TUBA1A MICROTUBULES ESTABLISH THE FOUNDATIONS FOR NEURONAL

FUNCTION

by

GEORGIA CHRISTINE BUSCAGLIA

B.S., Syracuse University, 2014

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Neuroscience Program

2020

This thesis for the Doctor of Philosophy degree by

Georgia Christine Buscaglia

has been approved for the

Neuroscience Program

by

Abigail Person, Chair

Jeffrey Moore

Santos Franco

Lee Niswander

Matthew Kennedy

Wendy Macklin

Emily Bates, Advisor

Date: May 22, 2020

ii Buscaglia, Georgia Christine (PhD., Neuroscience Program)

Tuba1a Microtubules Establish the Foundations for Neuronal Function

Thesis directed by Associate Professor Emily Bates

ABSTRACT

Developing neurons undergo dramatic morphological changes to appropriately migrate

and extend axons to make synaptic connections. The microtubule cytoskeleton, made of a/b-

tubulin dimers, drives neurite outgrowth, promotes neuronal growth cone responses, and

facilitates intracellular transport of critical cargoes during neurodevelopment and in adulthood.

The mechanisms by which neurons regulate microtubule network assembly and organization to

respond appropriately to external cues are largely unknown. Cells express different a- and b-tubulin genes, or isotypes, providing one potential level of microtubule regulation. TUBA1A is

the most abundant a-tubulin isotype in the developing brain and mutations to TUBA1A in

humans cause severe brain malformations. The role of TUBA1A in the adult brain is largely

unknown. We use a loss-of-function mutation to Tuba1a, Tuba1aN102D, to investigate

requirements for Tuba1a in developing and adult neurons. Tuba1aND substitution reduced the

amount of Tuba1a protein and prevented incorporation of Tuba1a into cellular microtubule

polymers. Tuba1aND/+ neurons with reduced Tuba1a had altered microtubule dynamics and

slower neuron outgrowth compared to controls. Reduced Tuba1a a-tubulin impacted axon extension and impaired formation of forebrain commissures in vivo, supporting a role for Tuba1a in axon guidance. Neurons deficient in Tuba1a failed to localize microtubule associated protein-

iii 1b (Map1b) to the developing growth cone, likely impacting reception of developmental

guidance cues. Further, we show that neurons with reduced Tuba1a had diminished axonal microtubule density and impaired intracellular transport. We demonstrate that Tuba1a is a major component of adult brain a-tubulin, implicating Tuba1a in adult neuronal microtubule function.

Tuba1aND/+ heterozygous mice had approximately half the amount of brain a-tubulin as wild-

type at birth, but normal a-tubulin abundance in adult Tuba1aND/+ brains. Despite this,

Tuba1aND/+ neuromuscular junction synapses deteriorated with age, leading to adult-onset

behavioral deficits. Overall, we show that reduced Tuba1a is sufficient to support neuronal

migration, but not axon guidance. We further show that the Tuba1a-rich microtubule tracks

assembled during development are essential for mature neuron function and maintenance of

synapses over time. These data provide mechanistic insight as to how TUBA1A tunes microtubule function to support neurodevelopment, and investigate novel functions of TUBA1A

in the adult nervous system.

Approved: Emily A. Bates

iv

For my father, John (Buzzy) Buscaglia. Thank you for always encouraging me, whether you

knew it or not, you inspired me to do this.

You are deeply missed.

v ACKNOWLEDGEMENTS

I would like to acknowledge the many people who have helped me through the years,

without all of you I wouldn’t be where I am today. I would like to thank my thesis advisor, Dr.

Emily Bates, for sharing her enthusiasm for discovery, constantly being my personal cheerleader,

and supporting me through all endeavors. I would like to thank my thesis advisory committee, Dr.

Abigail Person, Dr. Jeffrey Moore, Dr. Santos Franco, Dr. Matthew Kennedy, Dr. Wendy Macklin,

and Dr. Lee Niswander for their support, feedback, and enthusiastic discussion throughout this

process. I would also like to thank Dr. Jayne Aiken for creating the Tuba1a-His6 constructs, for sharing the GFP-MACF43 plasmids, and for her expertise and insightful conversations.

Thank you to Mark Gutierrez and Dr. Santos Franco (University of Colorado Anschutz)

for sharing the Myr-TdTomato plasmid. Thank you to Dr. John Caldwell (University of Colorado

Anschutz) for his assistance with rodent hindlimb muscle dissections and his kind gift of the rhodamine-labeled a-Bungarotoxin, used for NMJ-labeling. Thank you to Kyle Northington

(University of Colorado Anschutz) for his aid with the qRT-PCR experiments in Chapters II and

III. Thank you to Katie Hoff (University of Colorado Anschutz) for her assistance with the Tuba1a-

His6 rat neuron experiments in Chapter II. Thank you to Danae Mitchell (University of Colorado

Anschutz) for sectioning and staining spinal cord sections used in Chapter III. Thank you to Emily

Sullivan Gibson, Alexandra Lucas, Dr. Matthew Kennedy and Dr. Mark Dell’Acqua (University of Colorado Anschutz) for providing rat P0-P2 cortex and use of the AMAXA nucleofector. Thank you to Dr. Jennifer Bourne and the Electron Microscopy Core (University of Colorado Anschutz).

