Motor and Sensory Characterization of a Mouse Model of

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Motor and Sensory Characterization of a Mouse Model of Motor and Sensory Characterization of a Mouse Model of

Charcot-Marie-Tooth Type 2O Disease Charcot-Marie-Tooth Type 2O Disease

Swaran Nandini University of Central Florida

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MOTOR AND SENSORY CHARACTERIZATION OF A MOUSE MODEL OF CHARCOT-MARIE-TOOTH TYPE 2O DISEASE

by

SWARAN NANDINI M.S, University of Central Florida, 2013

B. Tech., Jaypee Institute of Technology (India), 2011

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy

in the Burnett School of Biomedical Sciences in the College of Medicine

at the University of Central Florida Orlando, Florida

Spring Term 2019

Major Professor: Stephen J. King

ii

©2019 SWARAN NANDINI

iii

ABSTRACT

Dynein is an essential motor protein required for the maintenance and survival of

cells. Dynein forms a motor complex to carry intracellular cargoes like organelles,

growth factors, peptides, and hormones along the microtubules inside the cells.

In neurons, the dynein is the retrograde motor protein that moves cargoes from

the neuronal tip to the neuronal soma along the length of an axon. Dynein has an

established role in neuronal nuclear migration, transport of neuronal survival

signals and growth factors, organelle positioning inside neurons etc. Hence, it is

not very surprising that numerous mutations in dynein have been reported in

association with neurodegenerative diseases in humans. The first human

mutation (H306R) in dynein heavy chain was reported to cause Charcot-Marie-

Tooth Type 2O disease (CMT2O) in humans. CMT2O patients display motor-

sensory neuropathy symptoms such as muscle weaknesses and wasting in legs,

skeletal deformities like pes cavus (high foot arching), difficulty in walking, and a

loss of sensation.

We developed a novel knock-in H304R mouse model with the corresponding

CMT2O linked dynein mutation to understand the disease’s molecular

mechanism. We investigated and characterized the motor-sensory phenotype of

the H304R mouse model (wildtype, heterozygous (H304R/+) and homozygous

(H304R/R) mice). First, we started with testing mice on motor skills behavior

tests such as tail suspension reflex, grip strength test, and rotarod test at 3, 6, 9

iv

and 12 months of age. Both male and female groups of heterozygous (H304R/+)

mice displayed mild defects in tail suspension reflex, grip strength, and rotarod

performance. In contrast, homozygous (H304R/R) mice exhibited severe defects

in the tail suspension reflex, grip strength, and rotarod performance right from an

early age. Next, I analyzed the sensory phenotype of the H304R mouse model.

Homozygous H304R/R mice appeared to have thinner sciatic nerves, reduced

total fascicular area of the sciatic nerve, and significantly quicker latency to tail

withdrawal from a pain stimulus than the wildtype and heterozygous H304R/+

mice.

Collectively, our motor and sensory characterization studies reveal that H304R

dynein mouse model recapitulates many of the phenotypes associated with CMT

symptoms. Hence, the H304R model is a useful tool in understanding the dynein

function in the onset and progression of CMT2O in humans.

v

I dedicate my PhD dissertation work to my grandparents, in honor of their

sacrifices, struggles, and hard work that afforded my family the privilege to

pursue our dreams.

vi

ACKNOWLEDGMENTS

First and foremost, I thank my PhD advisor, Dr. Stephen J. King for providing me

the opportunity to pursue my doctoral research in his lab. I specially thank him for

helping me develop my critical thinking and effective communication skills. His

mentorship has built me professionally both inside and outside the lab. Because

of his open-minded support, I was able to collaborate with other scientists and

learn many valuable scientific techniques and skills. In addition, I am also very

grateful to Dr. King for letting me be independent in pursuing my interests and

that has allowed me to build a strong leadership profile outside the lab. I also

thank him for creating a very welcoming and understanding environment in the

lab.

Next, I am very grateful to both the past and present members on my PhD

Dissertation Committee. Special thanks to Dr. Yoon-Seong Kim who has been a

long-term committee member for my M.S. thesis (2011-2013) and PhD

Dissertation (2013-2019) committees. Thank you, Dr. Kim, for showing me the

importance of thinking outside the box when looking for possible scientific

solutions. Special thanks to Dr. Annette Khaled for being a strong female role

model for me, both in science and in leadership roles. Thank you, Dr. Khaled, for

your direction and support for past so many years. Next, I want to thank

Dr. Stephen Lambert for being the most authentic person and well-rounded

scientist I know. Thank you, Dr. Lambert, for showing me the power of being

vii

passionately true to ourselves. Also, thank you for allowing me to learn the

specialized skills of mouse surgeries in your lab (thanks to Dr. Dale George and

Dr. Wesley Anderson). Last and of course not the least; I would like to thank Dr.

Alvaro Estevez who was a former M.S. and PhD committee member. Thank you,

Dr. Estevez for always having your office and lab door open for me walk in and

seek your guidance while you were at UCF. Thank you, Dr. Estevez, for always

believing in me. In addition, special thanks to Dr. Maria Clara Franco as well for

teaching me some novel techniques (mitochondrial isolation and oxygen

consumption assays) and the importance of designing in-depth experiments in

order to perform thorough science. Also, big thanks to Dr. Raheleh Ahangari for

being such a wonderful and kind advisor for my graduate teaching

assistantships. Thank you to all the above-mentioned scientists for challenging

me to be a well-trained PhD graduate student.

Thanks to the past and current members of King lab that made the environment

so collaborative and fun – Dr. Linda King, Rachal Love, Jami Conley Calderon,

Dr. Thywill Sabblah, Aaron Ledray, Julio Pasos, Alexis Ghersi, Rochelle Sadeghi,

Bryce Ordway, Keira Kraus, Emily Silva, Gabi Fiorino and Phil Hannoush. In

addition, special thanks to the former and current Burnett School of Biomedical

Sciences (BSBS) Program Leaders – Dr. Sampath Parthasarathy, Dr. Griffith

Parks, Dr. Saleh Naser, Dr. Jihe Zhao and Mr. Greg Norris for their support in

every step of the way of my graduate career. Many thanks to Lisa Vaughn,

viii

Allison Connally, Ka Yam, and Shannon Connally for being the strong backbone

of the BSBS PhD program.

Also special thanks to the former and current Lake Nona Staff members for their

efforts in making our lives easier so that we can solely focus on research – Lisa

Simcoe, David Frosch, Amy Postlewait, Jeanette Galloway, Abby Snipes, and

Susie Nisavic; the BSBS custodial staff (Ms. Lucy Bautista); the BSBS IT

(Robinson Pamplano and Alex Lazin); and the BSBS Engineering (Carol

Lanouette, Anthony Smith, and Joseph Myerson). And of course, special thanks

to the former and current BSBS Vivarium Staff members and Teri Krisch and

James Grant for making it so convenient for me to complete my experiments in

mice for past so many years. I also thank my peers in the BSBS program for

being so friendly and supportive of each other.

Serving multiple leadership roles in and outside of UCF has been a personal

career highlight for me. I am very thankful to all the leadership staff members I

had the opportunity to work with on the main campus. Moreover, I specially thank

them for always being so understanding and adaptable to my lab schedule and

letting me work on the leadership projects at my own schedule. Special thanks to

the UCF Recreation and Wellness Center (Mr. James Wilkening and the team),

UCF Internationalization Committee (Ms. Chantel Carter and the committee),

UCF Technology Fee Committee (Dr. Joel Hartman and the committee) for

wholeheartedly providing me with many valuable leadership opportunities for

past many years.

ix

Lastly, I want to thank my family and loved ones for being so selflessly patient

and understanding through the intense journey of the graduate school. Thank

you to all my friends who have shown me their unconditional support through

their actions, time and again. Last but not the least; deep heartfelt thanks to my

patrons (UCF students) that came to my group exercises classes every week for

past so many years. Their love, support, and energy are what kept me

encouraged and positive through this long journey of earning my doctoral degree.

x

TABLE OF CONTENTS

LIST OF FIGURES ..............................................................................................xv

LIST OF TABLES ............................................................................................. xviii

LIST OF ABBREVIATIONS .................................................................................xx

CHAPTER 1: INTRODUCTION ............................................................................ 1

1.1 Cytoplasmic Dynein Structure ................................................................. 3

1.1.1 Cytoplasmic Dynein Heavy Chain Structure ..................................... 4

1.2 Cytoplasmic Dynein Function .................................................................. 6

1.2.1 Dynein interactions with its binding partners ................................... 11

1.3 Charcot-Marie-Tooth Diseases in Humans ............................................ 20

1.3.1 Dynein mutation linked to CMT2O and SMA-LED .......................... 23

1.4 Mouse Models with Neurodegeneration Linked Dynein Mutations ........ 25

1.5 Peripheral Nervous System of Mouse ................................................... 30

1.5.1 Role of dynein mediated retrograde signaling in sensory neurons . 31

1.5.2 Sciatic Nerve anatomy in mice ........................................................ 35

1.5.3 Dorsal Root Ganglia anatomy in mice ............................................. 37

CHAPTER 2: MATERIALS AND METHODS ...................................................... 40

2.1 Generation of H304R mouse model ...................................................... 40

2.2 Mouse colony maintenance ................................................................... 42

xi

2.2.1 Mouse colony housing .................................................................... 42

2.2.2 Mouse colony breeding ................................................................... 42

2.2.3 Measurement of mouse body weight .............................................. 43

2.2.4 Mouse health supervision ............................................................... 43

2.3 Behavioral phenotype characterization test ........................................... 44

2.3.1 Tail suspension test ........................................................................ 44

2.3.2 Grip strength test ............................................................................ 46

2.3.3 Rotarod performance test ............................................................... 47

2.3.4 Tail flick test .................................................................................... 49

2.4 Mouse dissections (sciatic nerve and dorsal root ganglia) .................... 50

2.4.1 Tissue harvesting and cryopreservation ......................................... 50

2.4.2 Cryosectioning and immunohistochemistry of tissue samples ........ 50

2.4.3 Microscopy and data analysis ......................................................... 53

2.5 Motility analysis studies in primary H304R neurons .............................. 54

2.5.1 Tissue harvesting of embryonic primary neurons............................ 54

2.5.2 Live neuronal imaging with Rab7-GFP labeled vesicles ................. 55

2.5.3 Live neuronal imaging with Synaptophysin-RFP labeled vesicles .. 58

CHAPTER 3: BEHAVIORAL PHENOTYPE CHARACTERIZATIONS OF H304R

MICE .................................................................................................................. 59

xii

3.1 Introduction ............................................................................................ 59

3.2 Rationale ............................................................................................... 61

3.3 Results .................................................................................................. 64

3.3.1 H304R mice had normal life span ................................................... 64

3.3.2 H304R mice showed atypical tail suspension phenotype ............... 65

3.3.3 Heterozygous H304R/+ mice show mild defects in their motor

phenotype .................................................................................................... 69

3.3.4 Homozygous H304R/+ mice show severe defects in their motor

phenotype .................................................................................................... 75

3.4 Conclusions ........................................................................................... 81

CHAPTER 4: SENSORY PHENOTYPE CHARACTERIZATIONS OF H304R

MICE .................................................................................................................. 85

4.1 Introduction ............................................................................................ 85

4.1.1 Sensory deficits in previously published mutant dynein mice models

85

4.2 Rationale ............................................................................................... 88

4.3 Results .................................................................................................. 89

4.3.1 Altered latency to respond to pain stimulus in H304R mice ............ 89

4.3.2 Sciatic nerve appeared thinner in H304R mice ............................... 92

xiii

4.3.3 Reduced total sciatic nerve fascicular area in H304R mice ............ 93

4.3.4 Axonal counts in the sciatic nerve cross section ............................. 98

4.3.5 Identifying subpopulations of neurons in the adult DRG ............... 102

4.4 Conclusions ......................................................................................... 107

CHAPTER 5: MOTILITY STUDIES OF H304R MICE ...................................... 111

5.1 Introduction and rationale .................................................................... 111

5.2 Results ................................................................................................ 112

5.2.1 Synaptophysin-RFP positive vesicular motility in hippocampal

neurons ..................................................................................................... 112

5.2.2 Synaptophysin-RFP positive vesicular motility in cortical neurons 117

5.2.3 Rab7-GFP positive vesicular motility in cortical neurons .............. 122

5.3 Conclusions ......................................................................................... 125

CHAPTER 6: DISCUSSION ............................................................................. 127

6.1 All dynein mutations don’t act the same .............................................. 127

6.2 Relevance of H304R Mouse Model ..................................................... 129

6.3 Comparison of H304R Mouse Model with Other Dynein Mouse Models

133

6.4 Future Sensory Characterization Work of H304R Mouse Model ......... 136

6.5 Potential Dynein Disruptions in The H304R Mice and Future Studies . 139

xiv

APPENDIX A: DISSERTATION DEFENSE ANNOUCEMENT ......................... 142

REFERENCES ................................................................................................. 148

xv

LIST OF FIGURES

Figure 1-1 Intracellular motility inside a cell .......................................................... 2

Figure 1-2 Cytoplasmic Dynein Structure ............................................................. 3

Figure 1-3 Cytoplasmic Dynein Heavy Chain (DHC) peptide ............................... 4

Figure 1-4 Binding partners of dynein ................................................................. 13

Figure 1-5 Mutations in Dynein Heavy Chain (DHC) .......................................... 16

Figure 1-6 Clinical symptoms associated with Charcot Marie Tooth (CMT) ....... 21

Figure 1-7 CMT2O and SMA-LED linked mutation in dynein heavy chain ......... 23

Figure 1-8 Dynein heavy chain (DHC) mutations in mice ................................... 26

Figure 1-9 Pseudo-unipolar sensory neurons in DRGs ...................................... 30

Figure 1-10 Neurotrophic factors in sensory neurons development ................... 31

Figure 1-11 Neurotrophic factors activate Trk signaling in neurons .................... 32

Figure 1-12 Sciatic nerve anatomy in mice ......................................................... 36

Figure 1-13 Sciatic nerve is a mixed nerve ......................................................... 37

Figure 2-1 Generation of knock-in H304R mouse model .................................... 41

Figure 2-2 Tail suspension test ........................................................................... 45

Figure 2-3 Grip strength test meter ..................................................................... 46

Figure 2-4 Rotarod machine to measure motor coordination ............................. 47

Figure 2-5 Rotarod test profile ............................................................................ 48

Figure 2-6 Tail flick meter ................................................................................... 49

Figure 2-7 Schematic diagram of motility analysis using kymographs ................ 57

Figure 3-1 Comparison of dynein heavy chain peptide ....................................... 62

xvi

Figure 3-2 Gait phenotype of H304R mouse model ........................................... 65

Figure 3-3 Tail suspension phenotype of H304R mouse model ......................... 66

Figure 3-4 Atypical tail suspension response of H304R mouse model ............... 67

Figure 3-5 Grip strength phenotype of heterozygous H304R/+ mice .................. 70

Figure 3-6 Rotarod performance test of the H304R/+ mice ................................ 73

Figure 3-7 Grip strength phenotype of heterozygous H304R/R mice ................. 76

Figure 3-8 Rotarod performance test of the H304R/R mice ............................... 79

Figure 4-1 Sciatic nerve cross section ................................................................ 86

Figure 4-2 Tail flick pain sensitivity test of H304R male mice ............................. 89

Figure 4-3 Comparison of sciatic nerve thickness of H304R male mice ............. 93

Figure 4-4 Sciatic nerve cross sections of 3-month H304R male mice ............... 94

Figure 4-5 Sciatic nerve cross sections of 12-month H304R male mice ............. 95

Figure 4-6 Total fascicular area of sciatic nerve in H304R male mice ................ 96

Figure 4-7 The largest fascicle inside the sciatic nerve of each genotype .......... 99

Figure 4-8 Total count of axons inside a fascicle .............................................. 100

Figure 4-9 The percentage distribution of bundled axons vs single axons ....... 101

Figure 4-10 Serial sections of dorsal root ganglia of H304R mice .................... 103

Figure 4-11 Neuronal subpopulations staining of dorsal root ganglia ............... 106

Figure 5-1 Synaptophysin-RFP positive vesicles motility in hippocampal neurons

at DIV 10 .......................................................................................................... 114

Figure 5-2 Synaptophysin-RFP positive vesicles motility in hippocampal neurons

at DIV 14 .......................................................................................................... 116

xvii

Figure 5-3 Synaptophysin-RFP positive vesicles motility in cortical neurons at

DIV 10 .............................................................................................................. 119

Figure 5-4 Synaptophysin-RFP positive vesicles motility in cortical neurons at

DIV 14 .............................................................................................................. 121

Figure 5-5 Rab7-GFP positive vesicles motility in cortical neurons at DIV 10 .. 124

Figure 6-1 Severity of phenotypes associated with DHC mutations ................. 135

xviii

LIST OF TABLES

Table 1-1 Cellular functions of cytoplasmic dynein ............................................... 8

Table 1-2 Dynein heavy chain mutations in humans .......................................... 17

Table 1-2 Types of Charcot-Marie-Tooth disease in humans ............................. 22

Table 1-3 Dynein heavy chain mutations in mice ............................................... 27

Table 1-4 Major types of somatosensory neurons in the DRG ........................... 38

Table 2-1 Antibodies used for tissue immunohistochemistry .............................. 52

Table 2-2 Microscopic settings used for imaging ................................................ 53

Table 3-1 Tail suspension reflex of H304R mice ................................................ 68

Table 3-2 Grip strength in H304R/+ male mice .................................................. 71

Table 3-3 Grip strength in H304R/+ female mice ............................................... 72

Table 3-4 Rotarod performance in H304R/+ male mice ..................................... 74

Table 3-5 Rotarod performance in H304R/+ female mice .................................. 74

Table 3-6 Grip strength in H304R/R male mice .................................................. 77

Table 3-7 Grip strength in H304R/R female mice ............................................... 78

Table 3-8 Rotarod performance in H304R/R male mice ..................................... 80

Table 3-9 Rotarod performance in H304R/R female mice .................................. 80

Table 4-1 Tail flick data ...................................................................................... 91

Table 4-2 Total fascicular area in the sciatic nerve of H304R male mice ........... 97

Table 4-3 Axon counts inside the largest fascicle ............................................. 100

Table 4-4 Optimized conditions for identifying DRG neuronal subpopulations . 104

Table 4-5 Preliminary counts of wildtype DRG subpopulation .......................... 105

xix

Table 5-1 Synaptophysin-RFP positive vesicles motility in hippocampal neurons

at DIV 10 .......................................................................................................... 113

Table 5-2 Synaptophysin-RFP positive vesicles motility in hippocampal neurons

at DIV 14 .......................................................................................................... 115

Table 5-3 Synaptophysin-RFP positive vesicles motility in cortical neurons at DIV

10 ..................................................................................................................... 118

Table 5-4 Synaptophysin-RFP positive vesicles motility in cortical neurons at DIV

14 ..................................................................................................................... 120

Table 5-5 Rab7-GFP positive vesicles motility in cortical neurons at DIV 10 ... 123

xx

LIST OF ABBREVIATIONS

AAA+ - ATPase Associated Family of proteins

ATP – Adenosine Tri Phosphate

ALS – Amyotrophic Lateral Sclerosis

CMT – Charcot Marie Tooth

CMT2O – Charcot Marie Tooth Type 2O

Cra – Cramping

DHC – Dynein Heavy Chain

DIV – days in vitro

DRG – Dorsal Root Ganglia

GLUT1- Glucose transporter 1

H304R – Histidine 304 Arginine

H304R/+ - Heterozygous mutant of Histidine 304 Arginine

H304R/R – Homozygous mutant of Histidine 304 Arginine

H306R – Histidine 306 Arginine

ID – Intellectual Disability

Loa – Legs at odd angle

xxi

MCD – Malfunctions of Cortical Development

PBS – Phosphate Buffered Saline

SMA-LED – Spinal Muscular Atrophy with Lower Extremity Dominant

Swl – Sprawling

1

CHAPTER 1: INTRODUCTION

Dynein is a motor protein present ubiquitously in many living organisms ranging

from small microorganisms (yeast, fungi, parasite etc.) to humans. One of the

many functions of dynein is to carry intracellular cargo on microtubule tracks

(Paschal and Vallee, 1987). Microtubules are polarized cytoskeletal filaments

that are usually arranged with their growing plus ends at the cell periphery and

their minus ends at the microtubule-organizing center (MTOC) at the cell center.

Dynein utilizes adenosine triphosphate (ATP) as an energy source to power its

motor function and carries cellular cargoes from the cell periphery to the minus

end of the microtubule at the cell center (retrograde). In contrast, the motor

protein kinesin carries the cellular cargoes from the cell center to the plus end of

the microtubule that is usually located at the cell periphery (anterograde).

In eukaryotic cells, there are 3 classes of dynein – cytoplasmic dynein 1,

cytoplasmic dynein 2, and axonemal dynein (Wickstead and Gull, 2007) (Yagi,

2009). Cytoplasmic dynein 1 is involved in minus end directed transport within

the cell body and has extensive cellular functions ranging from intracellular

transport to cell division activities (listed in the Table 1-1). Cytoplasmic dynein 2

acts as a motor protein inside cilia/flagella and is involved in intraflagellar

transport of structural components for flagellar assembly (Pazour et al., 1999).

