increase in motor function may be the added ability to do a routine activity, for example being able to hold a pencil. Well, with multiple ongoing clinical trials, it is an exciting time to be in SMA research.

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