Finally, I would like to thank my boyfriend, family and friends for their unending encouragement of my scientific pursuits. Thank you for supporting me through a crazy five years, and for making sure I had some fun along the way.

vi TABLE OF CONTENTS CHAPTER

I. MICROTUBULES IN DEVELOPMENT AND DISEASE OF THE NERVOUS

SYSTEM ...... 1

Introduction ...... 1

Microtubule Properties and Regulation ...... 5

Microtubule Structure and Dynamic Instability ...... 5

Microtubule Polymerization ...... 6

Mechanisms of Microtubule Regulation: MAPs and the Tubulin Code ...... 8

Microtubule-Associated Proteins ...... 8

Tubulin Post-Translational Modifications ...... 9

Tubulin Isotypes and the “Tubulin Code” ...... 13

TUBA1A: The Neuronal a-tubulin ...... 16

TUBA1A Identity ...... 16

Tuba1a Expression Pattern ...... 19

Mechanisms Regulating TUBA1A Expression ...... 22

Tuba1a in Regeneration ...... 24

Microtubules in Developing and Adult Neurons ...... 26

Microtubules in Neural Development ...... 26

Adult Neuronal Microtubule Function ...... 28

Microtubules in Nervous System Disease ...... 30

vii Tubulinopathies Reveal Essential Role of TUBA1A in Brain Formation

and Function ...... 30

Mouse Models Provide Insight into neuronal microtubule function ...... 33

Microtubules in Neurodegeneration ...... 36

Conclusions ...... 40

II. TUBA1A PLAYS AN ESSENTIAL ROLE IN AXON PATHFINDING ...... 41

Introduction ...... 41

Materials and Methods ...... 43

Results ...... 50

TUBA1A is required for formation of midline commissural structures ... 50

Tuba1aND a-tubulin does not incorporate into neuronal microtubules .... 53

Reduced Tuba1a alters microtubule dynamic properties in neurons ...... 58

Tuba1a dictates neurite extension and cytoskeletal organization in growth

cone ...... 60

Tuba1aND neurons fail to localize critical developmental proteins to

growth cone ...... 65

Multi-faceted functions of TUBA1A during neurodevelopment ...... 67

Discussion ...... 67

Studying a-tubulin isotypes in vivo ...... 67

TUBA1A influences microtubule density, dynamics, and function in cells

...... 70

Models of Tuba1a-dysfunction reveal potential roles in developmental

signaling ...... 72 viii III. REDUCED TUBA1A TUBULIN CAUSES DEFECTS IN TRAFFICKING AND

IMPAIRED ADULT MOTOR BEHAVIOR ...... 75

Introduction ...... 75

Materials and Methods ...... 77

Results ...... 85

Tuba1a is a major component of adult a-tubulin mRNA ...... 85

Tuba1aND is a loss-of-function mutation in mice ...... 86

Reduced Tuba1a function results in fewer microtubule tracks in neurites 90

Tuba1a is required for functional intracellular transport ...... 90

Diminished Tuba1a causes late-onset behavioral deficits ...... 95

Tuba1a deficit impairs hindlimb behavior without causing cell death or

axon degeneration ...... 98

Neuromuscular junction synapses are diminished by prolonged

microtubule dysfunction ...... 101

Discussion ...... 104

Tuba1a is critical for developing and mature neurons ...... 104

Tuba1a is Required for Intracellular Transport ...... 104

Neuronal Tuba1a Establishes Microtubule Network Function ...... 106

Tuba1a Contributes to Function of Mature Neurons ...... 107

Microtubules in Neurodegeneration: Illuminating New Roles for TUBA1A

...... 108

Supplemental Figures ...... 110

IV. CONCLUSIONS ...... 112 ix Introduction ...... 112

TUBA1A in the Developing and Mature Brain ...... 113

TUBA1A is a major component of developing and adult neuronal

microtubules ...... 113

TUBA1A Lays the Groundwork for life-long neuronal microtubule

function ...... 114

Potential Roles for TUBA1A in Mature Neuronal Microtubules ...... 118

Haploinsufficiency vs. Dominant Negative Mechanisms of Disease ...... 121

Other Tubulin Isotypes Provide Insight into Mechanism of Disease ..... 121

Tuba1aND Mouse Model Illustrates Consequences of TUBA1A LOF ...... 125

Tuba1aND is a true LOF allele ...... 125

Developmental TUBA1A Haploinsufficiency does not replicate

tubulinopathy ...... 127

Concluding Remarks ...... 128

V. REFERENCES ...... 130

x CHAPTER I

MICROTUBULES IN DEVELOPMENT AND DISEASE OF THE NERVOUS SYSTEM1

Introduction

Neurons are highly specialized cells with elaborate morphologies that place unique

demands on their cytoskeletons. Microtubules are dynamic cytoskeletal polymers that help to

dictate neuronal morphology and enable several specialized neuronal functions. During brain

development, neurons utilize dynamic microtubule networks to proliferate, differentiate, migrate,

and extend axons and dendrites to form synapses [1, 2]. All of these processes must be spatially

and temporally coordinated and require dynamic interaction between individual cells and the larger

cellular environment to build a brain. Following brain development, neuronal microtubules

provide structural stability to axons and dendrites and form elaborate highways upon which

intracellular cargoes can be transported. Throughout their lifetime, neurons must regulate the

number, location, and dynamic properties of microtubules to promote appropriate neuronal

responses. Microtubules additionally play an important role in synaptic plasticity both by

facilitating morphological changes to dendritic spines in response to synaptic activity and by

promoting rapid delivery of proteins, mRNAs and organelles to the synapse [3-6]. This dissertation

investigates the mechanisms by which a neuronally-enriched a-tubulin protein, Tuba1a, tunes

microtubule function to support neuron growth, axon guidance, and lays the foundation for mature

neuronal microtubule function. I present evidence that Tuba1a is a major component of neuronal

a-tubulin in developing and adult neurons, I show that Tuba1a sets neuronal microtubule density

1 Portions of this chapter are published with permission from our previously published articles: Aiken, J., et al., The alpha-Tubulin gene TUBA1A in Brain Development: A Key Ingredient in the Neuronal Isotype Blend. J Dev Biol, 2017. 5(3). [15]; and Aiken, J., et al., Tubulin mutations in brain development disorders: Why haploinsufficiency does not explain TUBA1A tubulinopathies. Cytoskeleton (Hoboken), 2019 [210].