2

Figure 1-1 Intracellular motility inside a cell

Motor proteins dynein and kinesin carry cargoes like mitochondria and vesicles on the microtubule tracks. Dynein is minus end directed motor protein, while kinesin is plus end directed motor protein

Cytoplasmic dynein 2 transports cargoes from the tip to the base of cilia where

the minus end of microtubule is located (Porter et al., 1999). The axonemal

dynein is integrated with the microtubule to make the core structure of cilia called

axoneme (Gibbons and Rowe, 1965). Axonemal dynein is involved in powering

the ciliary beating to make the cilia and flagella motile. Cytoplasmic dynein 1 will

be referred just as ‘cytoplasmic dynein’ or ‘dynein’ from now on in this document.

3

1.1 Cytoplasmic Dynein Structure

Dynein is a large multimeric protein complex that is ~1.2 megaDalton in size. A

functional dynein motor unit (Figure 1.1) is comprised of many subunits – 1 pair

of dynein heavy chains (DHC), 1 pair of dynein intermediate chains (DIC), 2 pairs

of dynein light intermediate chains (DLIC), and 3 pairs of dynein light chains

(DLC). The dynein heavy chain (DHC) monomers dimerize at the N-terminal to

form a heavy chain dimer that interacts with other dynein subunits (DIC, DLIC,

DLC) to form a functional dynein motor complex.

Figure 1-2 Cytoplasmic Dynein Structure

Dynein is a multimeric protein comprising of subunits - dynein heavy chain (DHC), dynein light intermediate chain (DLIC), dynein intermediate chain (DIC) and dynein light chain (DLC). Dynein heavy chain has six ATPase heads and a microtubule (MT) binding domain.

Dynein heavy

chain (DHC)

DLIC

DIC

DLC

4

1.1.1 Cytoplasmic Dynein Heavy Chain Structure

The cytoplasmic dynein heavy chain (DHC) is encoded by the gene dync1h1 and

the peptide is ~532 kiloDalton in size. Cytoplasmic dynein heavy chain (DHC)

gene - dync1h1 is an essential gene. Animals with homozygous dynein null

mutations displayed embryonic lethality at embryonic day 8.5 where the embryo

development couldn’t proceed past the gastrulation stage due to disruptions in

Golgi trafficking and function (Harada et al., 1998). A single DHC monomer is

4646 amino acids long peptide in humans and 4644 amino acids long in mice.

The DHC can be divided into two main domains – N terminal tail domain and C

terminal globular head domain (Figure 1.2). Two monomers of DHC

homodimerize at the N-terminal tail region and interact with other subunits to

form a single functional unit of dynein. The C terminal globular head has 6

repeating AAA+ ATPase units forming a ring structure arrangement (Burgess et

al., 2004) (Sakato and King, 2004).

Figure 1-3 Cytoplasmic Dynein Heavy Chain (DHC) peptide

Dynein heavy chain has two domains – N terminal tail domain and C terminal head domain. There are also six AAA ATPase domains (AAA1-AAA6) that make up the motor dynein ring.

2

5

The six ATPase rings domains (AAA1-AAA6) are responsible for the motor

activity of DHC (Kon et al., 2012). Ring domain AAA1 has the p-loop motif that

can bind and hydrolyze ATP to generate energy for dynein motor activity

(Gibbons et al., 1991). Domains AAA2-AAA4 has p-loop motifs and ability to bind

ATP, whereas domains AAA5-AAA6 lack ATP binding sites. Overall, domains

AAA2-AAA6 lack ATP hydrolysis activity but do provide structural integrity and

regulate functions of the dynein motor ring (Mocz et al., 1991) (Neuwald et al.,

1999). A coiled-coil ‘stalk’ region is located between AAA4-AAA5 in the motor

ring and contains the microtubule binding domain (MTBD) of dynein heavy chain

at its tip (Koonce, 1997). ‘Linker’ domain lies across the six ATPase rings and it

changes its conformation based on the ATP binding state of dynein (Roberts et

al., 2012).

The cytoplasmic dynein undergoes a mechanochemical cycle where mechanical

conformational changes and chemical transitions regulate the motor activity. A

typical mechanochemical cycle of dynein consists of the following steps

(Burgess et al., 2003; Imamula et al., 2007; Kon et al., 2005) -

1. In the initial state, dynein is attached to the microtubule track via the

microtubule binding domain (MTBD) in the stalk region.

2. When an ATP molecule binds to the AAA+ ATPase domain of dynein

heavy chain, the MTBD of the stalk detaches from the microtubule.

3. Detachment of stalk from the microtubule causes a conformational change

in the linker domain and linker gets rotated.

6

4. Upon ATP hydrolysis, MTBD of stalk binds to a new position on the

microtubule and that leads to the release of ADP and Pi (inorganic

phosphate molecule).

5. Released Pi causes linker domain to straighten and generate a

“powerstroke” that extends dynein tail along with its cargo to move further

along towards the minus end of the microtubule track.

1.2 Cytoplasmic Dynein Function

Dynein has variety of roles and functions to play in a healthy functioning cell.

Dynein is involved in both cell division and interphase activities of the cell. Dynein

moves its cargoes on the microtubule in a “stop and go” manner. If the same

cargo has dynein and kinesin attached to it, then the transport of the cargo can

momentarily stop and the travel direction can get switched sometimes from the

retrograde to anterograde direction inside the cell (Steffen et al., 1997).

Sometimes the role of dynein is also distinct in different organisms – for example,

dynein transports vesicles in filamentous fungi (Sivagurunathan et al., 2012)

whereas dynein is responsible only for mitotic spindle positioning in yeast (Moore

et al., 2009). Even viruses utilize dynein driven transport of endocytotic vesicles

to reach the nucleus to replicate itself using the host machinery (Dodding and

Way, 2011). These functions of dynein are broadly categorized and summarized

in Table 1-1.

7

It is clear that cytoplasmic dynein is involved in many cellular functions that are

essential for the cell growth, maintenance, and survival. Total loss of dynein

function causes early embryonic lethality in Drosophila melanogaster (Dick et al.,

1996). Heterozygote dynein heavy chain knockout mice did not exhibit any

phenotype, however, the dynein-null homozygous mice were embryonic lethal

and did not survive past the gastrulation stage (Harada et al., 1998).

8

Table 1-1 Cellular functions of cytoplasmic dynein

Category Cargo Cellular functions References

Org

an

elle

tra

nsp

ort

Vesicles Transport of vesicles (Sivagurunathan et al., 2012),

(Koonce, 2000)

Early endosomes Receptor sorting and Rab5 positive early

endosome morphogenesis in HeLa cells

(Driskell et al., 2007)

Peroxisomes Coordinated movement of peroxisomes in

cultured Drosophila melanogaster cells

(Kural et al., 2005)

Melanosomes Regulation of melanosomes transport in

Xenopus melanophore cell line

(Gross et al., 2002)

Golgi vesicles Transport of vesicles from endoplasmic

reticulum to Golgi in mammalian cell lines

(Presley et al., 1997),

(Hoogenraad et al., 2001)

Mitochondria Transport of mitochondria in motor axons of

Drosophila melanogaster and in neurons

(Pilling et al., 2006),

(Schwarz, 2013)

Lysosomes and

late endosomes

Rab7-interacting lysosomal protein (RILP)

recruit dynein to Rab7 positive lysosomes and

late endosomes in mammalian cells

(Jordens et al., 2001),

(Cantalupo et al., 2001),

9


Recyclin

g a

nd

de

gra

datio

n

Phagosomes Dynein and dynactin regulate the maturation

and minus end directed motility of phagosomes

in mammalian cells

(Blocker et al., 1997)

Autophagosomes Maturation of autophagosomes (containing

organelles and soluble proteins for targeted

degradation) is regulated spatially by dynein in

primary neurons

(Maday et al., 2012)

Aggresomes Dynein drives the misfolded protein into

aggresomes for sequestration and eventual

degradation

(Johnston et al., 2002)

Cell

div

isio

n

Nuclear

positioning

Dynein positions nucleus in budding yeast

Saccharomyces cerevisiae during cell division

(Moore et al., 2009)

Spindle

positioning

Cortical dynein pulls and displaces microtubules

for mitosis and meiosis

(McNally, 2013)

Centrosomes Dynein transports and assembles gamma

tubulin and pericentrin at the centrosomes for

microtubule nucleation & organization

(Young et al., 2000)

Chromosome

separation

Dynein associated with the nuclear envelope

drives separation of chromosomes in prophase

(Raaijmakers et al., 2012)

10

Cyto

ske

leto

n &

pro

tein

tra

nspo

rt

Nuclear rotation Dynein rotates and directs the nucleus in the

leading edge of motile fibroblasts

(Levy and Holzbaur, 2008)

Microtubules Dynein maintains stable microtubule and

kinetochore orientation for aligning

chromosomes during metaphase

(Varma et al., 2008)

Neurofilaments Direct interaction and translocation of

neurofilaments by dynein in neuronal cells

(Shah et al., 2000)

Transcription

factors

Retrograde transport of transcription factor such

as glucocorticoid receptor from the cytoplasm to

the nucleus

(Harrell et al., 2004)

Ribonucleoprotein Correct mRNA localization by dynein mediated

apical transport in Drosophila melanogaster

(Wilkie and Davis, 2001)

Vira

l tr

an

spo

rt

HIV‐1 and HSV1 Dynein is hijacked by the viruses such as

Human immunodeficiency virus type 1 (HIV‐1)

and Herpes simplex virus (HSV1) to reach the

nucleus of the host cells for replication

(Dodding and Way, 2011)

Adenovirus Adenovirus capsid hexon directly interacts with

dynein for viral transport to the nucleus

(Bremner et al., 2009)


11

1.2.1 Dynein interactions with its binding partners

There are three families of cytoskeletal motor proteins inside a cell – microtubule-

based dynein, microtubule-based kinesin, and actin-based myosin. Multiple

subfamilies and isoforms of kinesins and myosin are available inside the cells to

perform distinct cellular functions (Vale, 2003). However, dynein has only three

classes inside the cell as mentioned previously – cytoplasmic dynein 1,

cytoplasmic dynein 2, and axonemal dynein. Cytoplasmic dynein 2 and axonemal

dynein are dedicated to ciliary and flagellar functions specifically. Therefore,

cytoplasmic dynein 1 (referred as dynein from now on) is solely responsible for

multiple cellular functions in all cell types besides cilia/flagella. It is no wonder

that dynein utilizes and interacts with various adaptor proteins to carry out

diverse cellular functions (Vallee et al., 2012).

Binding of dynein with its interacting partners controls different cellular functions

and is needed to load the cargo onto the motor complex. Dynein cargoes range

from membranous vesicles, organelles, proteins, cytoskeletal components and

even viruses. Furthermore, an active motor complex facilitates varied dynein-

based functions such are organelle transport and positioning (Burkhardt et al.,

1997), mitotic chromosome separation (Dujardin et al., 1998), and nuclear

migration (Levy and Holzbaur, 2008). Dynein interacts with many intracellular

binding partners via direct or indirect interactions (Figure 1-4). DHC participates

in the catalytic motor activity and other dynein subunits like LCs, ICs, LICs

12

interact with adaptor proteins to regulate various dynein functions in the cell

(Pfister, 2015).

The direct binding partners of dynein include nuclear distribution protein E

(NudE) (McKenney et al., 2011), lissencephaly (LIS 1) (Huang et al., 2012),

dynactin (dA) (Schroer, 2004), RAB-interacting lysosomal protein (RILP)

(Schroeder et al., 2014), Huntingtin protein (Htt) (Caviston and Holzbaur, 2009),

Snapin on late endosomes (Cai et al., 2010), adenoviral hexon protein (Bremner

et al., 2009), and pericentrin centrosomal protein (Tynan et al., 2000).

Dynactin regulates dynein driven motility by enhancing dynein’s binding with the

microtubule and making the transport more processive (Culver-Hanlon et al.,

2006). Additionally, dynactin also acts as an cargo adaptor for dynein and helps

dynein link with multiple cargo adaptors such as - Ankyrin 2 (AnkB) on

membranous cargoes (Lorenzo et al., 2014), RILP on Rab7 positive late

endosomes (Jordens et al., 2001), c-Jun N-terminal kinase-interacting proteins

(JIPs) on neuronal cargoes (like amyloid precursor protein) (Fu and Holzbaur,

2013), Huntington-associated protein 1 (HAP1) on neuronal vesicles (Engelender

et al., 1997), Spectrin on Golgi vesicles & late endosomes/lysosomes (Holleran

et al., 1996), bicaudal D/BICD2 on Golgi-ER vesicles (Liu et al., 2013; Matanis et

al., 2002), and TRAK/Milton adaptor proteins on mitochondria (van Spronsen et

al., 2013).

13

Figure 1-4 Binding partners of dynein

Dynein interacts with various cargo adaptors either directly (shown in blue arrows) or indirectly (shown in orange arrows) to achieve versatility in its cellular functions. Dynein heavy chain (DHC), dynein light intermediate chain (DLIC), dynein intermediate chain (DIC) and dynein light chain (DLC).

DHC

DLIC

DIC

DLC

Dynactin BicD, RILP, JIP, HAP1, AnkB, Spectrin, TRAK/Milton

Adenoviral hexon protein, pericentrin, RILP,

NudE/NudEL Lis1

Lis1

DIC Htt, Snapin

Direct interaction with dynein

Indirect interaction with dynein

14

1.2.2 Dynein mutations linked to neurodegeneration in humans

Neurons are highly polarized cells with axonal projections as long as up to a

meter in length. Consequently, it is understood that neurons need robust axonal

transport for survival. Dynein has an established role in retrograde transport of

cargoes from the neuronal tip to soma in neurons (Perlson et al., 2010).

Additionally, dynein interacts with dynactin to form a highly processive motor

complex (Culver-Hanlon et al., 2006) that has an important role in axonal

transport (Waterman-Storer et al., 1997). The dynein/dynactin motor protein

complex transports cargo from the tip of the axon towards the neuronal cell body

(retrograde direction). Other functions of dynein in neurons includes growth and

development of neurites (Barakat-Walter and Riederer, 1996), formation of

synapse in neurons (Cheng et al., 2006), axonal transport of microtubules &

neurofilaments (He et al., 2005) and intracellular organelles (Schnapp and

Reese, 1989), and neuronal migration inside the cortical layer of the brain

(Sasaki et al., 2000).

Seeing how crucial dynein is for neuronal growth and maintenance, it would not

seem very surprising to see a dynein heavy chain mutation leading to

neuropathies in humans (figure 1-5). The first dynein heavy chain mutation

H306R was reported in 2011, that lead to a hereditary motor-sensory

neurodegenerative disease called Charcot Marie Tooth Disease Type 2O in

humans (Weedon et.al. 2011). Since then at least 17 genetic mutations spanning

the motor and tail domains of the dynein heavy chain have been identified in

15

neurodegenerative diseases in humans (summarized in Table 1-2). The reported

dynein heavy chain mutation related diseases in humans can be broadly

classified as motor-sensory neuropathies, central nervous system deficits, and

congenital defects.

16

Figure 1-5 Mutations in Dynein Heavy Chain (DHC)

Mouse mutations (top, in blue text) and human mutations (bottom, in red text) in the DHC peptide. Mice mutations are reported in the tail domain of the DHC peptide, whereas the human mutations span the entire length of the DHC peptide.

17

Table 1-2 Dynein heavy chain mutations in humans

Disease DHC mutation Clinical symptoms Reference

Charcot Marie Tooth

Type 2O (CMT2O)

H306R muscle weakness and wasting, gait

abnormalities, skeletal deformities,

motor milestone delays, loss of

sensation

(Weedon et al.,

2011)

Spinal Muscular Atrophy

– Lower Extremity

Dominance (SMA-LED)

I584L, K671E, Y970C,

G1132E

Severe muscular atrophy in legs,

loss of motor neurons, delay in

motor milestones, waddling gait

(Harms et al.,

2012)

Malformation of Cortical

Development

deletion 659-662, K129I,

K3336N, R3384Q, R1567Q,

R3344Q, R1962C, K3241T,

R3344Q

Neuronal migration defects in

cerebellar cortex, microcephaly or

macrocephaly, polymicrogyria,

epilepsy/seizure in severe cases

(Poirier et al.,

2013)

18

Disease DHC mutation Clinical symptoms Reference

Intellectual disability E1518K, H3822P Defects in neuronal migration,

cortical development malfunctions,

impaired speech, reduced

intellectual and mental functions,

gait problems

(Willemsen et al.,

2012)

Congenital cataracts and

gut dysmotility

R2332C Cataract at birth, oral dysphagia,

dysmotility in gut, developmental

milestone delay, bifrontal

polymicrogyria

(Gelineau-Morel

et al., 2016)

19

It is interesting to note that even though these mutations lie in the same DHC

gene, they result in different clinical manifestations in humans. The DHC motor

domain mutations generally lead to cortical development type of defects,

whereas, the DHC tail domain mutations were observed to cause motor-sensory

type of defects in the reported human neuropathies. Moreover, it is important to

note that all dynein mutations in humans exhibit symptoms that are heavily

related to neuronal defects rather than any other type of defects in humans. This

suggests that dynein must have a specialized role in the neurons because even

the smallest single point mutation in dynein can disrupt the regular functions of

the central or peripheral nervous system. Hence, it is evident that mammalian

neurons must be particularly sensitive to the dynein mutations. It would be

valuable to understand how dynein heavy chain mutations cause diseases

specifically of the nervous systems in humans.

20

1.3 Charcot-Marie-Tooth Diseases in Humans

Charcot-Marie-Tooth (CMT) disease is the most common type of hereditary

motor-sensory neuropathy in humans with estimated prevalence rate is 1 in 2500

humans. CMT is characterized by symmetric distal limb weaknesses, muscular

atrophy in hands and/or feet, loss of sensation in lower limbs that may lead to

foot ulcerations, and skeletal defects like pes cavus (high arching of feet) (Figure

1-6).

More than 80 genes that are involved in neuronal structure maintenance signal

transduction, and axonal transport are reported to have mutations that are linked

with CMT (Bird, 1993). CMT patients are categorized into different types (table 1-

3) based on their nerve conduction velocity (NCV) and family history-based

inheritance of the disease. In each category, letters are assigned to represent the

gene involved. For example – CMT1A, CMT1B, CMT1C and so on, CMT2A,

CMT2B, CMT2C and so on. CMT1A indicates CMT Type 1A which is caused by

a mutation in PMP22 gene (myelination gene) (Patel et al., 1992) whereas

CMT1B indicates CMT Type 1B which is caused by a mutation in MPZ gene

(another myelination gene).

CMT is clinically diagnosed by molecular genetic testing and by identifying

peripheral neuropathy symptoms during their physical examinations and in their

previous medical history. CMT has a variable onset in patients, ranging from as

early as age at birth to as late as age 50. Currently there is no cure for CMT, and

the treatment is based on the clinical symptoms presented by the patients. CMT

21

patients often have to rely on surgical treatments, physical therapy, and use of

prosthetics like crutches/ankle braces to cope with the symptoms. About 5% of

all CMT patient are dependent on wheelchair for mobility.

Another challenge with CMT is the heterogeneity of the disease onset,

progression, and clinical symptoms even in related patients from the same family

pedigree. For example- in the Dominant Intermediate Charcot-Marie-Tooth

disease (DI-CMT), some patients from the same family may show demyelination

associated symptoms, whereas other patients from the same family may show

axonal degeneration, and some may even show both demyelination and axonal

demyelination form of the CMT symptoms.

Figure 1-6 Clinical symptoms associated with Charcot Marie Tooth (CMT)

CMT patients primarily show distal lower limb muscle wasting and atrophy, foot deformity (pes cavus), foot ulcerations, and gait abnormalities.

22

Table 1-3 Types of Charcot-Marie-Tooth disease in humans

Type of CMT Type of defect Inheritance Some examples of mutations (Bird, 1993)

CMT Type 1

(most common)

Demyelination with

reduced NCV of <35m/s

Autosomal

dominant

PMP22, MPZ, LITAF, ERG2, NEFL

CMT Type 2 Axonal degeneration with

normal NCV>45m/s

Autosomal

dominant

MFN2, RAB7, TRPV4, GARS, NEFL, MPZ,

DYNC1H1, GDAP1, HSPB8, LRSAM1, MORC2,

DI- CMT

(Dominant

Intermediate)

Demyelination and axonal

degeneration with NCV of

35-45m/s

Autosomal

dominant

GDAP1, MPZ, DNM2, DRP2, GNB4, INF2,

PLEKHG5, YARS,

CMT Type 4

(rare)

Demyelination and axonal


35-45m/s

Autosomal

recessive

GDAP1, MTMR2, SBF2, SH3TC2, NDRG1,

EGR2, PRX, FGD4, FIG4, HINT1

CMT Type X Demyelination and axonal


35-45m/s

X-linked DNM2, MPZ, YARS, GJB1

23

1.3.1 Dynein mutation linked to CMT2O and SMA-LED

A dynein heavy chain mutation linked to Charcot-Marie-Tooth type 2O (CMT2O)

in humans was first identified in 2011 (Weedon et al., 2011). Exome sequencing

of genes in a four generation European family identified a single point missense

substitution of Histidine to Arginine at position 306 (H306R) in cytoplasmic dynein

heavy chain 1 peptide (figure 1-7). The autosomal dominant H306R mutation

affected 23 members (male and female) in this family and they showed a varied

range of clinical symptoms.

Figure 1-7 CMT2O and SMA-LED linked mutation in dynein heavy chain

Missense substitution mutation (Histidine to Arginine) at position 306 in human cytoplasmic dynein heavy chain peptide has been linked to two distinct neuropathies – Charcot Marie Tooth type 2O (CMT2O) and spinal muscular atrophy with lower extremity dominant (SMA-LED).

CMT2O patients with autosomal dominant heterozygous H306R mutation have

varied disease onset ranging from early childhood to early adulthood. CMT2O

patients reportedly had delayed motor developmental milestones, problems in

walking or running, recurrent falls, slow progressing muscle weakness and

muscle wasting in lower legs, and skeletal deformities pes cavus (high arching of

24

foot). Physiological tests with CMT2O patients revealed normal nerve conduction

velocities, evidence for axonal degeneration in sural nerve biopsies, and

denervation defects in muscle biopsies. Several patients that were severely

affected with CMT2O also reported to have neuropathic pain and prickling

sensation (paresthesia) in their lower limbs. A few patients also reported loss of

reflexes and sensation, speech delays & behavioral problems, lumbar lordosis,

spinal and hip issues leading to waddling gait, and loss of proximal muscles in

the upper back (periscapular weakness).

In parallel, an independent study of a Japanese family with the same H306R

mutation presented clinical symptoms that were consistent with spinal muscular

atrophy with lower extremity dominant (SMA-LED) (Tsurusaki et al., 2012). The

patients in this family showed clinical features that were different from CMT2O

patients, such as proximal lower limb weakness instead of distal limb weakness

and non-progressive nature of muscular atrophy/wasting. SMA-LED patients with

H306R mutation showed motor neuron disease involvement but not sensory

neuron involvement. It is interesting to note that a family member in the

European family with CMT2O showed proximal limb atrophy like SMA-LED

patients in this Japanese family. In conclusion, the single point mis-sense

substitution of H306R in cytoplasmic dynein heavy chain has been reported to

cause to two separate neuropathies in humans – CMT2O and SMA-LED.

25

1.4 Mouse Models with Neurodegeneration Linked Dynein Mutations

There are three reported mouse models with DHC mutations – Legs at odd

angles (Loa), Cramping (Cra), Sprawling (Swl) (figure 1-8). These mice were

named after their abnormal hind limb clenching phenotype and their posture.

Heterozygotes of all three mouse models had normal lifespan, whereas the

homozygotes showed severe phenotype & did not survive embryonically or after

birth (Chen et al. (2007); (Hafezparast et al., 2003).

Loa and Cra mice were generated independently via N-ethyl-N-nitrosourea

(ENU) treated mice and were identified due to abnormal clenching of hind limbs

when lifted by tail (Hrabe de Angelis et al., 2000). Swl was created via radiation-

induced mutagenesis, also displayed abnormal clenching of hind limbs (Duchen,

1974). All three Loa, Cra, and Swl are autosomal dominant mutations that lead to

neurodegenerative phenotypes. Loa/+ and Cra/+ developed late-onset of motor

neuropathy, whereas Swl/+ showed early-onset of hereditary sensory

neuropathy. The key features of the three DHC mutant heterozygous mice are

listed below in table 1-4.

26

Figure 1-8 Dynein heavy chain (DHC) mutations in mice

Mouse mutations in the DHC peptide are reported in the tail domain and are associated with Loa, Cra, and Swl mice.

27

Table 1-4 Dynein heavy chain mutations in mice

DHC mutation Phenotype

Legs at odd angles (Loa) –

F580Y

(Hafezparast et al., 2003)

Loa/Loa homozygous lethal

late-onset motor neuron

degeneration in Loa/+

gait abnormalities

reduced front and hind limb strength,

thinner dorsal root and sciatic nerve

abnormal proprioception & absence of

H-reflex

lumbar DRG neurons loss

mild loss of spinal α- motor neurons

muscle spindle loss

neuronal migration defects

Cramping (Cra) – Y1055C

(Hafezparast et al., 2003)

Cra/Cra homozygous lethal

late-onset sensory neuron

degeneration in Cra/+

gait abnormalities

reduced hind limb strength

thinner dorsal root

abnormal muscle pathology,

28

mild loss of spinal α- motor neurons

behavioral hyperactivity

mitochondrial dysfunction in skeletal

muscles

Sprawling (Swl) –

[GIVT]1040[A]

(Chen et al., 2007)

Swl/Swl homozygous lethal

early-onset hereditary sensory

neuropathy in Swl/+

gait abnormalities

reduced hind limb strength,

thinner dorsal root and sciatic nerve

abnormal proprioception & absence of

H-reflex

lumbar DRG neurons loss

muscle spindle loss

To better understand the role of dynein in neurodegeneration, Loa/+, Cra/+, and

Swl/+ heterozygote mice were separately cross bred with the mutant SOD1G93A

mouse model of amyotrophic lateral sclerosis (ALS). ALS is a fast progressing

neurological disease marked by progressive loss of motor neurons, leading to a

total paralysis and lethality 3-5 years after onset. The gain of function mutation

G93A in superoxide dismutase 1 (SOD1) is autosomal dominant and kills motor

29

neurons rapidly (Valentine et al., 2005). It was interesting to note that when Loa/+

and Cra/+ were crossed separately with ALS SOD1G93A mice, their double

heterozygotes pups showed attenuation of the ALS SOD1G93A symptoms and

improved lifespan (Kieran et al., 2005) (Teuchert et al., 2006). This was a very

unsuspected finding and opposite of what was expected at the time. Loa/+ and

ALS SOD1G93A double heterozygotes also showed a remarkable increase in its

retrograde axonal transport. Swl/+ and SOD1G93A heterozygotes did not show any

attenuation or improvement of ALS SOD1G93A associated phenotype. It is

interesting to note that even though not all three mutations lie in the same tail

region of DHC, their attenuation response to ALS mutant SOD1G93A was the

same.

Undoubtedly, the three dynein heavy chain mutant mice (Loa, Cra, Swl) provided

many key insights into understanding the dynein function in neurodegenerative

disorders. However, Loa, Cra, and Swl mutations are not linked to any disease

alleles in humans and it is difficult to correlate their findings to human

neuropathies. Nevertheless, it would be very interesting to compare Loa, Cra,

and Swl mice studies with other DHC mice models that have dynein mutations

linked to an actual human disease.

30

1.5 Peripheral Nervous System of Mouse

The peripheral nervous system of mouse comprises of nerves and ganglia

arising out of the spinal cord. The dorsal root ganglia (DRG) is a cluster of

sensory neuron cell bodies located in pairs in the spinal column of mice that start

forming in the embryo at embryonic day 13 (E13) (Lawson and Biscoe, 1979).

The sensory neurons located in the DRGs are pseudo-unipolar in nature and do

not have dendrites. The DRG sensory neurons have a single axonal projection

that branches into two – peripheral and central branches (figure 1-9). The

peripheral branch brings in the nerve impulses from the innervated tissues (skin

or muscle) to the neuronal cell body located in the DRG. The central branch

carries the impulse away from the soma to the central nervous system i.e. the

spinal cord or brain. Both branches are myelinated and carry the nerve impulses

towards the central nervous system (shown by the arrows in figure 1-9).

Figure 1-9 Pseudo-unipolar sensory neurons in DRGs

Sensory neurons in the Dorsal Root Ganglia (DRG) are pseudo-unipolar in nature, i.e. it has one cell body and two functional axonal branches – peripheral and central branches.

31

1.5.1 Role of dynein mediated retrograde signaling in sensory neurons

Growth and maturation of sensory neurons inside the DRGs are dependent on

various neurotrophic factors such as nerve growth factor (NGF), glial derived

neurotrophic factors (GDNF), brain derived neurotrophic factors (BDNF), and

neurotrophins (NTs) (Kimpinski et al., 1997) (Markus et al., 2002) (Klusch et al.,

2018). The neurotrophic factors are secreted by the target tissue (muscle, skin,

or visceral organ) and stimulate the development and function of sensory

neurons (figure 1-10).

Figure 1-10 Neurotrophic factors in sensory neurons development

Sensory neurons and motor neurons growth, maturation and survival depends on the neurotrophic factors such as neurotrophins (NTs) and glial derived growth factors (GDNF) released by the target tissue they innervate.

The neurotrophic factors bind to the tyrosine kinases (Trk) receptors located on

the axonal terminals of the peripheral branch of the sensory neuron (table). NGF

32

specifically binds to TrkA receptors, NT4 and BDNF specifically bind to TrkB

receptors, and NT3 specifically binds to the TrkC receptors (figure 1-11).

Figure 1-11 Neurotrophic factors activate Trk signaling in neurons

Neurotrophic factors bind to a specific Trk receptor and activate downstream signaling of neuronal maintenance and survival factors. NGF= Nerve Growth Factor, BDNF= Brain Derived Neurotrophic Factors, NT3/4= Neurotrophins 3/4.

Binding of neurotrophic factors leads to dimerization of the receptors and their

autophosphorylation that recruits the effectors for downstream Trk signaling. The

activated Trk signals send survival factors such as mitogen-activated protein

kinase (MAPK); phosphatidylinositol 3-kinase (P13K); phospholipase (PLC-γ)

33

etc. (Chao, 2003) (Delcroix et al., 2003) to the neuronal cell body located in the

DRG via dynein mediated retrograde transport (Heerssen et al., 2004) (Ito and

Enomoto, 2016).

Neurotrophin mediated retrograde signaling also facilitates differentiation and

specification of neuronal subpopulations in the peripheral nervous system of

mice. Studies with BAX-/- and TrkA-/- double mutant mice lacking the NGF

mediated TrkA signaling showed an absence of calcitonin gene related peptide

(CGRP) expressing neuronal subpopulation in the mice DRGs (Suzuki et al.,

2010). Other studies have explored and established the role of target derived

neurotrophic factors (such as neurotrophins & BDNF) in determining the

cholinergic/peptidergic phenotype of the sensory neurons during their

development and differentiation in mice (Bergmann et al., 1997; Cellerino et al.,

1996).

Several in vitro studies have investigated and established the role and

importance of retrograde transport of neurotrophic factors. Neurotrophic factors

released by the target tissues bind to the specific receptors in the terminal ends

of the sensory axons innervating the tissue. Then the neurotrophic factors

mediated signaling cascade is activated at the axonal tip and it travels all the way

to the neuronal cell body in retrograde fashion.

Classical experiments with special compartmentalized cell chambers called

Campenot chambers were performed to investigate the spatial effect of

34

neurotrophin Nerve Growth Factor (NGF) on axonal growth (Campenot, 1977).

First, NGF was only supplied selectively to the compartment in the Campenot

chamber that had distal ends of axons growing in it. It was observed that axonal

growth was maintained even though the neuronal cell bodies did not receive

NGF. However, when the NGF was applied only to the Campenot compartment

containing neuronal cell bodies, the distal axons started retracting and

degenerating. These key experiments established that NGF mediated

neurotrophin signaling at the distal ends of neurons were required and adequate

for axonal growth and maintenance.

Subsequent studies then established that neurotrophins bound to the receptors

are internalized and packaged inside signaling endosomes for retrograde

transport to the neuronal cell bodies (Wang et al., 2016). Further investigation of

retrogradely transported signaling endosomes in rat sciatic nerves revealed that

these vesicles contained not only neurotrophin NGF and its receptor TrkA, but

also characteristic markers of early endosomes such as early endosome antigen

1 (EEA1) and Rab5 (small G-protein) (Deinhardt et al., 2006). Additionally,

components of mitogen-activated protein kinase (MAPK) and phosphatidylinositol

3-kinase (P13K) pathways were also found inside the signaling endosomes. The

same study also showed the evidence of dynein dependent retrograde transport

of these NGF-TrkA containing endosomes by colocalization, biochemical, and

nocodazole microtubule disruption assays. Several other independent studies

have also established the role of dynein in retrograde transport of neurotrophic

35

factors – colocalization of 14kDa Dynein Light Chain (DLC) with Trk receptors

(Yano et al., 2001), Rab7 positive vesicles in NGF-TrkA signaling (Saxena et al.,

2005), and p75 and TrkB signaling (Deinhardt et al., 2006).

In conclusion, dynein dependent retrograde signaling is necessary for the

development, specification and maintenance of sensory neurons and it is not

surprising that dynein disruption could cause sensory neuropathies. Numerous

studies have shown that disrupted neurotrophic factor mediated Trk signaling

causes severe sensory neuropathies in mice (Smeyne et al., 1994) (Patel et al.,

2000) (Sleigh et al., 2017).

1.5.2 Sciatic Nerve anatomy in mice

The sciatic nerve is the largest nerve innervating the hind limbs of mice. The

sciatic nerves arise from the sciatic notch near the spine, then continues all the

way from lumbosacral region to popliteal fossa (knee cavity) where it splits into

peroneal, sural and tibial nerves to innervate the lower hind limbs in mice (figure

1-12). The majority of sciatic nerves in C57BL/6J strain mice are made up of the

third lumbar (L3) and fourth lumbar (L4) spinal nerves and a lesser contribution is

received from the fifth lumbar (L5) spinal nerve (Rigaud et al., 2008). The

percentage contribution from each spinal nerve in a sciatic nerve is roughly about

L3 (~28%), L4 (~56%) and L5 (~16%) and no spinal nerve contribution is

reported from L6 at all.

36

Figure 1-12 Sciatic nerve anatomy in mice

Spinal nerves arising from lumbar vertebrae L3, L4, and L5 combine together to form a sciatic nerve in mice that splits up into peroneal, tibial, and sural nerves to innervate the lower hind limbs.

Because the sciatic nerve contains both motor and sensory neurons, it is also

regarded as a “mixed nerve” (Figure 1-13). Dorsal root supplies the sensory

neurons (afferent fibers) and ventral root (efferent fibers) supplies the motor

neurons at lumbar L3, L4, and L5 in mice to make up the “mixed” sciatic nerve in

mice. The sensory nerves bring in the afferent sensing signals from the periphery

target tissue to the spinal cord for processing. Sensing signals can be sensing

temperature, pain stimulus, touch, vibrations etc. In contrast, the motor neurons

send out the efferent action signals from the spinal to the target tissues. Action

signals can be voluntary or involuntary action of a muscle group to a stimulus.

Knee jerk test is a classic example of a reflex arc where the reflex hammer hits

37

the knee (stimulus) and the body reacts quickly by moving your leg away (action)

through rapid involuntary muscle contraction.

Figure 1-13 Sciatic nerve is a mixed nerve

Sciatic nerve has both sensory and motor neurons and is called a “mixed nerve”. Both dorsal and ventral roots supply spinal nerves to make up the sciatic nerve.

1.5.3 Dorsal Root Ganglia anatomy in mice

Neural crest migration gives rise to a cluster of sensory neurons called dorsal

root ganglia (DRG). There are 62 DRGs in total and they are located in pairs

along the spinal cord – 8 pairs in cervical, 13 pairs in thoracic, 6 pairs in lumbar

and 4 in sacral region in the C57BL/6J strain of mice (Malin et al., 2007). There

are many types of heterogeneous somatosensory neuronal subpopulations

present in the DRGs- nociceptors, mechanoreceptors, thermoreceptors, and

proprioceptors (table 1-4). Somatosensory neurons transmit sensory signals to

38

the spinal cord and brain from the target tissues that they innervate such as

muscle spindles, skin, and organs (Liu and Ma, 2011).

Table 1-5 Major types of somatosensory neurons in the DRG

Type of the neuron Size of the fiber Stimulus received

Nociceptors Small diameter (c-fibers) Pain stimulus

Mechanoreceptors Medium diameter (Aδ-

fibers)

Pressure, touch and

vibration

Thermoreceptors Small diameter (AΔ-fibers) Temperature (cold

and warm)

Proprioceptors Large diameter (Aβ-fibers) Muscle stretch and

orientation

Different somatosensory neurons express different types of receptor. Nociceptive

neurons have small sized diameter axons and express the TrkC receptors for

Nerve Growth Factor (NGF) (Suzuki et al., 2010). In addition, peptidergic

nociceptive neurons express calcitonin gene-related peptide (CGRP), whereas,

non-peptidergic nociceptive neurons express IsolecithinB4 (IB4) on its plasma

membrane (Bennett et al., 1996). Chemical stimuli such as capsaicin have been

reported to also stimulate responses in nociceptors expressing capsaicin

receptors called transient receptor potential cation channel subfamily V member

1 (TRPV1) channels (Amaya et al., 2004). Proprioceptive neurons have large

39

sized diameter axons and express TrkC receptors for neurotrophin 3 (NT-3)

uptake.

It is highly common for researchers to investigate the neuronal subpopulations in

the DRGs of mice exhibiting any sensory neurodegenerative symptoms. For

example - in sprawling (Swl) mice, it was reported that the heterozygous Swl/+

mice showed specific loss of its proprioceptive neurons and not in the nociceptive

neuron population in the lumbar dorsal root ganglia (Chen et al., 2007). As a

result, Swl/+ mice showed the corresponding phenotype of early-onset of

proprioceptive sensory neurological defects. In conclusion, investigation of sciatic

nerve anatomy and DRG neuronal subpopulation can provide key insights in

understanding molecular mechanisms behind sensory neuropathies.

40

CHAPTER 2: MATERIALS AND METHODS

2.1 Generation of H304R mouse model

The knock-in H304R mouse model was generated using gene targeting in mouse

embryonic cells (figure 2.1). Using 3-step PCR mutagenesis, the CMT2O disease

linked dynein mutation (CAC to CGC at position 304) was introduced in the

targeting vector with Neo cassette. Presence of the mutation was confirmed by

sequencing the PCR amplified targeted region. Then embryonic stem cells were

electroporated to take in the targeting vector. Embryonic stem cell clones with the

correct targeted single point mutation were confirmed using resistance selection

markers, PCR screening, and southern blot analysis. The embryonic stem cells

were then microinjected in C57BL/6 mouse blastocysts and implanted into

psuedopregnant foster host female mice. The offspring chimeric mice were

obtained, and cross bred to yield the heterozygous knock-in founder mice. Four

heterozygous founder mice with the desired single point mutation in dynein

heavy chain were outcrossed with B6129SF2/J mice (Jackson laboratories) to

produce a colony of different lineages. Heterozygous male and female mice from

different lineages were bred to obtain homozygous litters of mice.

41

Figure 2-1 Generation of knock-in H304R mouse model

Knock-in H304R mouse model was generated by introducing Neo cassette flanked targeting vector that was microinjected into embryonic stem cells. Chimeric mice were selected to obtain founder heterozygous knock-in founder mice.

42

2.2 Mouse colony maintenance

2.2.1 Mouse colony housing

All mice related procedures were carried out with the University of Central

Florida’s Institutional Animal Care and Use Committee (IACUC) guidance and

approval. The mice were checked daily by the trained vivarium staff and were

immediately treated in case of any health issues. Mice were housed in clear

plastic cages lined with sanitized bedding and were provided ad libitum access to

clean food and water. Mice cages were also provided with red colored housing

tube and a wooden block for enrichment of their surroundings. The cages were

kept in an environment-controlled room at 22 ± 2 °C and atmospheric humidity of

50 ± 10%. Mice were exposed to artificial light from 7:00am-7:00pm and to

darkness at 7:00pm-7:00am on a 12-hour diurnal light cycle.

2.2.2 Mouse colony breeding

Heterozygous H304R mice were repeatedly outcrossed to the B6129SF2/J mice

(Jackson Laboratories, stock#101045). Mice pups remained with their mother for

nursing until they were weaned on post-natal day 21 (3 weeks of age). Tail snips

were collected from the 3-week old pups upon weaning for PCR based

genotyping. Mice tails were also tattooed with unique identification numbers

using Labstamp Tattoo system (Somark Innovations).

43

2.2.3 Measurement of mouse body weight

Body weight of a mouse was recorded (in grams) every 2 weeks before the

mouse performed a behavior test. Bodyweight at 3 month of age were

statistically compared between the genotypes (wildtype, heterozygous,

homozygous) and gender (male and female) separately using One-way ANOVA

(GraphPad Prism 6.0 software).

2.2.4 Mouse health supervision

Mice colonies were kept under daily supervision of trained vivarium staff in case

any medical condition would arise. Mice were regularly monitored for their

posture, alertness, activeness, movement around their cages and consumption

of food and water. Physiological phenotypes of mice such as blood in urination,

and abscess on skins due to excessive scratching were immediately treated and

closely monitored. Mice teeth were routinely trimmed if they showed weight loss

due to inability to feed with longer teeth.

Other litter behaviors like fights with littermates, barbering each other, and

shredding of food were also noted. Phenotype like development of eye tumors,

recurrent seizures, and worsening health were also recorded. We immediately

euthanized any mice that showed any signs and symptoms of pain, suffering,

and unusual declining health.

44

2.3 Behavioral phenotype characterization test

All three groups of mice (wildtype, heterozygous H304R/+, homozygous

H304R/R) performed behavioral tests every two weeks starting from 1 month of

age to 24 months of age or until death (whichever came first). Later time points

such as 15, 18, 21 and 24 months of the longitudinal study yielded similar

phenotypic trend in mice at 12 months of age. Hence, the test readings until 12

months only were averaged in a 3-month bin to give time points of 3 month, 6

month, 9 month, and 12 month for data analysis.