1 and establishes microtubule dynamic properties, and demonstrate that Tuba1a is needed to support intracellular trafficking. At a cellular level, I show that Tuba1a supports neuron growth and axon pathfinding during development, and contributes to synaptic structure in mature neurons. Finally,

I present evidence that Tuba1a-induced microtubule dysfunction can be sufficient to cause adult- onset behavioral deficits, reminiscent of neurodegenerative disease. Collectively, the results presented in this dissertation expand the known roles of TUBA1A in the nervous system, and elucidate potential mechanisms of TUBA1A in disease.

How is the microtubule cytoskeleton regulated to perform diverse functions in developing and adult neurons? One potential mechanism of regulation occurs at the level of the specific tubulin protein building blocks that a cell expresses. Tubulin heterodimers, composed of α- and β-tubulin proteins, polymerize to form microtubules. In most eukaryotes, α and β tubulin proteins are encoded by a number of different genes, giving rise to unique tubulin proteins, termed isotypes.

As many as nine α- and ten β-tubulin isotypes have been identified by genome analysis in humans

[7, 8]. Incorporation of different tubulin isotypes can impact microtubule dynamic instability and modulate interactions with microtubule-associated proteins (MAPs)[9-13]. Thus, the specific α and β tubulin isotypes a cell expresses could potentially be used to modulate microtubule network properties. TUBA1A encodes an α-tubulin isotype that is highly expressed in the developing brain and has a demonstrated role in neurodevelopment due to its association with human brain malformations [14-16]. Heterozygous missense mutations that disrupt tubulin genes in humans cause a spectrum of severe neurodevelopmental phenotypes called tubulinopathies [17-22].

Tubulinopathy patients exhibit a wide range of brain malformations including cerebellar hypoplasia, commissural deficits, and cortical malformations such as lissencephaly, pachygyria, and polymicrogyria [17-22]. TUBA1A is the most commonly mutated gene in human

2 tubulinopathy, implicating this isotype in brain development. However, clinical TUBA1A variants

cause a broad range of brain phenotypes, making it difficult to pinpoint precisely how TUBA1A

contributes to neurodevelopment. Tubulinopathy-associated mutations to TUBA1A may impair specific interactions between microtubules and associated proteins, indicating that numerous molecular mechanisms may contribute to the diversity of clinical phenotypes [23-26]. Due to the complexity of tubulinopathy etiology, much remains unknown about the precise cellular and molecular mechanisms by which tubulinopathies occur. For example, it is unclear if the majority of TUBA1A variants dominantly disrupt microtubule function, or diminish TUBA1A via

haploinsufficiency. Additionally, recent work has elucidated mechanisms of cortical migration

errors in models of human tubulinopathy [25], but little is known about the origins of other aspects

of brain pathology. Agenesis and hypoplasia of the corpus callosum are highly associated with

tubulinopathy in humans [19, 20, 22]. However, it is largely unknown if corpus callosum

malformation occurs as a consequence of earlier cortical migration errors, or if separate

mechanisms contribute to axon guidance failures in tubulinopathy. This dissertation investigates

the requirements for TUBA1A in different aspects of neurodevelopment to address these gaps in

knowledge.

Despite ample evidence of a role for TUBA1A in neurodevelopment, very little is currently

known about the role of TUBA1A in adult neurons. Tuba1a expression decreases 20-fold between early-postnatal and adult stages in rodents [16], but certain areas of the brain, such as the cortex and hippocampus, retain high levels of Tuba1a expression in adulthood [27]. Thorough analysis of the a-tubulin mRNA composition in adult neurons is lacking, but TUBA1A is thought to be a minor component of the adult neuronal a-tubulin blend. However, Tuba1a does have a well- characterized role in adult nervous system regeneration, where Tuba1a expression is renewed in

3 response to neuronal injury [28-33]. Furthermore, microtubules have been implicated in the

pathology of a number of adult neurodegenerative diseases in humans. Specifically, altered

microtubule dynamics and disruptions in axonal transport have been noted in Alzheimer’s disease,

Parkinson’s disease, Huntington’s disease, and Amyotrophic Lateral Sclerosis among others [34-

41]. As such, microtubule dysfunction could serve as a common mechanism of pathogenesis

shared in many neurodegenerative diseases. Whether TUBA1A participates in microtubule dysfunction in neurodegeneration remains unknown. The work presented in this dissertation defines a role for TUBA1A in the adult nervous system, and explores the potential impact of

TUBA1A in adult-onset neurological disease.

Understanding what TUBA1A contributes to neuronal function will provide important insight as to how neuronal microtubules are tuned to support neurodevelopment and could illuminate a role for TUBA1A in adult neurons. A major area of investigation in microtubule biology aims to understand how tubulin isotypes play into the larger picture of microtubule network function. In this chapter, I will outline what is currently known about microtubule function in both health and disease of the nervous system. In particular, this chapter will focus on the current understanding of TUBA1A, including its expression, its dynamic properties, and its association with growth, injury and disease.