The behavior assay data was collected by the researchers that were blinded to

the genotype of the H304R mice. Mice were genotyped by other lab members

and the mice genotypes were revealed after the completion of the study for data

grouping and statistical analyses. Both testers alternated their turn in testing the

same test mouse every two weeks. The behavior tests - tail suspension, grip

strength, rotarod, and tail flick tests were performed on all three groups of H304R

mice (wildtype, heterozygous, homozygous) as described below.

2.3.1 Tail suspension test

The tail suspension test was performed to assess the postural reflex phenotype

of mice. A mouse was gently lifted by its tail and its positioning of hind limbs were

categorized as 'typical' or 'atypical' tail suspension reflex. In a 'typical' tail

suspension reflex (figure 2-2), the mouse splayed its hind limbs away from the

45

body after being suspended by its tail. Whereas, in 'atypical' tail suspension

reflex, the mouse clenched its hind limbs tightly near its body when suspended

by its tail. The data was collected and recorded from two views (ventral and

dorsal) of the mice to get a better evaluation of the phenotype.

Figure 2-2 Tail suspension test

Mouse is gently suspended by its tail and the hind limb response is categorized as typical (shown here) or atypical (hind limbs clenching together).

The percentage of mice displaying ‘atypical’ reflexes was calculated for all three

groups of mice (wildtype, heterozygous, homozygous) at each time point - 3, 6,

9, and 12 months. Then the ‘atypical’ reflex data were statistically compared

between the wildtype and heterozygous H304R/+ and between wildtype and

homozygous H304R/R mice at each time point using the Fisher’s exact test (two-

tailed distribution, p<0.05) on the GraphPad Prism 6.0 software.

46

2.3.2 Grip strength test

A standard grip test was used to measure the muscular limb strength in mice

using a grip strength meter from Harvard Apparatus (BIOSEB GS3# 76-1068). All

limbs or front limbs of mouse were placed on the metal grid attached to the

readout equipment. Then the mouse was gently pulled across the metal grid

using its base of the tail. The equipment recorded the maximum force exerted (in

grams) by the all limbs or front limbs of the mouse being tested (figure 2-3).

Figure 2-3 Grip strength test meter

Grip strength in limbs is measured using a standard grip strength meter that records the maximum force exerted by the mouse.

A test mouse performed 4 consecutive trial each of all limbs and front limbs grip

test. The 4 trials were averaged for all limbs and front limbs respectively to yield

one single readout each on a test day. Next, the averaged all limbs grip strength

readings and the averaged front limbs grip strength readings of a mouse were

47

combined per each genotype at 3-month time intervals - 3, 6, 9, and 12 months.

The all limbs and front limbs grip strength data were statistically compared

between the wildtype and heterozygous H304R/+ and between wildtype and

homozygous H304R/R mice at each time point using Student’s t-test (two-tailed

distribution, p<0.05) on the GraphPad Prism 6.0 software.

2.3.3 Rotarod performance test

Rotarod test is a standard test used to measure motor coordination, muscle

endurance, and motor balance in mice. We performed rotarod test on our

wildtype, heterozygous, and homozygous mice cohorts using rotarod equipment

from IITC Life Sciences (model# 755). The optimized test parameters were input

into the equipment console and 5 mice were placed individually on a stationary

rod inside 5 different lanes before the start of the test (figure 2-4).

Figure 2-4 Rotarod machine to measure motor coordination

Motor coordination in mice is measured by the rotarod performance test. Five mice can simultaneously run on the rotating rod to perform the rotarod performance test.

48

The rotarod started rotating at 5 RPM (Revolutions Per Minute) and gradually

ramped up to 40 RPM in 60 seconds and stayed constant for next 120 seconds

(figure 2-5). The entire run duration of rotarod was 180 seconds including 60

seconds of ramp time followed by 120 seconds of constant RPM. The equipment

recorded a mouse's fall on the plate below the rod during the 180 seconds total

rotarod run window.

Figure 2-5 Rotarod test profile

The rotarod machine starts at 5 RPM and then ramps up to 40 RPM in 60 seconds. Then it stays at 40 RPM for rest of the rotarod test duration.

A mouse's fall on the plate triggered the magnetic switches at the bottom of the

plate and accurately detected and recorded the latency of fall (in seconds).

Software Series 8 was provided with the equipment and it was used to record the

latency of fall for the mice being tested. Each mouse performed three

consecutive trials of the rotarod test, and all three latency of fall readings were

averaged to yield a single test readout for a mouse at a time point. Next, the

49

averaged latency of fall readings per mouse in the three groups of mice

(wildtype, heterozygous, and homozygous) were combined respectively at 3-

month time intervals - 3, 6, 9, and 12 months. The rotarod data were statistically

compared between the wildtype and heterozygous H304R/+ and between

wildtype and homozygous H304R/R mice at each time point using Student’s t-

test (two-tailed distribution, p<0.05) on the GraphPad Prism 6.0 software.

2.3.4 Tail flick test

Tail flick test was used to quantify the pain sensitivity response of H304R mice to

thermal pain stimulus using Panlab tail flick meter (Harvard Apparatus #

LE7106). A mouse is placed in the restrainer provided with the equipment and

allowed to get habituated inside the restrainer for 2 minutes. Then the mouse tail

is placed under a beam of thermal light and the meter records the time (in

seconds) taken by the mouse to move its tail away from the source of thermal

pain stimulus. Tail flick test data was collected and compared between 1, 3 and

12 month wildtype, heterozygous H304R/+, and homozygous H304R/R male

mice.

Figure 2-6 Tail flick meter

Sensitivity and response to thermal pain stimulus in mice is measured by tail flick meter.

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2.4 Mouse dissections (sciatic nerve and dorsal root ganglia)

Tissue from the peripheral nervous system of mice was dissected out by surgical

procedures. Sciatic nerves and the dorsal root ganglia were collected from mice

of 3 month and 12 months of age.

2.4.1 Tissue harvesting and cryopreservation

Mice were euthanized by administering CO2 and cervical dislocation. Sciatic

nerve and lumbar dorsal root ganglia (DRG) samples were collected from both

left and right side of the mice using Leica S8 AP0 Stereo Microscope. Collected

tissue was fixed in 4% paraformaldehyde solution (EMS catalog#15710) for 30

minutes at 4°C. Then the tissue was left in 30% Sucrose solution (weight/volume)

overnight at 4°C. Next day, the tissue samples were embedded in labelled

histology molds (Ted Pella, Inc. catalog # 27118) containing Tissue-Tek® O.C.T.

Compound (Sakura Finetek USA Inc. catalog #25608-930). The tissue samples

were rapidly frozen on dry ice and stored at -80°C in labelled freezer boxes.

2.4.2 Cryosectioning and immunohistochemistry of tissue samples

Frozen sciatic nerve or lumbar DRG was cryosectioned in 10µm size thin

sections on Leica Cryostat (model# CM 1850) and collected on glass slides. The

samples were then washed in 1X PBS (Gibco) thrice for 5 minutes each to clean

away any O.C.T compound around the tissue. Waterproof boundaries were

drawn around each tissue sample with ImmEdge Hydrophobic Barrier PAP Pen

(Vector laboratories, catalog #H-4000). The tissue was permeabilized for 10

minutes with permeabilization buffer (0.5% Triton-X100). Then block buffer (10%

51

normal goat serum and 0.1% Triton-X100 in PBS) was added for 60 minutes on

the samples. Tissue samples were incubated with appropriate dilutions of

primary antibodies (table 2-1) overnight at 4°C. Next day, the primary antibodies

were washed with block buffer twice and then incubated with respective

secondary antibodies for 2 hours at the room temperature. Unbound antibodies

were washed with block buffer twice, followed by 1X PBS wash twice. The tissue

samples were then mounted with ProLong™ Gold Antifade Mountant with DAPI

(ThermoFisher Scientific, catalog# P36931) and sealed with a glass coverslip.

52

Table 2-1 Antibodies used for tissue immunohistochemistry

Primary antibody Labels Catalog number Dilution

used

Mouse monoclonal

to Beta-III tubulin

(βIII tubulin)

Neuronal soma

and axons

Abcam # 78078 1:200

Rabbit polyclonal

to Myelin Basic

Protein (MBP)

Myelinated axons Abcam # 40390 1:200

Secondary

antibody

Labels Catalog number Dilution

used

Rabbit polyclonal

to Glucose

Transporter-1

(GLUT-1)

Perineurial glial

cells in the sciatic

nerve’s

perineurium

Abcam # 15309 1:200

Alexa Goat anti

mouse conjugated

with 488

Against mouse

antibodies

ThermoFisher

Scientific # A11001

1:200

Alexa Goat anti

rabbit conjugated

with 546

Against goat

antibodies

ThermoFisher

Scientific # A110035

1:200

53

2.4.3 Microscopy and data analysis

Images of sciatic nerve and dorsal root ganglia tissues were captured on the

Leica M205 FA Fluorescence Microscope with Leica DFC3000G CCD camera.

The microscope settings and configurations were kept uniform between all the

samples to avoid variability (table 2-2).

Table 2-2 Microscopic settings used for imaging

False color Wavelength

(nm)

Exposure

time

Gain Intensity Magnification

Green 515 nm 139 ms 2.5x 60% 80X

Red 660 nm 139 ms 2.5x 40% 80X

Blue 440 nm 2.6 sec 2.5x 100% 80X

Transmittance none 139 ms 2.5x 45% 80X

Collected sciatic nerve images were then analyzed on the ImageJ software

(National Institutes of Health and the Laboratory for Optical and Computational

Instrumentation) to calculate the cross-sectional area of the nerve. Threshold

function was used to trace out the boundary of the nerve that was stained by the

perineural marker GLUT-1 antibody. The area inside the traced-out threshold

was measured using the ‘measure area’ function. The measured area (in mm2)

was collected and compared between the wildtype, heterozygous H304R/+, and

homozygous H304R/R mice.

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2.5 Motility analysis studies in primary H304R neurons

Primary neurons from the embryonic brains of H304R mouse model were

dissected out and grown in vitro. Intracellular motility parameters were recorded

in in vitro primary neurons to characterize the functions of dynein in H304R

mouse model.

2.5.1 Tissue harvesting of embryonic primary neurons

Timed pregnancy breeding was set-up between the heterozygous male and

female mice. The embryos were removed at embryonic day 16.5 from a

euthanized pregnant female mouse. Brains were removed from each embryo and

were dissected further to collect the cerebellar cortex and hippocampus from

each lobe of the brain. Leica S8 AP0 Stereo Microscope was used for embryonic

dissections. Cortices and hippocampi of wildtype, heterozygous (H304R/+), and

homozygous (H304R/R) were then pooled together respectively after the

genotype of the embryos was determined by the PCR based method. Cortical

and hippocampal neurons were isolated respectively by enzymatic digestion

technique with Papain (Worthington® Biochemical Corp# LS003118) and DNase

(Worthington® Biochemical Corp # LK003172) in Hibernate® E minus

Ca2+/Mg2+ media (BrainBits® # HECAMG500). Neurons were plated on

0.3µg/ml poly-D-Lysine (Millipore Sigma) and 10µg/ml Laminin (ThermoFisher

Scientific # 23017-015) coated 25mm Fisherbrand® glass coverslips (Catalog #

12-545-102) in 6-well plates. The neurons were incubated at 37°C in warm

Neurobasal™ media (ThermoFisher Scientific # 21103049) supplemented with

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100X Antibiotic-Antimycotic (Gibco™ # 15240062), 50X B-27™ Supplement

(Gibco™ # 17504044), and 100x GlutaMAX™ (Gibco™ # 35050061). The

neurons were half fed by removing and replacing half of the media with fresh

neuronal media every other day.

2.5.2 Live neuronal imaging with Rab7-GFP labeled vesicles

Cortical neurons of all 3 genotypes (wildtype, heterozygous (H304R/+),

homozygous (H304R/R)) were nucleofected with Rab7-GFP plasmid using

standard protocol on the Nucleofector™ 2b Device (Lonza # AAB-1001). The

nucleofected neurons were then plated on the glass coverslips and incubated at

37°C in warm NeurobasalTM media (ThermoFisher Scientific # 21103049). The

nucleofected neurons were half fed with fresh media every other day.

Live neuronal movies were captured on the Nikon® Eclipse Ti Inverted

Microscope with Photometrics® Evolve EMCCD camera (#512) at D.I.V 10. The

live cell movies were recorded using the Metamorph® Automation and Image

Analysis Software (Molecular Devices). The movies were made at 100x

magnification with 500 frames captured at the frame rate of 10 fps (frames per

second) with the appropriate green fluorescence filter cube. The longest visible

projection of the neuron was selected visually and was used for making neuronal

movies to capture the motility events inside the neuronal projections.

Motility analysis on movies of neurons containing Rab7-GFP labeled vesicles

was done using the Kymograph function of the Metamorph® image analysis

56

software (Molecular Devices) and described in the next section. Motility

parameters of Rab7-GFP labeled vesicles such as total number of movements,

run length (µm), run velocity (µm/sec), and the direction of vesicular movement

were calculated (figure 2-7). Motility data from different genotypes (wildtype,

heterozygous (H304R/+), homozygous (H304R/R) were compared using one-

way ANOVA test on the GraphPad Prism®6 software.

Kymograph analysis description

The following Figure 2-7 is provided to illustrate the various features of

kymographs. For this purpose, a hypothetical set of two vesicles is shown.

Stationary vesicle is shown as a blue circle and a moving vesicle is shown as a

red circle inside a cell (figure 2-7). The red vesicle is shown moving on a

microtubule track shown with an orange line and the red vesicle is moving from

point B to A. A kymograph line is drawn from the cell center to the periphery

shown in the black line. The kymograph analysis of the black line is shown on the

right in the form of a time (sec) vs distance (µm) graph. The blue stationary

vesicle that does not move has a constant line as shown in blue indicating zero

displacement on the time vs distance graph. In contrast, the red vesicle that

moves shows the displacement as a slant line indicating a “run” on the time vs

distance graph. The slope of the slant line gives the run velocity (µm/sec) and the

slope angle gives the direction of the movement (towards or away from the cell

body).

57

Figure 2-7 Schematic diagram of motility analysis using kymographs

Kymograph is a time vs distance graph generated to analyze the motility parameters on vesicles (blue circle = stationary vesicle, red circle = moving vesicle in motion).

58

2.5.3 Live neuronal imaging with Synaptophysin-RFP labeled vesicles

Cortical and hippocampal primary neurons were each infected with Bacmam

Synaptophysin RFP (2µl/ml) (ThermoFisher Scientific # C10610) and incubated

for 48 hours. Live neuronal movies were captured as mentioned in the sections

above. Motility analysis of neurons containing Bacmam Synaptophysin RFP

labeled vesicles was done using the Kymograph function of the Metamorph®

Automation and Image Analysis Software (Molecular Devices). Motility

parameters of Synaptophysin RFP positive vesicles such as total number of

movements, run length (µm), run velocity (µm/sec), and the direction of vesicular

movement were calculated. Motility data from different genotypes of mice

(wildtype, heterozygous (H304R/+), homozygous (H304R/R)) were compared

using one-way ANOVA test on GraphPad Prism®6 software.

59

CHAPTER 3: BEHAVIORAL PHENOTYPE CHARACTERIZATIONS OF H304R MICE

3.1 Introduction

Prior to our study, three other dynein heavy chain mutant mice were reported in

the literature – legs at odd angle (Loa), Cramping (Cra), and Sprawling (Swl).

Loa and Cra mice were generated using ENU-mediated mutagenesis (Hrabe de

Angelis et al., 2000) and Swl was created using radiation induced mutagenesis

(Duchen, 1974). All three mice displayed abnormal posture and hind limb

clenching during the tail suspension. These transgenic mice were considered to

be the paradigm mouse models of neurological dysfunction and were extensively

studied. Some of the phenotypic studies on Loa, Cra, and Swl mice are outlined

below.

A multi-stage protocol called SHIRPA (Smith/Kline/Harwell/Imperial

College/Royal Hospital/Phenotype Assessment) was used to quantitatively asses

the phenotype of the Loa mouse in a longitudinal study (Rogers et al., 2001). The

Loa/+ test results were compared with age and gender matched wildtype control

mice in the behavioral and functional assessments done from 1 to 16 months of

age. During the SHIRPA screening, the heterozygote Loa/+ and wildtype mice

were assessed on tail suspension, grip strength, wire maneuver, and limb tone at

1, 3 and 14-16 months age.

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For grip strength testing, mice were gently pulled across a steel mesh grip and

the grip strength exerted was recorded qualitatively with arbitrary scores: no

effort= 0, slight semi-effective effort= 1, moderate effective effort= 2, active

effort= 4, and unusual effective effort =5. For the wire maneuver test, mice were

suspended on a horizontal wire by their fore limbs and the efforts to bring their

hind limbs to the wire were scored as “falls immediately” to “active grip” with hind

limbs. Lastly, in the hind limb tone assessment, mice were restrained in the

supine position and their resistance to the pressure on the hind paws (plantar

side) was recorded as “slight” or “moderate” or “heavy” resistance. Overall, the

Loa/+ mice exhibited abnormal posture of hind limb clenching during tail

suspension, significant reductions in hind limb (median score of 1= slight grip),

reduced performance in the wire maneuver test (median score of 1= difficulty in

grasping with hind limbs), and deficits in hind limb tone (median score of 1= slight

resistance) as compared to the wildtype control mice.

For another stage of SHIRPA protocol, Loa/+ mice performed the rotarod test

and spontaneous locomotor activity test. For the rotarod test, the test mouse was

placed on a rotating rotarod and time when a mouse would fall off from a rotarod

was recorded. In locomotor activity test, a mouse was placed in a clear Perspex

activity cage fitted with infra-red signals, and the infra-red beam breaks induced

by the mouse locomotion was detected and recorded over a 30 minutes test

window. The Loa/+ mice had a significant reduction in its rotarod performance at

1, 3 and 16 months’ time points as they were unable to stay on the rotarod as

61

long as the control wildtype mice. During locomotion test, the Loa/+ mice

interestingly showed more locomotor activity than the wildtype control mice. In

conclusion, the SHIRPA protocol of behavioral assessments provides a

comprehensive methodology for documenting progressive neurological

dysfunction in mouse models (Rogers et al., 2001).

The other two DHC mutant mice – Cra and Swl were also characterized using

SHIRPA protocol for their neurological functions. Just like Loa/+ mice, both Cra/+

and Swl/+ mice displayed hind limb clenching during tail suspension, gait

abnormalities, no reductions in front grip strength, and significantly reduced grip

strength in hind limbs (Banks and Fisher, 2008; Chen et al., 2007). The

homozygous Loa/Loa, Cra/Cra, and Swl/Swl mice were either embryonically

lethal or did not survive 24 hours post birth due to their inability to feed (Chen et

al., 2007; Hafezparast et al., 2003). Overall, the DHC mutant Loa, Cra, and Swl

mice models provided some interesting evidences in the role of dynein in

neurological dysfunction.

3.2 Rationale

The mutagenesis generated Loa, Cra and Swl mice have provided many key

insights in understanding the dynein function, however, the mutations in Loa

(F8580Y), Cra (Y1055C), and Swl ([GIVT]1040[A]) are not linked to any

neurodegenerative disease alleles in humans. It is hard to correlate their key

findings in understanding the dynein function in the context of neurodegenerative

62

diseases in humans. Discovery of the first dynein mutation (H306R) in human

neuropathy (CMT2O) was an opportunity to study dynein function. Hence, we

generated the novel H304R mouse model as a tool to understand the dynein

function and mechanism in human neurodegenerative diseases.

The H304R mouse model has a mutation in its dynein heavy chain that is

reported to lead to human neuropathies such as Charcot-Marie-Tooth Type 2O

disease (CMT2O) and Spinal Muscular Atrophy with Lower Extremity Dominant

(SMA-LED). Both CMT2O and SMA-LED patients exhibit similar clinical

symptoms such as they display lower limb weaknesses, muscle wasting, skeletal

and joint deformities of foot, joint contractures, and problems in walking. CMT2O

patients also show a loss of sensation symptoms in addition to their motor

defects. The disease linked Histidine to Arginine substitution mutation is at

position 306 in the human dynein heavy chain, and the corresponding mutation is

at position 304 in mouse dynein heavy chain (Figure 3-1).

Figure 3-1 Comparison of dynein heavy chain peptide

CMT2O disease linked mutation is a missense substitution of Histidine (H) to Arginine (R) at position 306 in human dynein heavy chain peptide. The corresponding mutation is at position 304 in mouse dynein heavy chain peptide.

63

We generated the novel H304R mouse model using knock-in gene targeting

technique (Sabblah et al., 2018). It is critical to remember that because a mouse

has a disease linked mutation, it does not mean that it would be a relevant model

to study that disease. A mouse model has to recapitulate the clinical

manifestations of the disease to be considered as a relevant model for studying

that disease. Therefore, a key question for the H304R mouse model was if the

H304R mice would develop and display the symptoms associated with CMT2O.

To answer this question, we performed extensive studies to investigate and

characterize the behavioral phenotype of the H304R mouse model. Several

phenotypic assessments and motor-skills tests were performed at multiple time

points to investigate if the H304R mouse model recapitulated the corresponding

phenotype associated with CMT2O. First, we started with breeding and growing

the heterozygous H304R/+ colony for preliminary behavioral assessments. The

initial assessments such as tail suspension reflex, grip strength, and rotarod test

were performed with the heterozygous H304R/+ male and female mice and the

data was compared with the wildtype mice. The preliminary findings between the

heterozygous H304R/+ and wildtype mice were published first (Sabblah et al.,

2018).

After seeing a promising phenotype with the heterozygous H304R/+ mice, we

decided to start another behavioral skills study with the homozygous H304R/R

mice. The homozygous H304R/R mice were then bred and grown into a colony

of mice. We compared the tail suspension reflex, grip strength, and rotarod data

64

between the homozygous H304R/R mice and the wildtype control mice at

multiple ages (manuscript in submission).

3.3 Results

3.3.1 H304R mice had normal life span

The founder H304R knock-in mice were bred to yield both male and female

heterozygous H304R/+ mice. The heterozygous H304R/+ pups had a normal

birth and normal lifespan of 2 years (Sabblah et al., 2018). The heterozygous

H304R/+ mice did not appear to have any significant bodyweight differences and

were able to move around the cage just like their littermate wildtype control

animals.

The homozygous H304R/R pups were able to survive after birth unlike the other

dynein heavy chain homozygous mutant pups (Loa/Loa, Cra/Cra, Swl/Swl) and

had a normal lifespan (Chen et al., 2007; Hafezparast et al., 2003). The

homozygous H304R/R mice appeared to have gait defects such as a high

steppage gait in hind limbs (figure 3-2). Also, they were not as inclined to move in

the cage as compared to their littermate wildtype control and heterozygous

H304R/+ mice. They did not have any significant bodyweight differences

compared to their littermates.

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Figure 3-2 Gait phenotype of H304R mouse model

Homozygous H304R/R mice showed gait defects such as high steppage gait in hind limbs (black arrow).

3.3.2 H304R mice showed atypical tail suspension phenotype

Tail suspension tests were performed to assess the phenotype of H304R mouse

model. Mice tend to splay their hind limbs away from their body when lifted by the

tails as their typical tail suspension reflex. However, in some cases, mice would

clench their hind limbs together and hold them close to their body as part of their

atypical tail suspension phenotype (Figure 3-3).

66

Figure 3-3 Tail suspension phenotype of H304R mouse model

Typical tails suspension reflex consists of splaying of hind limbs away from the body, whereas, atypical tail suspension involves clenching of hind limbs together closer to the body.

The tail suspension reflex test was performed on the H304R colony at multiple

ages and the mouse’s phenotype was scored as typical or atypical reflex. The tail

suspension reflex was then statistically compared between the 3 genotypes

using the Chi-square test (p<0.05). We observed that about 20-30% of male and

female heterozygous H304R/+ mice started to display the atypical tail suspension

phenotype at later ages (9 and 12 months) as compared to the wildtype control

67

mice (Figure 3-4). A majority (60-80%) of male and female homozygous

H304R/R mice significantly displayed the atypical tail suspension reflex

compared to the wildtype control and heterozygous H304R/+ mice right from an

early age. The tail suspension data is summarized in Table 3-1.

Figure 3-4 Atypical tail suspension response of H304R mouse model

Male and female heterozygous H304R/+ mice (gray bars) showed atypical tail suspension reflex at later time points as compared to the wildtype mice (black bars) (Chi-square test p<0.05). Whereas, both male and female homozygous H304R/R mice (white bars) showed atypical tail suspension reflex at all the time points (Chi-square test p<0.05). Note- * is the statistical significance as compared to the wildtype only and # is the statistical significance as compared to the wildtype and H304R/+.

* * * *

#

# # #

# # # #

68

Table 3-1 Tail suspension reflex of H304R mice

Tail

refl

ex

Time point

Percentage of mice showing atypical tail suspension

reflex

Number of animals (n) p-value (Chi-square

test)

Wild-type H304R/+ H304R/R Wild-type

H304R/+ H304R/R Wild-type vs

H304R/+

Wild-type vs

H304R/R

Mal

e m

ice

3 months

7.5% 2.0% 61.7% 34 25 26 0.0745 <0.0001

6 months

5.8% 11.2% 76.6% 46 34 20 0.0642 <0.0001

9 months

9.6% 21.6% 74.7% 29 27 20 0.0120 <0.0001

12 months

12.2% 27.3% 79.6% 40 42 18 0.0002 <0.0001

Fem

ale

mic

e

3 months

10.0% 15.4% 66.3% 9 8 27 0.5434 <0.0001

6 months

8.0% 12.5% 62.6% 35 21 20 0.2771 <0.0001

9 months

11.3% 18.6% 70.9% 54 37 19 0.0314 <0.0001

12 months

11.3% 18.5% 70.7% 56 44 19 0.0350 <0.0001

69

3.3.3 Heterozygous H304R/+ mice show mild defects in their motor phenotype

Initial behavioral assessments were done with the male and female heterozygous

H304R/+ mice and the data was statistically compared with age and gender

matched wildtype control mice. The behavioral test results are as follows:

Heterozygous H304R/+ mice show reductions in grip strength

Standard grip strength assay was performed to assess the muscular strength in

all limbs and front limbs of heterozygous H304R/+ mice. The male heterozygous

H304R/+ mice showed a statistically significant reduction of about 10% in all

limbs grip strength at all ages as compared to the wildtype control mice

(Student’s t-test, p<0.05) (Figure 3-5). Female heterozygous H304R/+ mice

showed a statistically significant reduction of about 5% in all limbs grip strength

at only 6 and 9 month ages as compared to the wildtype control animals

(Student’s t-test, p<0.05).

Male heterozygous H304R/+ mice showed a statistically significant reduction of

about 7% in its front grip strength at later ages of 9 and 12 months as compared

to the wildtype mice (Student’s t-test, p<0.05). The female heterozygous

H304R/+ mice did not have any reductions in their front grip strength as

compared to the wildtype mice at any age. The grip strength data for male

H304R/+ and female H304R/+ mice are summarized in table 3-2 and table 3-3

respectively.

70

Figure 3-5 Grip strength phenotype of heterozygous H304R/+ mice

Both male and female heterozygous H304R/+ mice showed significant reductions in all limbs grip strength as compared to the wildtype mice (A & B). Only male heterozygous H304R/+ mice showed significant reductions in front limbs grip strength at later time points (C). The female heterozygous H304R/+ mice did not show any reductions in front grip strength as compared to the wildtype control mice (D). Note- * Student’s t-test, p<0.05.

71

Table 3-2 Grip strength in H304R/+ male mice

Test (male mice)

Time point

Mean ± std. dev (grams) Number of animals (n)

p-value (Student’s t-test)

Wild-type H304R/+ Wild-type

H304R/+ Wild-type vs

H304R/+

All limbs

3 months

212.5 ± 23.2 190.4 ± 26.8 50 43 0.0001

6 months

225.1 ± 30.8 201.0 ± 29.9 62 58 0.0001

9 months

230.6 ± 37.9 206.8 ± 33.4 41 42 0.0033

12 months

244.5 ± 32.1 225.3 ± 33.5 40 42 0.0096

Front limbs

3 months

99.2 ± 12.9 95.6 ± 17.4 50 43 0.2654

6 months

88.8 ± 14.3 83.8 ± 15.0 62 58 0.0667

9 months

89.8 ± 12.9 82.1 ± 11.1 41 42 0.0045

12 months

92.0 ± 11.9 86.1 ± 13.4 40 42 0.0362

72

Table 3-3 Grip strength in H304R/+ female mice

Test (female mice)

Time point





H304R/+

All limbs

3 months

175.5 ± 24.0 171.9 ± 22.0 51 35 0.4820

6 months

192.9 ± 15.9 185.7 ± 16.9 56 41 0.0324

9 months

209.8 ± 18.8 199.8 ± 16.5 55 41 0.0068

12 months

222.2 ± 22.2 216.4 ± 20.0 54 41 0.1843

Front limbs

3 months

91.0 ± 12.5 89.1 ± 12.2 51 35 0.4801

6 months

81.5 ± 9.8 81.8 ± 11.0 56 41 0.8959

9 months

81.3 ± 11.8 79.8 ± 11.1 55 41 0.5148

12 months

85.9 ± 13.4 85.4 ± 13.1 54 41 0.8681

73

Heterozygous H304R/+ mice show deficits in the rotarod performance

Rotarod test was performed to assess the motor coordination phenotype of

H304R/+ mice. Mice were placed inside the rotarod equipment and their falling

off from the rotarod before the completion of the rotarod cycle was measured as

latency to fall (in seconds). The male heterozygous H304R/+ mice were falling

from the moving rotarod quicker than the wildtype mice (Table 3-5). They showed

significant reductions in their latency to fall (by about 25%) as compared to the

wildtype control animals at 3, 6 and 9 months of age (Student’s t-test, p value

<0.05) (Figure 3-6). Female heterozygous H304R/+ mice also showed a similar

trend (Table 3-5), but had a significant reduction of 20% in their latency to fall

only at an early age of 3 months as compared to the wildtype mice (Student’s t-

test, p value <0.05).

Figure 3-6 Rotarod performance test of the H304R/+ mice

Male and female heterozygous H304R/+ mice showed significant defects in their rotarod performance as compared to the wildtype at some time points (*Student’s t-test, p<0.05).

74

Table 3-4 Rotarod performance in H304R/+ male mice

Test (male mice)

Time point Mean ± std. dev (seconds)

Number of animals (n)




H304R/+

Rotarod

3 months 78.8 ± 40.9 55.2 ± 30.4 51 43 0.0019

6 months 95.5 ± 49.1 73.0 ± 42.4 48 41 0.0223

9 months 101.9 ± 52.9 74.6 ± 44.8 28 33 0.0359

12 months 100.7 ± 48.6 82.4 ± 45.7 20 25 0.2035

Table 3-5 Rotarod performance in H304R/+ female mice

Test (female mice)






H304R/+

Rotarod

3 months 103.6 ± 47.6 82.7 ± 34.3 51 35 0.0205

6 months 113.7 ± 54.9 104.9 ± 45.2 51 35 0.4210

9 months 106.8 ± 61.4 105.9 ± 54.0 51 35 0.9386

12 months 98.8 ± 62.6 87.7 ± 51.3 50 35 0.3711

75

3.3.4 Homozygous H304R/+ mice show severe defects in their motor phenotype

After seeing the altered phenotype of motor skills in the heterozygous H304R/+

mice, we decided to perform behavioral assessments of the male and female

homozygous H304R/R mice. The data was statistically compared with age and

gender matched with the wildtype control mice. The behavioral test results are as

follows:

Homozygous H304R/R mice show reductions in grip strength

Standard grip strength assay was performed to assess the muscular strength in

all limbs and front limbs of heterozygous H304R/R mice. Male homozygous

H304R/R mice showed statistically significant reductions of about 30% in all

limbs grip strength as compared to the wildtype animals at all ages (Student’s t-

test, p value <0.05) (Figure 3-7). Female homozygous H304R/R mice also

showed statistically significant reductions of about 20% in all limbs grip strength

as compared to the wildtype animals at all ages tested (Student’s t-test, p value

<0.05). Both male and female homozygous H304R/R mice did not show any

reductions in their front grip strength as compared to the wildtype mice at all

ages. Grip strength data for male homozygous H304R/R (Table 3-6) and female

homozygous H304R/R (Table 3-7) mice are summarized in the table below.

76

Figure 3-7 Grip strength phenotype of heterozygous H304R/R mice

Both male and female homozygous H304R/R mice showed significant reductions in all limbs grip strength (A & B) as compared to the wildtype controls. No grip strength reductions were observed in the front limbs of both male and female homozygous H304R/R mice (C&D). Note- * Student’s t-test, p value <0.05.

77

Table 3-6 Grip strength in H304R/R male mice

Test (male mice)

Time point



Wild-type H304R/R Wild-type

H304R/R Wild-type vs

H304R/R

All limbs

3 months

212.5 ± 23.2 162.0 ± 46.3 50 26 <0.0001

6 months

225.1 ± 30.8 162.9 ± 31.4 62 20 <0.0001

9 months

230.6 ± 37.9 162.2 ± 38.6 41 20 <0.0001

12 months

244.5 ± 32.1 167.0 ± 31.3 40 18 <0.0001

Front limbs

3 months

99.2 ± 12.9 96.0 ± 23.6 50 26 0.9995

6 months

88.8 ± 14.3 84.6 ± 21.7 62 20 0.9970

9 months

89.8 ± 12.9 85.8 ± 18.8 41 20 0.9987

12 months

92.0 ± 11.9 84.1 ± 20.2 40 18 0.7910

78

Table 3-7 Grip strength in H304R/R female mice

Test (female mice)

Time point





H304R/R

All limbs

3 months

175.5 ± 24.0 154.5 ± 23.0 51 27 0.0004

6 months

192.9 ± 15.9 164.9 ± 28.5 56 20 0.0004

9 months

209.8 ± 18.8 161.0 ± 29.5 55 19 <0.0001

12 months

222.2 ± 22.2 173.2 ± 39.2 54 19 <0.0001

Front limbs

3 months

91.0 ± 12.5 91.5 ± 15.5 51 27 >0.9999

6 months

81.5 ± 9.8 81.7 ± 17.0 56 20 >0.9999

9 months

81.3 ± 11.8 81.6 ± 17.9 55 19 >0.9999

12 months

85.9 ± 13.4 86.5 ± 21.1 54 19 >0.9999

79

Homozygous H304R/R mice show deficits in the rotarod performance

Homozygous H304R/R mice were unable to keep up with the rotating rotarod

and were falling off the rod a lot quicker than the wildtype mice. Male

homozygous H304R/R mice showed statistically significant reductions in their

latency to fall as compared to the wildtype at all ages (Student’s t-test, p value

<0.05) (Figure 3-8). Similarly, female homozygous H304R/R mice also showed

statistically significant reductions in their rotarod performance as compared to the

wildtype mice at all ages tested (Student’s t-test, p value<0.05). Both male (Table

3-8) and female (Table 3-9) homozygous H304R/R mice showed about 65%

reduction in their latency to fall as compared to the wildtype mice.

Figure 3-8 Rotarod performance test of the H304R/R mice

Both male and female homozygous H304R/R mice showed significant defects in their rotarod performance as compared to the wildtype at all ages (* - Student’s t-test, p value <0.05).

80

Table 3-8 Rotarod performance in H304R/R male mice

Test (male mice)






H304R/R

Rotarod

3 months 78.8 ± 40.9 25.0 ± 9.6 51 26 <0.0001

6 months 95.5 ± 49.1 22.4 ± 9.7 48 20 <0.0001

9 months 101.9 ± 52.9 17.8 ± 9.6 28 20 <0.0001

12 months 100.7 ± 48.6 18.8 ± 9.7 20 18 <0.0001

Table 3-9 Rotarod performance in H304R/R female mice

Test (female mice)

Time point Mean ± std. dev (seconds) Number of animals (n)



H304R/R

Wild-type vs

H304R/R

Rotarod

3 months 103.6 ± 47.6 28.1 ± 9.1 51 27 <0.0001

6 months 113.7 ± 54.9 30.8 ± 18.7 51 20 <0.0001

9 months 106.8 ± 61.4 23.9 ± 12.0 51 19 <0.0001

12 months 98.8 ± 62.6 23.2 ± 9.8 50 19 <0.0001

81

3.4 Conclusions

We generated a novel H304R mouse model for Charcot-Marie-Tooth type 2O

(CMT2O) with a disease mutation in its dynein heavy chain. We performed

several phenotypic and behavioral assessments on male and female

heterozygous H304R/+ and homozygous H304R/R mice. The heterozygous

H304R/+ had a normal lifespan and survived until 2 years (end point of the

study). Similarly, the homozygous H304R/R mice had a normal lifespan unlike

the other dynein heavy chain mutant mice (Loa/Loa, Cra/Cra, and Swl/Swl) that

were embryonic or post-natal lethal (Chen et al. (2007); (Hrabe de Angelis et al.,

2000).

Male and female heterozygous H304R/+ mice showed an atypical tail suspension

reflex of hind limb clenching at later ages (9 and 12 months). The atypical tail

suspension phenotype could be indicative of motor dysfunction in lower

extremities (or hind limbs) of the mice. Male and female heterozygous H304R/+

mice showed significant reductions of 5-10% in their hind limbs grip strength.

Only male heterozygous H304R/+ mice showed significant reductions in the front

limbs grip strength at later ages (9 and 12 months). Additionally, male

heterozygous H304R/+ mice also showed significant ~25% reduction in the

rotarod performance almost at all the time points. Female heterozygous H304R/+

mice showed a significant ~20% reduction in the rotarod performance test at an

early age only (3 months).

82

Male and female homozygous H304R/R mice appeared to have defects in their

gaits and were inclined to move less inside the cage as compared to their

littermates. The majority of homozygous H304R/R significantly showed atypical

tail suspension reflex right from an early age 3 months to all the way up to 12

months. Male and female homozygous H304R/R mice also showed significant

20-30% reductions in all limbs grip strength as compared to the wildtype mice,

but not in front limbs grip strength. Both male and female homozygous H304R/R

mice also performed poorly on the rotarod performance test with ~65% reduction

in latency to fall because they were unable to stay longer on the moving rod than

the wildtype control mice.

So far, our initial investigation and characterization of the novel H304R mice with

CMT2O mutation in DHC offered the following insights:

The H304R mouse model is a novel mouse model with a mutation in the

dynein heavy chain that is known to cause neuropathies in humans

(CMT2O and SMA-LED) (Tsurusaki et al. (2012); (Weedon et al., 2011).

The H304R mouse model was able to recapitulate and express the

symptoms of human neuropathies (CMT2O and SMA-LED) such as

muscle weaknesses, and motor coordination defects.

The heterozygous H304R/+ mutation expressed and recapitulated the

heterogeneity of motor phenotype in mice, similar to the heterogeneity of

83

clinical symptoms seen in humans with the autosomal dominant CMT2O

mutation.

Quantitatively, the heterozygous H304R/+ mice displayed a milder

reduction in its motor skills phenotype, whereas, the homozygous

H304R/R mice displayed a severe reduction in its motor skills phenotype.

The male heterozygous H304R/+ mice appeared to be affected more

significantly by the mutant dynein function than the female heterozygous

H304R/+ mice.

The male heterozygous H304R/+ mice had a variable onset of the motor

phenotype, sometimes at an early age (reductions in all limbs grip strength

and rotarod tests) and sometimes at a later age (atypical tail suspension

and reduced front limbs grip strength).

Both male and female homozygous H304R/R mice appeared to be

affected equally by the mutant dynein function.

The homozygous H304R/R mice consistently had an early onset of motor

phenotype that was severely deficient right from the start. In addition, the

homozygous H304R/R mutation phenotype was also static and non-

progressive in nature, i.e. it was not worsening drastically as the mice

aged.

A dynein molecule contains a dimer of two DHC monomeric peptides

linked together. A unique advantage of the homozygous H304R/R

mutation is that the homozygous mice will have a uniform composition of

84

dynein molecules with DHC dimers containing both mutant monomers. In

contrast, the heterozygous H304R/+ mutation leads to a mixed pool of

dynein in heterozygous mice, i.e. a quarter population of DHC dimers will

have both wildtype monomers, a quarter of DHC with both mutant

monomers, and half of the DHC dimers with one wildtype and one mutant

monomer.

In conclusion, the H304R mouse model appears to serve as a useful tool to

understand the role of dynein in neurodegenerative diseases. Future biochemical

studies with H304R mice can unveil the molecular mechanism of the mutant

dynein function in neurons.

85

CHAPTER 4: SENSORY PHENOTYPE CHARACTERIZATIONS OF H304R MICE

4.1 Introduction

Many CMT patients show the symptoms of sensory neuropathy along with motor

neuropathy. Common sensory deficits reported in CMT patients include loss of

sensation in distal limbs, paresthesia (tingling sensation), reduced

proprioception, and some patients also displayed reduction in their fine touch and

vibration sense (Weedon et al., 2011). Due to loss of sensation in their lower

limbs, some patients lose sensitivity to pain due to any foot wounding.

Consequently, that further leads to complications of foot ulcerations and usually a

follow up surgery is needed to remove the damaged foot/toes. It is worth noting

that previously published dynein heavy chain mutant mice (Loa, Cra and Swl)

reported sensory deficits in their phenotype (Chen et al., 2007; Dupuis et al.,

2009) as outlined below.

4.1.1 Sensory deficits in previously published mutant dynein mice models

The sciatic nerve is the largest nerve in the body, and it innervates the lower limb

muscles of mice. The outer layer of dense irregular connective tissue around a

sciatic nerve is called epineurium (Figure 4-1). The sciatic nerve is made up of

fascicles i.e. the several bundles of neuronal axons that are enclosed together.

The outer layer of a fascicle is lined with perineural glial cells and is called the

perineurium. The extracellular matrix space between neuronal axons inside a

fascicle is called the endoneurium.

86

Figure 4-1 Sciatic nerve cross section

The sciatic nerve is covered with a layer of connective tissue called epineurium and contains several nerve bundles called fascicles. The fascicles are lined with glial cells that make up a layer called perineurium. The extracellular space around the bundles of axons inside the fascicles is called endoneurium.