4 Microtubule Properties and Regulation

Microtubule Structure and Dynamic Instability

Microtubules are composed of tubulin proteins, which are obligate heterodimers of α- and

β-tubulin polypeptides. α- and β-tubulin genes are conserved across eukaryotic evolution and can

be traced back to related filament-

forming proteins in prokaryotes and

bacteriophage [42, 43].

Microtubules, and related filament-

forming proteins, are defined by their

characteristic property ‘dynamic

instability’, meaning that these

Figure 1.1 Tubulin Heterodimer Structure. polymers undergo phases of a- and b-tubulin proteins dimerize to form the tubulin heterodimer. Labels indicate GTP-binding sites, carboxy-terminal tails of a- and b-tubulin proteins, and structural terminology. heterodimer addition or removal,

termed polymerization and depolymerization, respectively. Dynamic instability properties of

microtubule polymers are fueled by binding and hydrolysis of GTP. Within the tubulin

heterodimer there exist two distinct sites capable of binding GTP: the “N-site” and “E-site”. The

N-site, or non-exchangeable GTP site, is found between α- and β-tubulin proteins within the

heterodimer, termed the intradimer interface, and exchanges GTP at a slow rate (Fig. 1.1) [44].

As the N-site is buried within the tubulin dimer and is constitutively bound to GTP, this site is

thought to play a largely structural role within the microtubule polymer [45]. Conversely, the E-

site, or exchangeable GTP site, binds and rapidly hydrolyzes GTP to GDP. The E-site is located

at the interdimer interface, formed between the β-tubulin protein of one heterodimer and the α- tubulin protein of the adjacent heterodimer (Fig. 1.1). Unlike the N-site, the nucleotide-binding

5 state of the E-site directly impacts the conformation of the heterodimer and affects interactions with other heterodimers within the microtubule polymer.

Microtubule Polymerization

The currently accepted model of tubulin polymerization states that the α-tubulin protein of an incoming heterodimer binds to the exposed β-tubulin protein on an existing polymer, creating an E-site at the interdimer interface where GTP can bind (Fig 1.1, 1.2). E-site GTP is rapidly hydrolyzed to GDP, causing structural compaction of the newly added heterodimer, referred to as tubulin maturation [46-48]. Importantly, GDP bound at the E-site decreases the affinity of interaction between tubulin heterodimers, and drives the switch from microtubule polymerization to depolymerization, termed microtubule catastrophe [49, 50]. Mature GDP- tubulin is prone to disassembly but is locked into a polymer state by the continued addition of

GTP-tubulin heterodimers to the growing end of the microtubule, called a “GTP cap”. When

GTP hydrolysis to GDP in assembled heterodimers outpaces new GTP-bound heterodimer addition, the microtubule undergoes catastrophe and will stochastically transition from polymerization to depolymerization. Microtubule rescue describes the transition from tubulin disassembly to renewed microtubule polymerization. The precise mechanisms regulating microtubule catastrophe and rescue are still under active investigation.

6

Figure 1.2 Microtubule Dynamic Instability. Microtubules exhibit dynamic instability, or stochastic switches between phases of heterodimer addition and microtubule growth (polymerizing) and heterodimer removal and microtubule shrinkage (depolymerization). Microtubules are most dynamic at the plus-end, or the end of the microtubule with exposed b-tubulin.

Polymerization and depolymerization are restricted to microtubule ends, however each

end of the microtubule exhibits different dynamic properties. As microtubules are formed from

α/β-heterodimers, these polymers have inherent structural polarity. The end of the microtubule

with exposed α-tubulin is referred to as the “minus-end”, while the end with exposed β-tubulin is

called the “plus-end” (Fig. 1.2). Microtubule minus-ends are relatively stable and exhibit much

slower dynamics than plus-ends in vitro [51, 52]. In cells, microtubule minus-ends are stably anchored to sites of microtubule nucleation, and as such dictate the spatial organization of microtubules within the cell, but are largely prohibited from exhibiting dynamic instability [53,

54]. For this reason, microtubule plus-ends are viewed as the dynamic end of the microtubule, as nearly all polymerization and depolymerization will occur at this site. Both the inherent structural polarity of microtubules and their dynamic instability properties are used by cells to construct elaborate and malleable structural networks.

7 Mechanisms of Microtubule Regulation: MAPs and the Tubulin Code

Microtubule-Associated Proteins

In eukaryotes, cells harness microtubule dynamic instability to form highly organized networks for intracellular cargo transport and force generation. Individual microtubules can be stabilized, removed, or transported to suit the needs of the cell. The organizational control of microtubule networks, and generation of movement or force along microtubules is largely mediated through interactions with microtubule-associated proteins (MAPs). Several classes of

MAPs exist within eukaryotic cells that facilitate a variety of microtubule-based processes.

Structural MAPs, like Tau and Map2, bind to microtubules and stabilize them to prevent stochastic microtubule disassembly [55]. In cells, microtubules are highly decorated with structural MAPs, and evidence is emerging that the MAP-bound state of a microtubule can dictate its interactions with other MAPs and with the larger intracellular environment [55-60].

Conversely, enzymatic severing MAPs, like katanin and spastin, bind to and break microtubule polymers into short segments that can then be transported or removed [61, 62]. The presence of both stabilizing and severing MAPs gives the cell exquisite control over the organization and dynamic properties of its microtubule network.