Deficits in the sciatic nerve anatomy have been linked to sensory dysfunction in a

previously reported dynein heavy chain mutant mouse (Chen et al., 2007). Both

heterozygote Loa/+ and heterozygote Swl/+ mice reportedly had thinner sciatic

nerves as compared to wildtype animals. Both ventral root and dorsal root

contribute their spinal nerves (neuronal axons) to make up the sciatic nerve. It

was observed and determined that the dorsal root size was smaller in both

heterozygote Loa/+ and Swl/+ mice as compared to wildtype controls, whereas

the ventral root size was unchanged. The Cra/+ mice also showed thinner dorsal

roots and loss of large caliber axons (Dupuis et al., 2009)

87

A dorsal root ganglion is a cluster of neuronal cell bodies or somas of pseudo-

unipolar sensory neurons. The dorsal root ganglia (DRG) receive input signals

from sensory neurons innervating target tissue such skin, muscles, and organs.

Input nerve signals travel along the sensory neuron axons in ‘afferent direction’

i.e. from target tissue to dorsal root ganglia and further passed onto the spinal

column for signal processing. Additionally, the dorsal root ganglion is composed

of different neuronal subpopulation involved in sensory stimulus reception. For

example, nociceptive neuronal subpopulation inside a dorsal root ganglion is

responsible for sensing pain stimulus from the target tissue such as skin. Another

example inside a dorsal root ganglion is the proprioceptive neuronal

subpopulation that is involved in sensing the muscle stretch and orientation.

In the study mentioned above, the DRG in heterozygote Loa/+ and heterozygote

Swl/+ mice were also examined, and they revealed interesting insights into the

DRG neuronal subpopulation composition (Chen et al., 2007). It was determined

that the total number of neurons were significantly reduced in dorsal root ganglia

from lumbar (L4) spinal region of heterozygote Loa/+ and heterozygote Swl/+

mice. Specifically, it was determined that proprioceptive neuronal subpopulations

were reduced by 69% in the dorsal root ganglia (L4) of newly born heterozygote

Swl/+ animals. Proprioceptive neuronal subpopulation count was not studied in

heterozygote Loa/+ mice. In conclusion, the sensory studies on heterozygote

Loa/+ and Swl/+ mice laid out the basis for investigating the sensory phenotype

of the H304R mouse model.

88

4.2 Rationale

We focused on the motor behavior skills characterization of the novel H304R

mouse in our initial studies and observed deficits in the tail suspension, grip

strength and the motor coordination phenotype of the mice. Here I start

investigating the sensory phenotypes of H304R mice to understand and compare

those to the sensory phenotypes of previously published dynein heavy chain

mutant mice (Loa/+ and Swl/+).

First, I started with assessing the sensory behavior phenotype by exposing

wildtype, heterozygous H304R/+, and homozygous H304R/R mice (both male

and female) to the tail flick sensitivity test and recorded their response to pain

stimulus at early (1 month and 3 months) and a later (12 months) time point.

Based on the tail flick data, I selected the male H304R mice cohort at 3 and 12

months’ time points for further sensory tissue investigation. I examined the gross

anatomy of sciatic nerve in the wildtype, heterozygous H304R/+, and

homozygous H304R/R male mice at 3 and 12 months of age and compared its

thickness in all three genotypes. I dived deeper into quantitative analysis of

sciatic nerve thickness by comparing the total fascicular area in the sciatic nerve

from the male H304R mice of all three genotypes. Because the dorsal root

ganglia (DRG) contribute its axons to the sciatic nerve, I refined and optimized

conditions to study the DRG composition to further understand the sciatic nerve

phenotype of the H304R mouse model.

89

4.3 Results

4.3.1 Altered latency to respond to pain stimulus in H304R mice

The tail flick test was used to measure the sensitivity of H304R mice to thermal

pain stimulus. A test mouse was restrained safely in a restrainer and its tail was

briefly exposed to a thermal light beam as a source of pain stimulus at a

moderate intensity setting (50 out of 99 on the setting scale). The tail flick meter

recorded the time (in seconds) that a mouse would take to remove its tail out of

the thermal light beam path (pain stimulus). Both male and female cohorts of

H304R mice were tested on the tail flick test at multiple time points and their data

was compared respectively between the three genotypes.

Figure 4-2 Tail flick pain sensitivity test of H304R male mice

Male homozygous H304R/R mice displayed a significantly quicker response to the pain stimulus at 3 months and 12 months of age as compared to their respective wildtype control male mice (One-way ANOVA, p<0.05). Only 3 month female homozygous H304R/R mice displayed a significant quicker response to the pain stimulus as compared to the wildtype control female mice (One-way ANOVA, p<0.05). 12 month old female homozygous H304R/R data is not presented due to the unavailability of mice.

Male mice Female mice

HH

HR

RR

0

5

1 0

1 5

1 m o n th m a le ta il f lic k

Tim

e t

o r

es

po

nd

(s

ec

)

HH

HR

RR

0

5

1 0

1 5

3 m o n th m a le ta il f l ic k w /o s in g le g e n o

Tim

e t

o r

es

po

nd

(s

ec

)

HH

HR

RR

0

5

1 0

1 5

3 m o n th fe m a le ta il f l ic k w /o s in g le g e n o

Tim

e t

o r

es

po

nd

(s

ec

)

HH

HR

RR

0

5

1 0

1 5

1 m o n th fe m a le ta il f lic k

Tim

e t

o r

es

po

nd

(s

ec

)

* *

HH

HR

RR

0

5

1 0

1 5


Tim

e t

o r

es

po

nd

(s

ec

)

*

HH

HR

0

5

1 0

1 5


Tim

e t

o r

es

po

nd

(s

ec

)

Res

po

nse

tim

e (s

ec)

Res

po

nse

tim

e (s

ec)

90

Interestingly, both heterozygous H304R/+ and homozygous H304R/R male mice

took less time to respond to the pain stimulus than the wildtype control mice

(Figure 4-2) at multiple time points. However, the quick response time was

statistically significant only in the homozygous H304R/R male mice at 3 months

and 12 months of age (One-way ANOVA, p<0.05) as compared to the wildtype

mice. There was no significant difference between the pain response time of

heterozygous H304R/+ and homozygous H304R/R male mice at any time points

tested (Table 4-1).

Female heterozygous H304R/+ and homozygous H304R/R mice did not show

any significant differences in their tail flick response times as compared to the

wildtype control female mice at 1 month of age. However, at 3 months of age,

both heterozygous H304R/+ and homozygous H304R/R female mice took lesser

time to respond to the pain stimulus. But only homozygous H304R/R female mice

had a statistically significantly quicker response time as compared to the wildtype

control mice (One-way ANOVA, p<0.05). The tail flick data for 12 month old

female homozygous H304R/R mice is not presented due to a lack of the mouse

availability. There was no significant difference between the pain response time

of heterozygous H304R/+ and homozygous H304R/R female mice at all time

points.

91

Table 4-1 Tail flick data

Average time of response (sec), number of animals (n), and the one-way ANOVA test p-values (with Tukey’s multiple

comparison test) for the wild-type, heterozygous (H304R/+) and homozygous (H304R/R) mice

Tail

flick

test

Time

point

(age)

Average time of response

to the pain stimulus

(seconds)

Number of animals (n) p-value (Tukey’s multiple

comparison)

Wild-

type

H304R/+ H304R/R Wild-

type

H304R/+ H304R/R Wild-

type vs

H304R/+

Wild-type

vs

H304R/R

H304R/+

vs

H304R/R

Ma

le m

ice 1 month 8.13 7.28 4.46 3 13 2 0.9051 0.4203 0.4743

3 months 8.64 6.40 5.21 7 18 6 0.0843 0.0284 0.5117

12 months 8.43 7.17 4.48 4 6 5 0.6272 0.0370 0.1227

fem

ale

mic

e 1 month 7.60 7.77 7.86 5 9 4 0.9876 0.9796 0.9968

3 months 8.43 7.04 4.59 9 12 4 0.3108 0.0163 0.1335

12 months 10.43 7.83 N/A 4 4 N/A N/A N/A N/A

92

4.3.2 Sciatic nerve appeared thinner in H304R mice

The sciatic nerve is the largest nerve in the body and it innervates the lower

limbs of mice. The sciatic nerve arises from spinal nerves L3, L4, and L5 from the

lumbar spinal column in mice. Dissections were performed on euthanized

wildtype, heterozygous H304R/+, and homozygous H304R/R male mice to

expose & collect both left and right sciatic nerves. Then, the tissue was frozen,

sectioned and stained with immunohistochemistry-based methods to compare

the sciatic nerve thickness between the three genotypes.

Sciatic nerve dissections were done on two litters each of 3 months and 12

month old wildtype, heterozygous H304R/+, and homozygous H304R/R male

mice. Visually, it appeared that both the left and right sciatic nerves were thinner

in the male homozygous H304R/R mice as compared to the wildtype and

heterozygous H304R/+ littermate control (Figure 4-3). Heterozygous H304R/+

mice appeared to have sciatic nerves of similar or slightly reduced thickness as

compared to the wildtype littermate control animals. These visual observations of

sciatic nerve were then further validated by quantitatively measuring the nerve

thickness and axonal counts using immunohistochemistry-based staining

methods as described next.

93

Figure 4-3 Comparison of sciatic nerve thickness of H304R male mice

The sciatic nerve appeared to be thinner in 3-month homozygous H304R/R male mice as compared to the wildtype and heterozygous H304R/+ littermates.

4.3.3 Reduced total sciatic nerve fascicular area in H304R mice

After seeing apparent differences in sciatic nerve thickness of H304R mice, I

wanted to quantify this observed difference by measuring the total fascicular

area. The sciatic nerve is made up of fascicles or bundle of neuronal axons

enclosed within the perineurium layer. Fascicular area was calculated by

measuring the area enclosed inside perineurium layer of fascicles that was

stained by a perineural marker - glucose transporter GLUT-1 antibody (details in

Chapter 2: Materials and methods). Total area of all the fascicles were added

together and compared between the 3-month (Figure 4-4) and 12-month (Figure

4-5) old wildtype, heterozygous H304R/+, and homozygous H304R/R male mice

respectively.

Wild-type

H304R/+

H304R/R

94

Figure 4-4 Sciatic nerve cross sections of 3-month H304R male mice

Sciatic nerve cross sections were stained with perineural glial maker – GLUT 1 (shown here in red) to label individual fascicles. Total fascicular area was calculated for all fascicles and compared between the genotypes. Homozygous H304R/R mice showed a significant reduction in the total fascicular area as compared to wildtype littermate control. No significance differences were observed between heterozygous H304R/+ and wildtype mice.

Wild-type H304R/+ H304R/R

Litter #1

Litter #2

95

Figure 4-5 Sciatic nerve cross sections of 12-month H304R male mice

Sciatic nerve cross sections were stained with perineural glial maker – GLUT 1 (shown here in red) to label individual fascicles. Total fascicular area was calculated for all fascicles and compared between the genotypes. Homozygous H304R/R mice showed a significant reduction in the total fascicular area as compared to wildtype littermate control. No significance differences were observed between heterozygous H304/R+ and wildtype mice.

Wild-type H304R/+ H304R/R

Litter #1

Litter #2

96

By visual observation, the sciatic nerve appeared thinner in homozygous

H304R/R male mice as compared to the wildtype littermate control animal at both

3 months and 12 months of age. This result was quantitatively confirmed by

observing a significant overall reduction of about 40% in total fascicular area of

homozygous H304R/R mice as compared to the wildtype control (One-way

ANOVA, Tukey’s multiple comparisons test, p<0.05) at 3 months and 12 months

age (Figure 4-6). Heterozygous H304R/+ mice also showed about 16% reduction

in the total fascicular area as compared to the wildtype littermate control animal

at 3 months and 12 months age, but this difference was not statistically

significant. Total fascicular area differences between homozygous H304R/R and

heterozygous H304R/+ male mice were not significant (Table 4-2).

Figure 4-6 Total fascicular area of sciatic nerve in H304R male mice

Homozygous H304R/R male mice had a statistically significant reduction of ~40% in its total fascicular area as compared to the wildtype control mice (1-way ANOVA, Tukey’s multiple comparisons test, p<0.05).

0

0.05

0.1

0.15

0.2

3 months 12 months

Tota

l fas

cicu

lar

area

(m

m2)

Sciatic nerve thickness

Wildtype

H304R/+

H304R/R

* *

97

Table 4-2 Total fascicular area in the sciatic nerve of H304R male mice

Total

fascicular area

Wildtype Heterozygous

H304R/+

Homozygous

H304R/R

3-month

littermate # 0.20 mm2 0.16 mm2 0.12 mm2

3-month

littermate #2 0.16 mm2 0.14 mm2 0.09 mm2

Average 3 -

month area 0.18 mm2 0.15 mm2 0.11 mm2

Total

fascicular area

Wildtype Heterozygous

H304R/+

Homozygous

H304R/R

12-month


12-month


Average 12 -

month area 0.18 mm2 0.15 mm2 0.11 mm2

98

4.3.4 Axonal counts in the sciatic nerve cross section

After seeing a reduction in the total fascicular area of the sciatic nerve, it was

apparent that the axon numbers inside the fascicles were needed to be counted

to reveal any alterations that may have led to this reduction. Alterations in total

fascicular area could be a result of reduced number of the axons or a reduction in

the myelination that could ultimately lead to the thinner nerves of heterozygous

H304R/+ and homozygous H304R/R mice.

The largest βIII tubulin stained fascicle from each genotype of 3-month littermate

male mice was selected (Figure 4-7). The axons were counted and recorded in

two categories based on their appearances – single axons and bundled axons. A

small and uniform green dot was counted as a single axon, while the larger

shaped green dot was counted as a bundle of multiple axons inside the fascicle.

Then the percentage of bundled axon over total axons and the percentage of

single axons over total axons were calculated and compared separately for all

three genotypes (Table 4-3). The preliminary results revealed a reduction in the

number of axons in both the heterozygous H304R/+ and homozygous H304R/R

mice (Figure 4-8). However, the percentage distribution of the total axons into

bundled axons or single axons categories were similar in all three genotypes

(Figure 4-9).

99

Figure 4-7 The largest fascicle inside the sciatic nerve of each genotype

The cross-sectional area of the largest fascicle inside each sciatic nerve was stained with the βIII tubulin antibody. Each green dot was counted as a single axon and a bunch of green dots were counted as a bundle of multiple axons.

Wildtype H304R/+ H304R/R

100

Table 4-3 Axon counts inside the largest fascicle

3 months Wildtype H304R/+ H304R/R

Bundled axons 629 461 362

Single axons 579 397 282

Total axons 1208 858 644

% of bundled axons 52.07% 53.73% 56.21%

% of single axons 47.93% 46.27% 43.79%

Figure 4-8 Total count of axons inside a fascicle

The total number of axons was counted in the cross-sectional area of the largest fascicle inside each sciatic nerve.

101

Figure 4-9 The percentage distribution of bundled axons vs single axons

The distribution of the total axons in the categories – bundled axons and single axons were similar in all three genotypes.

102

4.3.5 Identifying subpopulations of neurons in the adult DRG

After observing a significant reduction in sciatic nerve thickness and axonal count

in H304R mice, I wanted to determine if there was a corresponding alteration in

the dorsal root ganglia (DRG) composition. The DRG contribute its neuronal

axons to make up the sciatic nerves. The DRG contain a mixed subpopulation of

neurons such as nociceptive neurons (to sense pain) and proprioceptive neurons

(to sense muscle stretch and orientation). DRGs collected from the lumbar (L4)

region of wildtype, heterozygous H304R/+, and homozygous H304R/R male mice

were sectioned serially every 10µm and stained with anti-βIII tubulin antibody to

label all the neuronal somas (Figure 4-10).

103

Figure 4-10 Serial sections of dorsal root ganglia of H304R mice

The dorsal root ganglia from L4 lumbar region were serially sectioned. Best matched sections shown here in blue and orange boxes were picked for further analysis between the wildtype, heterozygous H304R/+ and H304R/R male mice.

Wil

dty

pe

H

30

4R

/+

H30

4R

/R

104

Best matched sections of L4 dorsal root ganglia were picked from wildtype,

heterozygous H304R/+, and homozygous H304R/R male mice for further staining

with neuronal subpopulation specific makers (Figure 4-11). I successfully

optimized the working conditions to stain dorsal root ganglia sections with anti-

CGRP antibody to label nociceptive neurons and with anti-Parvalbumin (PV)

antibody to label the proprioceptive neurons (Figure 4-11). This optimized

methodology (Table 4-4) can be further used to stain and count the anti-PV

positive neurons (proprioceptors) and the anti-CGRP positive neurons

(nociceptors).

Table 4-4 Optimized conditions for identifying DRG neuronal subpopulations

Primary antibody

Working concentration Notes

Mouse anti- βIII tubulin 1:200 Primary antibody

dilutions are to be

prepared in the Block

buffer prepared with

Bovine Serum Albumin

(BSA).

Overnight incubation at

4°C.

Goat anti-CGRP 1:100

Rabbit anti- parvalbumin 1:100

Secondary antibody

Working concentration Notes

Donkey anti- mouse 350 1:200 Goat secondary

antibody dilutions are to

be prepared in the Block

buffer prepared with

Bovine Serum Albumin

(BSA).

3 hours incubation at

room temperature.

Donkey anti- goat 488 1:100

Donkey anti- rabbit 568 1:200

105

Counting subpopulation numbers of anti-PV positive neurons and anti-CGRP

positive neurons can be used to calculate and compare the percentage of each

subpopulation over total neurons present in the DRG of H304R mice. Preliminary

data for the wildtype DRG is presented in table 4-5. Out of all the βIII tubulin

stained neurons, ~ 36% of the neurons were anti-PV positive and ~59% of the

neurons were anti-CGRP positive in the same section of the wildtype DRG.

Table 4-5 Preliminary counts of wildtype DRG subpopulation

Subpopulation Counts

Total cell bodies 39

anti-PV positive cell bodies 14

anti-CGRP positive cell bodies 23

% anti-PV positive cell bodies 35.90%

% anti-CGRP positive cell bodies 58.97%

106

Figure 4-11 Neuronal subpopulations staining of dorsal root ganglia

The dorsal root ganglia section from wildtype mice was stained with anti-βIII tubulin to identify all sensory neurons. The section was co-labelled with anti-parvalbumin and anti-CGRP primary antibodies to identify proprioceptive and nociceptive neuronal subpopulations respectively.

107

4.4 Conclusions

Charcot-Marie-Tooth patients exhibit sensory neuropathy symptoms in addition

to the motor neuropathy symptoms. In parallel, previously published dynein

heavy chain mutant Loa/+, Cra/+ and Swl/+ mice have also reported sensory

deficits (Chen et al., 2007; Dupuis et al., 2009). Based on the reports in human

CMT patients and previously published mutant dynein mice, I initiated the

sensory phenotype assessments of the H304R mouse model.

First, I started with characterizing the pain sensitivity response of H304R mice

with the tail flick test and found that both male and female homozygous H304R/R

mice had a statistically significant quicker response time to the pain stimulus than

the wildtype mice at 3 months of age. Male homozygous H304R/R mice also

showed a statistically significant quicker response time at 12 months of age as

compared to the wildtype mice. However, due to unavailability of female

homozygous H304R/R mice, no data was presented for comparison at 12

months. Heterozygous H304R/+ male mice also showed a quicker response to

tail flick test but didn’t have any significant differences as compared to the

wildtype control or homozygous H304R/R littermate mice at multiple time points.

It was previously reported that the heterozygote Loa/+ mice took a longer time to

respond to the tail flick test (unpublished data, (Banks and Fisher, 2008)). In

contrast, the H304R mutant mice took lesser time to respond on the tail flick test.

These contrasting observations indicate that the H304R mutation in the dynein

108

heavy chain might be affecting nociceptive neurons differently than the Loa

mutation (F580Y) in the dynein heavy chain.

Next, a closer examination of sciatic nerve anatomy revealed that homozygous

H304R/R mice had thinner sciatic nerves as compared to both the wildtype and

heterozygous H304R/+ littermate mice. Quantitatively homozygous H304R/R

mice had an overall reduction of ~40% in total fascicular area of its sciatic nerve

as compared to the wildtype littermates at 3 months and 12 months of age.

Heterozygous H304R/+ mice displayed a slight 16% reduction in the total

fascicular area at 3 months and 12 months of age, but this difference was not

statistically significant as compared to the wildtype littermate control animals.

Reductions in the sciatic nerve thickness of H304R mice were similar to the

observations of thinner sciatic nerves in the Loa/+ and Swl/+ mice (unpublished

data, (Banks and Fisher, 2008)). Repetition of sciatic nerve experiments needs to

be performed on more number of animals to confirm this preliminary observation.

Observations of the sciatic nerve appearing thinner and reduction in the total

fascicular area in the heterozygous H304R/+ and homozygous H304R/R mice

opened many interesting follow up questions. That is why counting the number of

axons inside the largest fascicle was the next key step to determine the potential

reason behind the observed reduction in the total fascicular area. The preliminary

data from a littermate set revealed that there was a reduction in the number of

the axons in both the heterozygous H304R/+ (~29%) and homozygous H304R/R

(47%) mice as compared to the wildtype control. However, the percentage

109

distribution of the bundled axons vs single axons was same in the fascicles from

all three genotypes. Reduced number of axons could be a potential reason

behind the thinner sciatic nerves inside the mutant H304R mice; however, further

investigations are necessary to establish this preliminary observation.