Motor proteins are a class of ATPase MAPs that utilize ATP hydrolysis to drive directional movement along microtubule polymers [63]. Motor protein ATPases fall into two major classes: dynein and kinesin superfamily proteins [64, 65]. While kinesins and dyneins operate by different mechanisms, they both contain ATPase activity, microtubule-binding, and cargo-binding domains that allow them to bind intracellular cargoes and hydrolyze ATP to generate movement along microtubule tracks [63]. The specific cargoes bound by different dynein and kinesin family members are known in some cases, but are still under active

8 investigation. The concept that specific motor proteins may preferentially bind different cargoes provides cells with an additional layer of regulation over intracellular transport, depending on the subcellular localization and cell-specific expression of motor proteins.

In addition to the variety of different types of MAPs, many MAPs exhibit distinct expression and localization patterns. Variation in spatial and temporal expression of MAPs indicates that certain MAPs may play larger roles in different cell types and at different stages of life. For example, developing neurons express Map1b at much higher levels than adult neurons.

Studies into the function of Map1b have revealed that this MAP plays a critical role in many important developmental processes, such as cortical migration, neurite branching, axon elongation and guidance [57, 66-71]. Additionally, many MAPs can fulfill different roles in different contexts. For example, dynein facilitates chromosome segregation in dividing cells, but also promotes nuclear migration in post-mitotic neurons [65, 72]. The abundance and variety of

MAPs, and their ability to work cooperatively have made it challenging to determine the precise function of individual MAPs. It is evident that MAPs enact important modifications to microtubule function and are a critical component of cellular microtubule regulation.

Tubulin Post-Translational Modifications

Post-translational modifications (PTMs) can also change properties of microtubules.

Different PTMs are enriched in specific cell types and sub-cellular structures [73-77]. PTMs commonly found on neuronal microtubules include detyrosination/tyrosination, polyglutamylation, acetylation, and polyamination (Fig. 1.3). Many of these modifications to tubulin occur in the 15–27 amino acids at the very carboxy-terminus; a region known as the

Carboxy-Terminal Tail (CTT; Figs. 1.1, 1.3). CTTs decorate the outer surface of the

9 microtubule, contain an abundance of amino acids with negatively-charged side chains, and are major sites of PTMs. Detyrosination/tyrosination refers to the enzymatic removal of tyrosine from the carboxy-terminal tail (CTT) of α-tubulin (detyrosination), and subsequent re-ligation

(tyrosination). This tyrosine is genetically encoded by a subset of α-tubulin isotypes in humans, meaning that only certain tubulin isotypes can exist in a tyrosinated/detyrosinated state.

Although the enzyme that catalyzes the detyrosination reaction in vivo has not been identified, it is well-established that tyrosination is catalyzed by Tubulin Tyrosine Ligase (TTL), which exclusively acts on free heterodimers to ligate tyrosine onto a detyrosinated α-tubulin [78, 79].

Tyrosinated or detyrosinated α-tubulin can be differentially enriched on specific microtubules within a network, or specific regions of an individual microtubule, and appear to correlate with the age of the microtubule lattice [74, 76, 80]. Accordingly, neurons exhibit an enrichment of tyrosinated α-tubulin at regions containing more newly-assembled microtubules (e.g., neurites and the distal ends of axons) while detyrosinated α-tubulin is primarily enriched in regions with older and highly stable microtubules (the axon shaft) [81-83]. Neurons lacking the TTL enzyme undergo aberrant neuronal development, indicating an important role for the cycle of detyrosination/tyrosination [84]. Currently, there is little to no evidence that tyrosination status influences the intrinsic stability of a microtubule. In contrast, tyrosination status does impact the binding of various proteins to the microtubule surface. Cytoskeleton-Associated Protein Glycine- rich (CAP-Gly) domains selectively bind to EEY/F motifs, such as those found in the CTT of α- tubulins, and this binding strongly depends on the aromatic side chain of the terminal Y/F residue [85-88]. Detyrosination therefore inhibits the microtubule-binding activity of CAP-Gly- domains. CAP-Gly domains are found in several microtubule-associated proteins, including

CLIP170, the tubulin binding co-factors TBCB and TBCE, and the p150glued subunit of dynactin.

10 Recent studies show that the binding of p150glued and CLIP170 to tyrosinated tubulin promotes the initiation of retrograde dynein-dependent transport in vitro and at the distal ends of axons

[89, 90]. Thus, detyrosination/tyrosination provides a system for local control of microtubule-

MAP interactions and transport within a microtubule network.

Figure 1.3 Sites of post-translational modifications to tubulin. a- and b-tubulin proteins can be highly decorated with post-translational modifications (PTMs). PTMs such as tyrosination/detyrosination (blue) and polyglutamylation (magenta) are added to residues in the C-terminal tail region of tubulin proteins, which extend outwards from the surface of microtubules. Acetylation (orange) primarily occurs on the luminal surface of a-tubulin proteins that are incorporated into microtubule polymer, but additional non-canonical acetylation sites may exist. This figure was adapted from [119].

Polyglutamylation is the most abundant tubulin PTM within the brain and involves the addition of variable-length chains of glutamate residues to genetically-encoded glutamates in the

CTT regions of α- or β-tubulin. Polyglutamylation was originally found to extend from α-tubulin peptide sequences that are only present in the TUBA1A and TUBA1B isotypes, with the glutamate chains extending from the γ-carboxyl group of the genetically-encoded glutamate at position 445 [91]. This may represent the primary polyglutamylation site on α-tubulin; however, alternative glutamates in the α-tubulin CTT may also be targeted for modification. The enzymes

11 that catalyze polyglutamylation belong to the Tubulin Tyrosine Ligase Like (TTLL) family [92].