Because the sciatic nerve is a “mixed nerve”, it receives axons from both the

dorsal root (sensory neurons) and ventral root (motor neurons) of the spinal

column. The reduction in the diameter of sciatic nerve could be due to the loss of

axonal projections from the dorsal root and/or the ventral root. In order to answer

this question, I picked the sensory route or the dorsal root ganglia (DRG) to study

further.

As mentioned previously in this chapter, DRG has a mixed neuronal

subpopulation of sensory neurons that could be identified for quantitative

analysis. I successfully optimized the working conditions to label and identify the

neuronal subpopulation in the DRG of H304R mice. Future experiments using

these optimized conditions can be used to label all neurons with the βIII-tubulin

marker, proprioceptive neurons with the parvalbumin marker, and nociceptive

neurons with the calcitonin gene related protein (CGRP) marker. Next, these

labelled neuronal subpopulations can be quantitatively analyzed to calculate the

percentage of proprioceptive or nociceptive subpopulation over all the neurons

inside a DRG. Preliminary calculations with the wildtype DRG showed that ~36%

of all neuronal cell bodies were anti-parvalbumin positive cell bodies and nearly

~59% of all neurons were anti-CGRP positive cell bodies in the same stained

110

section of the DRG. Further quantitative subpopulation comparisons may reveal

some key differences in the composition of DRG in the three genotypes of the

H304R mice.

Overall, my preliminary work to characterize the sensory phenotype of H304R

mice revealed some interesting differences in the pain sensitivity response and

sciatic nerve anatomy of mutant heterozygous H304R/+ and homozygous

H304R/R mice. The sciatic nerve anatomy of mutant heterozygous H304R/+ and

homozygous H304R/R mice was also comparable with previously published

dynein mice (Loa/+ & Swl/+) as outlined above. Further in-depth studies with a

greater number of animals will be required to establish the preliminary

observation of reduced sciatic nerve thickness in the H304R mice. Subsequent

studies exploring and comparing the DRG neuronal subpopulations between the

three genotypes would be needed to get a deeper explanation of the preliminary

observations in the sensory phenotype of H304R mice.

111

CHAPTER 5: MOTILITY STUDIES OF H304R MICE

5.1 Introduction and rationale

Dynein is the key motor protein involved in the retrograde transport of cargoes

along the neuronal axon (Hirokawa et al., 1990). Direct role of dynein has been

established in the retrograde transport of neurotrophin mediated trk receptors

towards the neuronal soma for survival response (Heerssen et al., 2004; Yano et

al., 2001), fast axonal transport (Waterman-Storer et al., 1997), neurite growth

and organization (Ahmad et al., 1998), and neuronal migration (Tanaka et al.,

2004). Therefore, it is not surprising that the mutations in dynein have been

reported to show the impaired axonal transport in the DHC mutant Loa/+ mice

(Ori-McKenney et al., 2010) and in Swl/+ mice (Zhao et al., 2016).

Hence, we were also interested in investigating the intracellular motility

phenotype of H304R mouse model. Because previously studied DHC mutant

Loa/Loa mice showed defects in the development of its cortical and hippocampal

regions (Ori-McKenney and Vallee, 2011), we chose neurons from these areas

for our initial motility studies. We started with isolating cortical and hippocampal

neurons from H304R embryos (at E16.5) because embryonic neurons are easier

to grow in vitro than the adult mice neurons. Motility studies with established

dynein cargoes such as synaptophysin containing synaptic vesicles (Li et al.,

2000) and Rab7 positive vesicles (Johansson et al., 2007) were performed in

cortical and hippocampal embryonic neuronal cultures.

112

5.2 Results

5.2.1 Synaptophysin-RFP positive vesicular motility in hippocampal neurons

Hippocampal neurons from H304R mice were isolated and plated on coverslips

for intracellular motility experiments. The neurons from the same genotype were

pooled together independent of the gender of the embryo. Hippocampal neurons

were then infected with BacMam synaptophysin-RFP and were imaged at the

ages DIV 10 and 14 (more details in the methods chapter section 2.5). Various

parameters such as the number of movements in the inward (towards the cell

body) and outward (away from the cell body) directions, run length or distance

(µm), and velocity (µm/sec) were recorded of the synaptophysin-RFP positive

vesicles that were moving. The data was then analyzed and statistically

compared between the three genotypes. The wildtype mice are referred as HH,

the heterozygous mice are referred as HR, and the homozygous mice are

referred as RR in this chapter.

At DIV 10, synaptophysin-RFP positive vesicles in homozygous H304R/R mice

showed an unusual distribution of movement patterns i.e. about 83% of vesicles

were moving in the inward direction as compared to about 45% of inward

direction bound vesicles in both the wildtype and heterozygous H304R/+ mice

(Table 5-1). Subsequently, about 17% of vesicles were outward direction bound

in the homozygous H304R/R mice as compared to about 55% outward direction

vesicles in both the wildtype and heterozygous H304R/+ mice.

113

Table 5-1 Synaptophysin-RFP positive vesicles motility in hippocampal neurons at DIV 10

Furthermore, both inward and outward bound synaptophysin-RFP positive

vesicles in heterozygous H304R/+ mice showed a slight (but insignificant)

increase in the distance (µm) covered as compared to the wildtype and

homozygous H304R/R mice (Figure 5-1). Both heterozygous H304R/+ and

homozygous H304R/R mice showed a slight reduction in the velocity (µm/sec) of

the inward and outward bound synaptophysin-RFP positive vesicles as

compared to the wildtype mice, but this reduction was not found to be statistically

significant (Figure 5-1).

114

Figure 5-1 Synaptophysin-RFP positive vesicles motility in hippocampal neurons at DIV 10

The graphs show the distance (µm) covered by the vesicles in the inward (A) and outward (B) bound directions. The graphs show the velocity (µm/sec) of the vesicles in the inward (C) and outward (D) bound directions. Wildtype (black bars), heterozygous H304R/+ mice (gray bars), and homozygous H304R/R mice (blue bars).

At DIV 14, synaptophysin-RFP positive vesicles in homozygous H304R/R mice

showed an altered distribution of movement patterns. About 32% of

Synaptophysin-RFP positive vesicles were moving in inward direction as

compared to about 50-60% of inward bound vesicles in the wildtype and

heterozygous H304R/+ mice (Table 5-2).

A B

C D

115

Subsequently, 68% of synaptophysin-RFP positive vesicles were moving in

outward direction as compared to about 40-50% of outward-bound vesicles in the

wildtype and heterozygous H304R/+ mice.

Table 5-2 Synaptophysin-RFP positive vesicles motility in hippocampal neurons at DIV 14

Interestingly, synaptophysin-RFP positive vesicles in homozygous H304R/R mice

displayed a significant increase in the distance covered (µm) as compared to the

wildtype and heterozygous H304R/+ mice (One-way ANOVA, p<0.05) (Figure 5-

2). Heterozygous H304R/+ mice showed slight but insignificant increase in its

vesicular distance coverage as compared to the wildtype mice. However, the

116

synaptophysin-RFP positive vesicles in heterozygous H304R/+ mice showed a

significant decrease in its velocity (µm/sec) as compared to both wildtype and

homozygous H304R/R mice (One-way ANOVA, p<0.05) (Figure 5-2).

Figure 5-2 Synaptophysin-RFP positive vesicles motility in hippocampal neurons at DIV 14

The graphs show the distance (µm) covered by the vesicles in the inward (A) and outward (B) bound directions. The graphs show the velocity (µm/sec) of the vesicles in the inward (C) and outward (D) bound directions. Wildtype (black bars), heterozygous H304R/+ mice (gray bars), and homozygous H304R/R mice (blue bars). Note: * shows statistical significance (1-way ANOVA, p<0.05).

A B

C D

117

5.2.2 Synaptophysin-RFP positive vesicular motility in cortical neurons

Cortical neurons from H304R mice were isolated and plated on coverslips for

intracellular motility experiments. The neurons from the same genotype were

pooled together independent of the gender of the embryo. Cortical neurons were

then infected with BacMam synaptophysin-RFP and were imaged at the ages

DIV 10 and 14. Various parameters such as the number of movements in the

inward (towards the cell body) and outward (away from the cell body) directions,

run length or distance (µm), and velocity (µm/sec) were recorded of the

synaptophysin-RFP positive vesicles that were moving. The data was then

analyzed and statistically compared between the three genotypes.

At DIV 10, synaptophysin-RFP positive vesicles in both heterozygous H304R/+

and homozygous H304R/R mice showed an altered distribution of movement

patterns. About 63% of the vesicles in H304R mutant mice were moving in the

inward direction as compared to about 82% of inward direction bound vesicles in

the wildtype mice (Table 5-3). Subsequently, about 40% of vesicles were

outward direction bound in both heterozygous H304R/+ and homozygous

H304R/R mice as compared to only about 18% outward direction vesicles in the

wildtype mice.

118

Table 5-3 Synaptophysin-RFP positive vesicles motility in cortical neurons at DIV 10

Furthermore, both inward and outward bound synaptophysin-RFP positive

vesicles in heterozygous H304R/+ and homozygous H304R/R mice showed an

alteration in the distance (µm) covered as compared to the wildtype, but this

alteration was not significantly different statistically (Figure 5-3). Heterozygous

H304R/+ mice showed a slight (but insignificant) increase in the velocity (µm/sec)

of the inward and outward bound synaptophysin-RFP positive vesicles as

compared to the wildtype and homozygous H304R/R mice (Figure 5-3).

119

Figure 5-3 Synaptophysin-RFP positive vesicles motility in cortical neurons at DIV 10

The graphs show the distance (µm) covered by the vesicles in the inward (A) and outward (B) bound directions. The bottom graphs show the velocity (µm/sec) of the vesicles in the inward (C) and outward (D) bound directions. Wildtype (black bars), heterozygous H304R/+ mice (gray bars), and homozygous H304R/R mice (blue bars).

At DIV 14, synaptophysin-RFP positive vesicles showed a similar distribution of

movement patterns in all 3 genotypes. About 50-60% of the vesicles in H304R

mice was moving in the inward direction and about 40-50% of the vesicles were

outward direction bound in all three genotypes (Table 5-4).

A B

C D

120

Table 5-4 Synaptophysin-RFP positive vesicles motility in cortical neurons at DIV 14

Both inward and outward bound synaptophysin-RFP positive vesicles had similar

distance (µm) covered in all 3 genotypes as well (Figure 5-4). Heterozygous

H304R/+ mice showed a peculiar pattern in the velocity (µm/sec) characteristic of

the inward and outward bound synaptophysin-RFP positive vesicles as

compared to the wildtype and homozygous H304R/R mice (Figure 5-4). Velocity

of synaptophysin-RFP positive vesicles was decreased in the inward bound

121

direction and increased in the outward bound direction as compared to both the

wildtype and homozygous H304R/R mice, but this difference was not statistically

significant (Figure 5-4).

Figure 5-4 Synaptophysin-RFP positive vesicles motility in cortical neurons at DIV 14

The graphs show the distance (µm) covered by the vesicles in the inward (A) and outward (B) bound directions. The bottom graphs show the velocity (µm/sec) of the vesicles in the inward (C) and outward (D) bound directions. Wildtype (black bars), heterozygous H304R/+ mice (gray bars), and homozygous H304R/R mice (blue bars).

A B

C D

122

5.2.3 Rab7-GFP positive vesicular motility in cortical neurons

Cortical neurons from H304R mice were isolated and plated on coverslips for

intracellular motility experiments. The neurons from the same genotype were

pooled together independent of the gender of the embryo. Cortical neurons were

nucleofected with Rab7-GFP and were imaged at the age DIV 10. Various

parameters such as the number of movements in the inward (towards the cell

body) and outward (away from the cell body) directions, run length or distance

(µm), and velocity (µm/sec) were recorded of the synaptophysin-RFP positive

vesicles that were moving. The data was then analyzed and statistically

compared between the three genotypes.

At DIV 10, Rab7-GFP positive vesicles in both heterozygous H304R/+ and

homozygous H304R/R mice showed a similar distribution of movement patterns

in both inward and outward directions as compared to the wildtype mice (Table 5-

5). The Rab7-GFP positive vesicles in homozygous H304R/R mice showed an

increase in the distance (µm) covered by them in the inward direction, but this

difference was not significant compared to the wildtype and heterozygous

H304R/+ mice.

123

Table 5-5 Rab7-GFP positive vesicles motility in cortical neurons at DIV 10

124

Figure 5-5 Rab7-GFP positive vesicles motility in cortical neurons at DIV 10

The graphs show the distance (µm) covered by the vesicles in the inward (A) and outward (B) bound directions. The bottom graphs show the velocity (µm/sec) of the vesicles in the inward (C) and outward (D) bound directions. Wildtype (black bars), heterozygous H304R/+ mice (gray bars), and homozygous H304R/R mice (blue bars). Note: * shows statistical significance (1-way ANOVA, p<0.05).

The outward bound Rab7-GFP positive vesicles in heterozygous H304R/+

showed a significant increase in the distance (µm) covered as compared to the

wildtype and homozygous H304R/R mice (One-way ANOVA, p<0.05) (Figure 5-

5). The velocities of both inward and outward bound Rab7-GFP vesicles were

comparable between all three genotypes (Figure 5-5).

A B

C D

125

5.3 Conclusions

The preliminary motility studies with synaptophysin-RFP and Rab7-GFP positive

vesicles in the hippocampal and cortical primary neurons from the H304R mice

presented highly variable and sometimes contradictory results. There could be

multiple reasons behind these variable results, such as:

Pooling of the primary neurons from the male and female embryos

together for each genotype might have masked any motility differences,

especially in the heterozygous H304R/+ mice neuronal cultures. We

previously showed that the male heterozygous H304R/+ mice are affected

more than the female heterozygous H304R/+ mice in their motor skills

phenotype. A better approach would be to perform the motility studies on

primary neuron cultures that are grown separately based on the genotype

and gender of the embryos.

The primary neurons were collected from the E16.5 developing embryos

that are hard to isolate and grow consistently in vitro.

The motility videos were collected from the longest visible axonal

projection in culture for the data analysis. The classification of “inward”

and “outward” direction bound moving vesicles may not be accurate if the

longest visible axonal projection was not an axon. Only the axons are

known to have an established microtubule polarity and the dendrites have

microtubules arranged in a random fashion with mixed polarities. Hence,

126

this could have introduced another source of variability in the direction

based motility data comparison.

The motility analysis was done on the primary neurons that were grown in

vitro for 10 (DIV 10) or 14 (DIV 14) days. Performing motility analysis at a

range of early and late time points might reveal some alterations in the

motility phenotype.

In conclusion, performing future motility experiments with the abovementioned

considerations might help resolve the variability issue observed in the preliminary

study results. Additionally, at the time of this preliminary study we chose neurons

from the central nervous system (CNS) due to the established experimental

protocols in CNS neurons. Future motility studies should also be performed on

the motor and sensory neurons from the peripheral nervous system in the H304R

mouse model.

127

CHAPTER 6: DISCUSSION

6.1 All dynein mutations don’t act the same

The motor protein dynein is responsible for retrograde transport of crucial

cargoes from the axonal tip to the soma in neurons. Various studies show the

specific role of dynein in neuronal functions such axonal transport (He et al.,

2005), retrograde signaling (Perlson et al., 2009), protein degradation and

recycling (Ligon et al., 2005), and neuronal migration (Ori-McKenney and Vallee,

2011). It is evident that neurons need a robust retrograde mediated transport for

its growth, maintenance, and survival. Hence, it is not surprising to see that

several mutations in cytoplasmic dynein have been reported in human

neurodegenerative diseases. These dynein mutations (Table 1-2) are associated

with motor and sensory neuropathies such as Charcot-Marie-Tooth type 2O

(CMT2O) (Weedon et al., 2011), Spinal Muscular Atrophy with Lower Extremity

Dominant (SMA-LED) (Harms et al., 2012) (Tsurusaki et al., 2012), in

malformations of cortical development (MCD) (Poirier et al., 2013), in intellectual

disability (Willemsen et al., 2012), and congenital cataract and gut dysmotility

(Gelineau-Morel et al., 2016).

The spectrum of neurodegenerative diseases in humans point out the

susceptibility of neurons to the dynein mutations in the peripheral and central

nervous systems. Perhaps, it is the distinct structure and function of neurons that

make them very sensitive to the altered functions of mutant dynein. For example,

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the motor neurons have very long axons that can grow up to a meter in humans

and require robust axonal transport for its maintenance (Levy and Holzbaur,

2006). Not surprisingly, about 18 distinct mutations in dynein heavy chain are

reported thus far in SMA-LED patients with peripheral motor neuron loss (Scoto

et al., 2015). The same study also reported about 16 mutations in dynein heavy

chain associated with malformations of cortical development (MCD), signifying

the vulnerability of neurons in the central nervous system to the mutant dynein

functions.

The dynein heavy chain (DHC) mutations in motor domain and tail domain alter

the dynein function differentially, thus giving rise to a spectrum of human

neuropathies spanning both the peripheral and central nervous systems in

humans (Scoto et al., 2015). Interestingly, the same DHC mutation can cause

two different diseases as well. For examples, the H306R mutation in DHC is

associated with both CMT2O (Weedon et al., 2011) and SMA-LED (Tsurusaki et

al., 2012). In fact, both CMT2O and SMA-LED patients exhibit similar clinical

symptoms such as muscle weakness and wasting in legs, developmental

milestone delays, skeletal deformities, joint contractures, and gait problems. The

only difference is that the CMT2O patients also exhibit sensory loss symptoms, in

addition to the motor deficits.

Various mutations exist in the DHC peptide with overlapping phenotypes, but

they all do not cause just one exact same neuropathy in humans. Rather these

DHC mutations cause different type of neuropathies, thus signaling that the

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mutations alter proteins in different ways. Our previous studies with filamentous

fungi N.crassa also recapitulated similar observations where various dynein

mutations resulted in distinct fungal morphology and vesicular motility

(Sivagurunathan et al., 2012). Because different dynein mutations do not behave

the same and cause the same phenotype in humans, there is a possibility that

there might be a distinct molecular mechanism of disease behind each dynein

mutation.

6.2 Relevance of H304R Mouse Model

Relevance of heterozygous H304R/+ mice

One key feature of Charcot-Marie-Tooth type 2O (CMT2O) disease is the

heterogeneity of its onset and the severity of the clinical symptoms in patients.

Family members with the same CMT2O dynein mutation reported variable ages

of onset and the severity of the clinical symptoms expressed. Similar trends were

observed in our study with the heterozygous H304R/+ mice behavioral and

phenotypic characteristics. The heterozygous H304R/+ mice showed variable

onset of phenotype, i.e. early onset of reduced all limbs grip strength and rotorod

test defects, while late onset of reduced front grip strength and tail suspension

reflex. Some heterozygous H304R/+ mice expressed a mild phenotype, whereas

some expressed a more severe phenotype just like as seen in the human

patients with the corresponding autosomal dominant CMT2O mutation.

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A possible explanation to this heterogeneity might lie in the fact that dynein

heavy chains (DHC) dimerize with each other and interact with other subunits

(like intermediate chains, light intermediate chain, & light chains) to form a

functional dynein motor complex. In the heterozygous H304R/+ mice, there

would be a mix of DHC monomers that have been translated from the

heterozygous DHC alleles (H304R/+). From the mix of wildtype and H304R

mutant DHC monomers, three possible combinations of dynein population may

arise –

A quarter population of dynein containing wildtype DHC dimers only

A quarter population of dynein containing mutant DHC dimers only

And half of the population of dynein containing DHC dimers of 1 wildtype

monomer dimerized with 1 mutant DHC monomer

Thus, at any given time, a heterozygous H304R/+ mouse cells will have all three

types of DHC dimers combination – wildtype, homozygous, and heterozygous

DHC dimers. There could be a “dose-dependent” modulation of dynein motor

function depending on what percentage of wildtype, homozygous, and

heterozygous DHC dimers are present and interacting functionally with other

dynein subunits at a given time. The variable level of DHC dimers may also affect

the interaction of the dynein motor complex with its other binding partners and/or

cargoes. Future biochemical tests with purified dynein from H304R mouse model

can reveal key insights on the cellular dynamics that happen when different

“doses” of wildtype, homozygous, heterozygous DHC dimers are around in a cell.

131

One key experiment will be to test the binding efficiency of wildtype,

homozygous, and heterozygous DHC dimers with other dynein subunits and key

binding partners of dynein such as dynactin, BicD, Lis1, Nudel etc..

Another key observation in the heterozygous H304R/+ mice was the gender-

based differences in the exhibition of the motor skills phenotypic defects. Overall,

the male heterozygous H304R/+ mice appeared to be more affected by the

dynein mutation as evident in the various motor-skills assays. In contrast, the

female heterozygous H304R/+ mice showed mild to no defects in the exhibition

of the motor skills phenotypic defects. A potential reason behind this gender-

based observation could be the presence of the neuroprotective effects of

estrogen in the female mice (Green and Simpkins 2000). Thus, the female

heterozygous H304R/+ mice appear to be less sensitive to the effects of the

dynein mutation on their motor skills phenotype. Similar observations were made

in a small study with CMT1A patients where the male patients were shown to

have significantly reduced muscle strength in their lower limbs as compared to

the female patients (Wozniak and Gess, 2015).