To date, twelve mouse TTLLs have been identified based on sequence homology, although only

six of these exhibit polyglutamylase activity in vitro [93]. Among these, TTLLs 5, 6, 11, and 13 selectively modify α-tubulin [92-94]. The removal of glutamate residues is catalyzed by

Cytosolic Carboxypeptidase (CCP) enzymes, which can either shorten polyglutamate chains or remove the genetically encoded, penultimate glutamate from detyrosinated α-tubulin, creating a truncated species known as ∆2-tubulin [95]. Similar to detyrosination/tyrosination, polyglutamylation is thought to alter the interactions of MAPs and motors at the microtubule surface. The clearest example is the regulation of microtubule severing by spastin. Here, the length of glutamate chains provides a tunable signal for directing spastin’s severing activity [96,

97]. Although the mechanistic role of polyglutamylation in neurons is still unclear, it appears to be developmentally regulated and correlate with degeneration. Both in vivo and in vitro

experiments show that levels of polyglutamylation vary in different brain regions and increase

over development, reaching their highest level in mature neurons [98]. Loss of function

mutations affecting the CCP1 enzyme disrupt the normal program of tubulin PTMs in mouse

neurons, increasing the amount of polyglutamylated tubulin and decreasing the ∆2-tubulin

species [95]. CCP1 disruption causes Purkinje cell degeneration in mice, in a manner that

depends on increased polyglutamylation [99-101]. This suggests an important yet poorly

understand role for regulated polyglutamylation in neurons.

The presence or absence of specific PTMs can directly affect the stability of the

microtubule. For example, cold temperatures (~4ºC) will generally induce microtubule

depolymerization, but neurons contain a pool of “cold-stable” microtubules that are resistant to

temperature-induced depolymerization. The adult brain contains a much higher cold-stable

12 population of microtubules than the developing brain, and this increase in the proportion of cold-

stable microtubules has been attributed to an accumulation of polyamination on neuronal

microtubules with age [102, 103]. Polyamination describes the addition of polyamine to

multiple, genetically-encoded glutamine residues in α- and β-tubulin, by the transglutaminase

enzyme TG2 [103]. Polyamination is sufficient to stabilize microtubules in vitro, and TG2 protein and activity levels increase postnatally, suggesting a role in neuronal maturation [103].

Acetylation of α-tubulin also correlates with microtubule stability, and staining with antibodies to acetylated tubulin is commonly used as a measure of microtubule stability in cells. However, while long-lived microtubules tend to be acetylated, the direct impact of acetylation on microtubule stability is not well established. Two recent studies indicate that acetylation of α- tubulin may promote microtubule stability through an unexpected mechanism—softening the microtubule lattice and allowing it to withstand bending without breaking and depolymerizing

[104, 105]. In addition to direct effects on the microtubule lattice, acetylation has been reported to promote the activities of kinesin-1 motors in vivo [106] and dynein motors in vitro [107]. This is intriguing, since the canonical acetylation site at lysine residue 40 of α-tubulin is located on the luminal side of the microtubule cylinder. How acetylation impacts lattice stability, interactions on the microtubule surface, and the larger implications of microtubule acetylation in vivo remain active areas of research.

Tubulin Isotypes and the “Tubulin Code”

A final level of microtubule network regulation comes from the specific tubulin genes that a cell expresses. Nearly all eukaryotes express multiple, distinct genes for α- and β-tubulin, known as tubulin isotypes. Recent analysis of the human genome identified nine α-tubulin isotypes and ten β-tubulin isotypes, along with dozens of pseudogenes [7, 8]. In addition to their

13 different chromosomal locations, isotypes can be distinguished by the amino acid sequences they encode, and their expression levels in different tissues or developmental stages. Some tubulin isotypes are ubiquitously expressed, but others have divergent, tissue-specific expression patterns, meaning that certain isotypes likely never interact in vivo. Despite their different expression profiles, tubulin isotypes readily co-polymerize to form microtubules with heterogeneous mixtures of isotypes in vitro. a- or b-tubulin isotypes are very similar at the amino acid sequence level, with the exception of the diverse CTT regions, and have been shown to promiscuously heterodimerize and polymerize into microtubules to form a variety of cellular structures [108-111]. In fact, even isotypes with the most divergent structure and restricted expression, such as the highly divergent β-tubulin isotype TUBB1 normally restricted to certain hematopoietic cells, also incorporate into microtubules in their normal cellular context as well as when ectopically expressed in HeLa cells [109]. Similarly, α-tubulin isotypes Tuba3a/b, Tuba1c, and Tuba4a all co-assemble in both interphase and spindle microtubules in HeLa and NIH 3T3 cells, despite their endogenous expression in non-overlapping tissues [111]. This suggests that cellular microtubules consist of blends of the available tubulin isotypes, rather than the segregation of isotypes into distinct microtubules.

While tubulin isotypes are capable of co-polymerizing, there is evidence that the different

α- and b-isotypes may not be completely interchangeable in terms of microtubule function.

Microtubules formed from purified tubulin isotypes under in vitro reconstituted conditions reveal differential dynamics properties between β-tubulin isotypes [11, 13, 112, 113]. Studies employing the microtubule stabilizing drug taxol provide additional evidence of differential dynamics properties of β-tubulin isotypes in vitro [114]. The Drosophila β-tubulin isotype, β2- tubulin, is specifically required for axoneme function, and cannot be functionally rescued by the

14 substitution of β3-tubulin [115]. Additionally, there is evidence of preferential use of certain β-

tubulin isotypes in distinct compartments of the neuron [116, 117]. There is less evidence of

functionally distinct α-tubulin isotypes. Recently, however, it was shown that Tuba8, an α-

tubulin that is typically not expressed in the brain, cannot compensate for loss of Tuba1a during

brain development [26]. These studies suggest that while tubulin isotypes may co-polymerize to

form microtubules, the microtubules that are formed from distinct blends of tubulin isotypes may

exhibit different dynamics and provide different functions. However, similarities between other

isotypes may allow for compensation if one isotype is not expressed adequately.

Given the availability of different a- and b-tubulin isotypes, microtubule networks may be uniquely poised to compensate for reduction of a particular tubulin isotype. In other words, if one isotype gene acquires a loss of function mutation and no longer provides the appropriate level of tubulin protein, the cell may be able to compensate by increasing the expression of alternative isotypes. Importantly, this model implies that the cell must have a way to sense an insufficient tubulin pool and subsequently upregulate other isotypes until a sufficient level of tubulin is produced. There is evidence that regulation of tubulin isotypes does occur in neurons

[118]; however, the mechanisms by which a cell can detect tubulin levels and induce compensatory expression remain largely unexplored. In the current state of the field, it is unclear whether upregulation of an alternative isotype will fully compensate for a loss of another isotype, or whether changes in the tubulin isotype blend within a neuron could lead to functional consequences.

Current models propose that the molecular diversity generated by genetically-encoded differences and PTMs in the CTTs act as a “tubulin code” that regulates the activities of MAPs and motors at the microtubule surface [119, 120]. Consistent with this model, several studies

15 show that altering the amino acid sequence within the CTT causes changes in microtubule

function in vivo [85, 121-124]. The tubulin code is a compelling model of how different cell types can utilize genetic differences in tubulin isotypes and PTMs to tailor microtubule network function for specialized tasks.

TUBA1A: The Neuronal a-tubulin

TUBA1A Identity

Tubulin isotypes have divergent expression profiles, with unique blends of isotypes

expressed in different cell types. In developing mammalian neurons, Tuba1a is the primary α- tubulin isotype, representing approximately 95% of a-tubulin mRNA during brain development

[15, 16, 125, 126]. During later stages, the expression of Tuba1a declines [125]. In contrast, the

β-tubulin blend in developing neurons is not dominated by a single isotype, with five different β-

tubulin mRNAs expressed at varying levels [16, 125, 126]. While the levels of a- and b-isotype mRNAs have been confirmed by multiple studies, the protein levels of specific isotypes are not well characterized. Nevertheless, we assume that the tubulin pool in young neurons consists of heterodimers formed from mostly TUBA1A a-tubulin and a mix of b-tubulin isotypes. This

suggests that during the developmental window, neurons are building axons and dendrites using

the available TUBA1A α-tubulin isotype. Studies interrogating the longevity of neuronal

proteins have identified Tuba1a and other tubulins as long-lived proteins, with Tuba1a recorded

as having an average half-life of 7.2 days in rodent neurons [127]. It is therefore important to

consider that TUBA1A used during the initial establishment of neuronal microtubule networks

may remain in microtubules after initial phases of brain development such as axon extension and

16 synapse formation are complete, and play important roles in the maintenance of neuronal connectivity.

Identity to Human Gene Protein Identity to Mouse CTT Amino Acid Sequence Human Gene Accession Accession TUBA1A Gene Isotype TUBA1A NM_006009 NP_006000 - MAALEKDYEEVGVDSVEGEGEEEGEEY Tuba1a 451/451 TUBA1B NM_006082 NP_006073 449/451 MAALEKDYEEVGVDSVEGEGEEEGEEY Tuba1b 451/451 TUBA1C NM_032704 NP_116093 442/451 MAALEKDYEEVGADSADGEDEGEEY Tuba1c 446/449 TUBA3C NM_006001 NP_005992 440/451 LAALEKDYEEVGVDSVEAEAEEGEEY TUBA3D NM_080386 NP_525125 440/451 LAALEKDYEEVGVDSVEAEAEEGEEY TUBA3E NM_207312 NP_997195 435/451 LAALEKDCEEVGVDSVEAEAEEGEEY TUBA4A NM_006000 NP_005991 432/450 MAALEKDYEEVGIDSYEDEDEGEE Tuba4a 448/448 TUBA8 NM_018943 NP_061816 399/439 LAALEKDYEEVGTDSFEEENEGEEF Tuba8 446/449 TUBAL3 NM_024803 NP_079079 329/444 LAALERDYEEVAQSF Tubal3 369/446

Table 1.1. Isotypes of α-tubulin. Human genes identified by [7] are listed, along with accession IDs for DNA and protein. CTT sequences depict the last 15–27 genetically encoded amino acids. Underlined residue is the major site of polyglutamylation [91]. Gray denotes residues that differ from TUBA1A.

The amino acid sequence of α-tubulin is strongly conserved across eukaryotic evolution.