Relevance of homozygous H304R/R mice

The homozygous H304R/R mice exhibited severe symptoms of atypical tail

suspension, reduced grip strength and rotorod deficits right from an early age.

The onset and the severity of homozygous H304R/R phenotype recapitulated the

severity of SMA-LED symptoms as seen in another and the only reported

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homozygous DHC mutation (R399G) (Scoto et al., 2015). Offspring with this

DHC homozygous R399G mutation showed an early onset and severe motor

symptoms as compared to the parents that were heterozygous for the same DHC

mutation.

Furthermore, both male and female homozygous H304R/R mice showed similar

trends of deficiencies in their behavioral phenotype, unlike the heterozygous

H304R/+ mice that showed gender-based deficiencies in their phenotype

(Sabblah et al., 2018). As in the previous publication, only the male heterozygous

H304R/+ mice primarily exhibited an altered motor skills phenotype in most

behavioral tests. The female heterozygous H304R/+ mice exhibited little to no

difference in its behavioral phenotype as compared to the wildtype mice. This key

difference between the heterozygous H304R/+ and homozygous H304R/R mice

indicates that the homozygous H304R/R mutation has a distinct and more severe

effect on the dynein function that may not altered by the gender differences of the

mice.

Overall relevance of H304R mice in understanding the dynein function

In conclusion, both heterozygous H304R/+ and homozygous H304R/R mice with

DHC mutations recapitulate the onset and the phenotypes as seen with the

reported heterozygous (H306R) and homozygous (R399G) dynein mutations in

humans. Hence, our H304R mouse model would serve as a relevant and useful

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mouse model to understand the molecular mechanism of mutant dynein in

human neuropathies.

6.3 Comparison of H304R Mouse Model with Other Dynein Mouse Models

Previously reported mutant DHC mice were generated using ENU or radiation

based mutagenesis method that exhibited dominant neurological phenotypes

(Hrabe de Angelis et al., 2000) (Duchen, 1974). The mutant DHC mice were

Legs at odd angle (Loa) mice with F580Y mutation, Cramping (Cra) mice with

Y1055C mutation, and Sprawling (Swl) mice with a nine base-pair deletion

[GIVT]1040[A] mutation in the cytoplasmic dynein heavy chain gene. Even

though all three mutations were located in the tail domain of the dynein heavy

chain, the Loa, Cra and Swl did not display the same exact phenotype.

A key difference is that the H304R mouse model has the corresponding dynein

mutation (H306R) that is reported to cause neuropathies in humans (CMT2O and

SMA-LED) (Tsurusaki et al., 2012; Weedon et al., 2011). In contrast, the

mutagenesis generated Loa, Cra, and Swl mice models do not have a

corresponding mutation that is associated with any human neuropathies. Hence,

it is hard to correlate their findings as a basis for human neuropathy. Another key

difference between H304R mice and other DHC mice is that homozygous

Loa/Loa, Cra/Cra, and Swl/Swl were embryonically lethal or did not survive 24

hours post birth due to inability to move and feed (Hafezparast et al., 2003)

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(Zhao et al., 2016). In contrast, the homozygous H304R/R mice survived post

birth and had a normal lifespan.

The H304R mouse model shares many commonalities with other published

models of dynein heavy chain mutation Loa, Cra, and Swl. The first commonality

is that all the mutations are located in the tail domain of the dynein heavy chain.

Shared phenotypes between H304R model and Loa/+, Cra/+ and Swl/+ mice are

– atypical reflex of clenching hind limbs closer to the body upon tail suspension,

normal life span, gait abnormalities, reduced grip strength in hind limbs, and

thinner sciatic nerves. Additionally, H304R/+ and H304R/R mice showed deficits

in rotarod performance similar to Loa/+ mice.

In conclusion, the H304R model and previously published DHC mice model

display a varying range of phenotypes. The severity of the mouse DHC mutations

on the dynein function varies in the following order – the homozygous Loa/Loa,

Cra/Cra and Swl/Swl are the most severe mutations, followed by the

homozygous H304R/R and heterozygous Loa/+, Cra/+ and Swl/+ moderate

mutations, and finally the heterozygous H304R/+ mild mutation (Figure 6-1).

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Figure 6-1 Severity of phenotypes associated with DHC mutations

Comparison of the severity of mouse DHC mutations with the human DHC mutations. The severity of the

phenotype associated with the DHC mutation increases from left to the right.

MOUSE

MUTATIONS

HUMAN MUTATIONS

Loa/+

Cra/+

Swl/+ H304R/+ H304R/R

H306R/+

(CMT2O)

Loa/Loa

Cra/Cra

Swl/Swl

LETHAL

R399G/G

No Mutation

NO SYMPTOM

No Mutation

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6.4 Future Sensory Characterization Work of H304R Mouse Model

Collectively behavioral assessments like tail suspension, grip strength, and

rotarod performance tests have given us some robust insights into the motor

phenotype of H304R mouse model. Neuromuscular junction (NMJ) analysis gave

additional evidence supporting the defective motor phenotype of H304R mouse

model (Sabblah et al., 2018). NMJ analysis revealed the deficiencies in the

morphology and complexity in heterozygous H304R/+ and homozygous

H304R/R male mice. The synaptophysin occupancy in NMJs were also found to

be faulty, indicating a loss of motor neuron functionality in heterozygous H304R/+

and homozygous H304R/R male mice. The reason why the motor neurons are

not functional in mutant H304R mice is yet to be explored.

However, investigating the motor characterization is only one part of the equation

in understanding a motor-sensory neuropathy like CMT2O. Multiple studies have

indicated dynein’s role in sensory neuron maintenance and survival (Deinhardt et

al. (2006); (Saxena et al., 2005; Yano et al., 2001). In parallel, the previously

published dynein heavy chain mutant Loa/+, Cra/+ and Swl/+ mice have also

reported sensory deficits. Heterozygote Loa/+ and Swl/+ mice reportedly had

reduced size of dorsal root and fewer neurons in their lumbar dorsal root ganglia

(Chen et al., 2007). Furthermore, Loa/+ and Swl/+ also showed abnormal

proprioceptive phenotype as indicated by the absence of H-reflex and specific

loss of proprioceptive sensory neurons as compared to the wildtype mice.

137

Interestingly, Loa/+ and Swl/+ mice also had thinner sciatic nerves as compared

to the wildtype control mice. The Swl/+ mice showed no difference in the latency

to tail withdrawal as compared to the wildtype control mice. The Cra/+ mice

displayed subtler sensory phenotype than the Loa/+ and Swl/+ mice and had

reduced size of its dorsal root due to the loss of large caliber axon (potentially

proprioceptive) neurons (Dupuis et al. (2009). In conclusion, CMT2O patients

and all the other dynein mutant mice (Loa/+, Cra/+, and Swl/+) showed mild to

severe sensory phenotype and subsequently raised a question on the sensory

phenotype of the H304R mouse model. Hence, I started investigating the

sensory phenotype of H304R mice with tail flick test, sciatic nerve thickness

measurements, and optimizing the protocol to identify the dorsal root ganglia

neuronal subpopulation.

Preliminary results from the tail flick tests showed a quicker latency to tail

withdrawal from the pain stimulus in the homozygous H304R/R mice. One

possible explanation is that the homozygous H304R/R mice have hypersensitivity

to thermal pain stimulus due to reduced pain threshold as commonly seen in

mice models with sciatic nerve injury and persistent sensory neuropathic pain

(Wang et al., 2009; Yajima et al., 2002). Incidentally, severely affected CMT2O

patients also reported having neuropathic pain (Weedon et al., 2011).

Next, a closer examination of peripheral nervous system revealed thinner sciatic

nerves and reduced axonal counts in both the heterozygous H304R/+ and

homozygous H304R/R mice. It would be interesting to learn if the sciatic nerve

138

thinning in homozygous H304R/R is caused by the loss of motor and/or sensory

neurons. One way to test this would be to stain the sciatic nerve cross sections

with unique markers for both motor and sensory neurons. Then the number of

motor and sensory axons passing through the sciatic nerve can be counted and

normalized against the total number of neurons (labeled with a pan-neuronal

marker). This would help in identifying if any alteration in the number of motor

and/or sensory neurons led to thinner sciatic nerve in homozygous H304R/R

mutant mice.

To advance the sensory characterization, it would be of critical importance to

identify the neuronal subpopulation inside the dorsal root ganglia (DRG) as well

of the H304R mouse model. Previous work on Swl/+ mice revealed a significant

loss of proprioceptive neurons in the lumbar DRG (Chen et al., 2007). My

optimization work established the working protocol of how to stain and identify

nociceptive and proprioceptive subpopulations in the same section of the DRG of

H304R mice. The next step would be to stain DRG sections from the wildtype,

heterozygous H304R/+, and homozygous H304R/R mice with the unique

markers for each neuronal subpopulation. Then the number of neuronal cells

positive for nociceptive marker or proprioceptive marker can be counted and

normalized over the total count of neurons in the DRG. The results would yield if

there are any differences in the neuronal subpopulations count in the H304R

mouse model, and potentially reveal the neuropathology of mutant dynein in

sensory dysfunction.

139

6.5 Potential Dynein Disruptions in the H304R Mice and Future Studies

Our preliminary studies revealed both motor and sensory defects in our H304R

mouse model. Because large sized motor and sensory neurons have high

demands for energy and require robust axonal transport, it is not surprising that

motor and sensory neurons are vulnerable to the mutations that may alter the

dynein functions in retrograde axonal transports (LaMonte et al., 2002; Ori-

McKenney et al., 2010). Dynein has an established role in transporting nerve

growth factors via activated trk signaling from the neuronal tip to the soma for

neuronal survival (Heerssen et al., 2004; Mitchell et al., 2012; Yano et al., 2001).

Impaired retrograde signaling of growth factors or neurotropic factors in sensory

neurons has been shown to develop peripheral neuropathy phenotypes in mice

(Rishal et al., 2012). Based on this literature evidence, I speculate three different

potential mechanism of the H304R mutation in the disease progression.

The first disease mechanism is that the dynein mutation affects the retrograde

signaling of endosomes with the neurotrophic factors. This would lead to a loss of

sensory neurons in the H304R mice. As evident from my preliminary work with

the sensory phenotype characterization, I think the nociceptive (pain sensing)

subpopulation of the sensory neurons might be affected more as seen in the

quick tail withdrawal latency in the mutant H304R mice. Hence, impaired

retrograde signaling leading to the loss of nociceptive sensory neurons ultimately

causes the altered sensory phenotypes such as altered tail withdrawal, thinner

sciatic nerves, and a loss of axonal counts in the heterozygous H304R/+ and

140

homozygous H304R/R mice. Investigating the robustness of retrograde signaling

of trk receptors upon activation of growth factors such as brain derived

neurotrophic factors (BDNF), nerve growth factor (NGF), or neurotrophins (NTs)

would be of critical importance in understanding the role of mutant dynein in the

H304R mice.

The second disease mechanism is the impaired retrograde transport of length

sensing cellular factors leading to an altered growth of sensory neurons. This is

based on an interesting study that revealed a role of dynein in maintaining the

sensory neurons length (Ori-McKenney et al., 2010). In this study, adult sensory

neurons in the heterozygote Loa/+ mice had longer neurites than the wildtype

controls when grown in vitro. Additionally, heterozygote Loa/+ mice showed a

similar trend of significantly longer axis from cell periphery to the nucleus in

embryonic fibroblasts as compared to wildtype controls. These results indicated

that dynein might have a role in transporting the length sensing cellular factors

needed for sensory neuron axonal length maintenance. Based on the evidence

presented above I speculate that the H304R mice may have an impaired

retrograde transport of length sensing cellular factors. And that this may lead to

an alteration in the sensory neuron lengths. Hence, it would be an exciting idea

to measure and compare the lengths of the sensory neurons in vitro from the

H304R mice to reveal any alterations in the sensory neuron lengths.

Lastly, the third possible mechanism for the dynein dysfunction in the H304R

mice could be a biochemical interaction defect based on the location of the

141

H304R mutation on the dynein heavy chain (DHC) peptide. The H304R mutation

is located in the tail domain of the DHC peptide. The H304R mutation lies within

the homodimerization region of the tail domain in the DHC. The presence of the

H304R mutation in the homodimerization region may cause a hindrance to the

binding of two DHC monomers to form a functional DHC dimer. The H304R

mutation could also contribute to interference in the interactions of the DHC to

other dynein subunits like dynein intermediate chains (DICs) and dynein light

chains (DLCs). The interactions with DICs and DLCs allow for the indirect

interaction of the DHCs to its binding factors such as dynactin, Lis1, BicD and

these binding factors have been implicated in the proper functioning of the

neurons such as neurofilament transport, clearance of misfolded proteins, and

neuronal migration. Thus, it would be insightful to perform future studies to

investigate the functional binding of the mutant DHC with other dynein subunits

and binding factors using biochemical assays. These biochemical assays may

potentially uncover any altered dynein interaction that may contribute to the

pathogenesis of the disease as discussed above.

142

APPENDIX A: DISSERTATION DEFENSE ANNOUCEMENT

143

Announcing the Final Examination of Ms. Swaran Nandini for the degree of Doctor

of Philosophy in Biomedical Sciences

Dissertation Title: “MOTOR AND SENSORY CHARACTERIZATION OF A

MOUSE MODEL OF CHARCOT-MARIE-TOOTH TYPE 2O DISEASE”

Date: 04/03/2019 Time: 3:00pm

Location: Lake Nona BBS 103 (Live)/ BMS 136 (Simulcast)

ABSTRACT

Dynein is an essential motor protein required for the maintenance and survival of the

cells. Dynein forms a motor complex to carry intracellular cargoes like organelles,

growth factors, peptides, hormones etc. along the microtubules inside the cells. In

neurons, the dynein is the retrograde motor protein that moves cargoes from the neuronal

tip to the neuronal soma along the length of an axon. Dynein has an established role in

the neuronal nuclei migration, transport of neuronal survival signals and growth factors,

organelle positioning inside neurons etc. Hence, it is not very surprising that numerous

mutations in dynein have been reported in association with neurodegenerative diseases in

humans. The first mutation (H306R) in dynein heavy chain was reported to cause

Charcot-Marie-Tooth Type 2O disease (CMT2O) in humans. CMT2O patients display

motor-sensory neuropathy symptoms such as muscle weaknesses and wasting in legs,

skeletal deformities like pes cavus (high foot arching), difficulty in walking, and a loss of

sensation.

We developed a novel knock-in H304R mouse model with the corresponding CMT2O

linked dynein mutation to understand the disease’s molecular mechanism. We

investigated and characterized the motor-sensory phenotype of the H304R mouse model

(wildtype, heterozygous (H304R/+) and homozygous (H304R/R) mice). First, we started

with testing mice on motor skills behavior tests such as tail suspension reflex, grip

strength test, and rotarod test at 3, 6, 9 and 12 months of age. Both male and female

groups of heterozygous (H304R/+) mice displayed mild defects in tail suspension reflex,

grip strength, and rotarod performance at a later age time point. In contrast, homozygous

(H304R/R) mice exhibited severe defects in the tail suspension reflex, grip strength, and

rotarod performance right from an early age. Next, I analyzed the sensory phenotype of

the H304R mouse model. Homozygous H304R/R mice appeared to have thinner sciatic

nerve, reduced total fascicular area of the sciatic nerve, and significantly quicker latency

to tail withdrawal from a pain stimulus.

Collectively, our motor and sensory characterization studies reveal that H304R dynein

mouse model recapitulates the phenotype associated with CMT symptoms. Hence, the

H304R model is a useful tool in understanding the dynein function in the onset and

progression of CMT2O in humans.

144

Committee members:

Stephen J. King, Ph.D. (Committee Chair)

Annette Khaled, Ph.D.

Yoon-Seong Kim, M.D., Ph.D.

Stephen Lambert, Ph.D.

Publications:

Nandini, S.*, Conley Calderon, J.*, Sabblah, T. T., Love, R., King, L. E., King,

S. J. (2019). Mice with an autosomal dominant Charcot-Marie-Tooth type 2O

disease mutation in both dynein alleles display severe moto-sensory phenotypes.

(Submitted to Scientific Reports, * equal contribution)

Sabblah, T. T.*, Nandini, S.*, Ledray, A. P., Pasos, J., Conley Calderon, J., Love,

R., King, L. E., King, S. J. (2018). A novel mouse model carrying a human

cytoplasmic dynein mutation shows motor behavior deficits consistent with

Charcot-Marie-Tooth type 2O disease. Scientific reports, 8(1), 1739. (* equal

contribution)

Höing S, Yeh TY, Baumann M, Martinez NE, Habenberger P, Kremer L, Drexler

HCA, Küchler P, Reinhardt P, Choidas A, Zischinsky ML, Zischinsky G,

Nandini S., Ledray AP, Ketcham SA, Reinhardt L, Abo-Rady M, Glatza M, King

SJ, Nussbaumer P, Ziegler S, Klebl B, Schroer TA, Schöler HR, Waldmann H,

Sterneckert J. (2018) Dynarrestin, a Novel Inhibitor of Cytoplasmic Dynein. Cell

Chem Biol.,25(4), 357-369.

Sivagurunathan, S., Schnittker, R. R., Nandini, S., Plamann, M. D., & King, S. J.

(2012). A mouse neurodegenerative dynein heavy chain mutation alters dynein

motility and localization in Neurospora crassa. Cytoskeleton (Hoboken, N.J.),

69(9), 613-24.

Sivagurunathan, S., Schnittker, R. R., Razafsky, D. S., Nandini, S., Plamann, M.

D., & King, S. J. (2012). Analyses of dynein heavy chain mutations reveal

complex interactions between dynein motor domains and cellular dynein

functions. Genetics, 191(4), 1157-79.

Approved for distribution by Dr. Stephen J. King, Committee Chair

The public is welcome to attend.

145

APPENDIX B: RESUME

146

SWARAN NANDINI

EDUCATION & RESEARCH EXPERIENCES

Doctor of Philosophy - Biomedical Sciences, College of Medicine, University of Central Florida, FL o Received the UCF Graduate Dean's Dissertation Completion Fellowship o 7 research awards/scholarships, published 4 research papers & 5th paper

in submission, presented 18 research posters & 4 research talks

2013 - 2019

Master of Science - Biotechnology, College of Medicine, University of Central Florida (UCF), FL o Received the UCF COM Outstanding Master’s Thesis Award

2011 - 2013

Bachelor of Technology - Biotechnology, Jaypee Institute of Information Technology, Delhi, India o Completed Thesis research project and 2 summer research internships

2007 - 2011

LEADERSHIP EXPERIENCES (TOP 5 OUT OF 8)

American Society for Cell Biology (ASCB) – Ambassador & Regional Symposium Organizer o Involved in Annual conference promotion, moderated sessions, onsite

conference assistance o Partnered with the host school to organize the 2nd Annual FL Cell Biology

Symposium for 130+ attendees, recruited poster presenters & panel speakers, hosted career development workshop

2014 - 2019

University of Central Florida Recreation and Wellness Center (RWC) – Health & Fitness Intern o Achieved 100% client retention rate as a Group exercise instructor o Audited facility risks & developed creative solutions to mitigate risks (risk

management efforts) o Executed strategic projects to grow the diverse representation in the

RWC staff employment, and presented on LGBTQ+ clients customer service training (diversity & inclusion efforts)

2016 - 2019

University of Central Florida Technology Fee Committee – Graduate Student Representative o Reviewed 233 tech proposals to allocate committee funds of $22 million

2016 - 2019

National Association of Graduate Professional Students (NAGPS) – Vice Chair & Conference Chair o Organized the 2016 NAGPS Southeastern (SE) Regional Conference that

was hosted for the 1st time in FL, achieved the highest SE regional conference attendance in 4 years with participation from 7 out-of-state guest schools

2015 - 2017

147

Biomedical Sciences Graduate Student Association (BSGSA) at UCF – Student President o Achieved the highest student member enrollment ever o Recruited 6 guest speakers to talk on varied topics (bioentrepreneurship,

academia, industry) o Successfully led a cross functional team to organize the first ever 2-day

Career Symposium with limited budget, sold out registration, 1 attendee received an employment offer out of it

2014 - 2015

AWARDS

Received 16 awards and scholarships totaling $15,400 in amount. Key

highlights here-

2012 - 2019

o Graduate Dean's Dissertation Completion Fellowship University of Central Florida (UCF)

2019

o Risk Management Champion of the year UCF Recreation and Wellness Center

2018

o Diversity & Inclusion Champion of the year UCF Recreation and Wellness Center

2017

o Region II Top Educational Session Award National Intramural Recreational Sports Association

2016

o Volunteer UCF Service Award University of Central Florida (UCF)

2015

o Graduate Student Travel Award American Society for Cell Biology Education Committee

2014

COMMUNITY SERVICE

Biology Integrated Orlando Training and Enrichment Camp (BIOTEC) – Organizer & instructor Collaborated with multi-specialty teams to host 4 science outreach camps for high school students

2015 - 2018

Volunteer UCF – Event organizer & marketing specialist Led 2 different taskforce to organize two 24-hr long community service events for 200+ attendees

2015 - 2017

YMCA of Central Florida Teen Achievers Program – STEM Careers Outreach at a local high school

2015 - 2016

148

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