For example, the human α-tubulin TUBA1A exhibits 86% sequence identity to α-tubulin in the unicellular eukaryote Giardia lambia and 75% identity to the α-tubulin in the budding yeast,

Saccharomyces cerevisiae. Within the human α-tubulin isotypes, there are a small number of amino acid sequence differences (Table 1.1). Importantly, these differences are conserved in isotype homologues across species, suggesting that selective pressure may maintain isotype- specific sequence differences. The majority of these differences are found in the CTT, but human

α-tubulin isotypes also exhibit amino acid differences at other positions. For example, the human

TUBA1A and TUBA1B isotypes have identical CTTs, but have different amino acids at position

232 and 340 (Table 1.1). Position 232 is buried deep within α-tubulin, while position 340 lies on the microtubule surface, near the interdimer interface [128]. Whether these amino acid differences lead to functional differences between TUBA1A and TUBA1B has not been

17 investigated. This exemplifies a general deficit in our understanding of α-tubulin isotypes— although we have known of amino acid differences between α-tubulin isotypes for over 30 years, the functional consequences of these differences remain largely unexplored.

Another distinguishing feature of tubulin isotypes is their pattern of expression across different tissues and developmental stages. The β-tubulin isotypes have been extensively mapped to different tissues, cell types, and, in some cases, sub-cellular localization [116, 129-132]. We have a comparatively poor understanding of the distributions of α-tubulin isotypes, with the exception of TUBA1A. Tuba1a is strongly and specifically expressed in the developing nervous system, and provides over 95% of the α-tubulin in the embryonic brain [16, 125]. TUBA1A expression and its regulation will be extensively discussed in subsequent sections. Spatial and temporal expression data on α-tubulin isotypes remain sparse and may need to be readdressed.

Tracking the expression of tubulin isotypes at the protein level is particularly challenging due to the high degree of homology, and will benefit from new approaches to selectively label isotypes without impairing their functions.

Why different cell types express specific tubulin isotypes is a long-standing question. On one hand, the genetically-encoded differences between isotypes could impart functional differences within microtubule networks. The strongest evidence for specific functional roles for

α-tubulin isotypes comes from studies of in Drosophila and C. elegans, which demonstrate isotype-specific requirements for generating proper axonemal structures within cilia and flagella

[133-135]. These findings underscore the possibility that isotypes may play specific roles in building complex microtubule architectures, and raise the question of whether specific isotypes could be required to build other complex microtubule structures, such as those in neurons. An alternative, but not mutually exclusive explanation for differential isotype expression is that it

18 provides a convenient mechanism to regulate the levels of tubulin protein in a cell. This is a

particularly important challenge considering that cells must balance the levels of α- and β-tubulin

to form heterodimers, and excess monomer, particularly β-tubulin, can be toxic [136-138].

Studies investigating tubulin isotypes have provided some answers to these questions, but our

overall understanding of isotype biology is still in its infancy.

Tuba1a Expression Pattern

TUBA1A was originally identified in a cDNA library from human fetal brain tissue [139].

Subsequent northern blot studies in cell lines showed that TUBA1A is expressed in neural-

derived cells and is particularly abundant in cell lines with characteristic long cytoplasmic

processes (such as adherent neuroblastomas), but not in cell lines that exhibit small, round

morphologies [140]. Since then, numerous groups have tracked the expression of TUBA1A and

its vertebrate homologues to neuronal populations after terminal mitosis, as they begin to extend

long processes.

A series of classic studies elucidated the expression program of mouse Tuba1a mRNA across tissues and developmental stages. The earliest study investigating mouse Tuba1a expression via northern blot with probes recognizing the distinct Tuba1a 3′-UTR provides the most extensive inquiry into which organs express Tuba1a postnatally [125]. This study identified

Tuba1a mRNA in the adult brain, lung, and at low levels in the testes; however, only alternative

α-tubulin mRNAs were detected in the heart, kidney, liver, muscles, spleen, stomach, and

thymus. A time course analysis of brain-derived samples revealed that Tuba1a is abundant

through early postnatal stages, begins to decline by postnatal day 10 (P10), but still is expressed

beyond P32 through adulthood [125]. Subsequent studies sought to elucidate the program of

19 Tuba1a expression in the brain; to our knowledge there are no other published studies confirming the presence of Tuba1a in the adult lung or testes.

Much of our understanding of Tuba1a expression in the developing brain comes from

studies conducted by Miller and colleagues, who inserted lacZ behind the native Tuba1a

promoter and used this Tuba1a:nlacZ transgenic mouse to monitor Tuba1a expression in vivo

[14, 27, 28]. The Tuba1a:nlacZ transgenic mouse and a later transgenic reporter line using

Tuba1a promoter-driven EYFP expression [141] were used to characterize Tuba1a expression

patterns in rodents. These studies show that Tuba1a is expressed highly throughout the nervous

system over the course of development, starting around embryonic day 9.5 (E9.5) and continuing

through early postnatal stages [14, 28, 125, 141]. This trend holds true through the developing

central and peripheral nervous system, with Tuba1a induction onset correlating to neurogenesis

of numerous neuronal structures including the forebrain, midbrain, hindbrain, spinal cord, retina,

and cranial nerves, among others [14]. Primary cultures generated from the transgenic reporter

mice revealed that Tuba1a is expressed in cells positive for neuronal markers neuron-specific

enolase (NSE) and βIII tubulin, but not in cells positive for stem cell or glial markers [14, 28,

141, 142]. More recent RNA-seq studies from the Barres lab show that Tuba1a remains the most

abundant α-tubulin isotype mRNA in neurons late in brain development (P7) [126]. Based on

these studies, the current model is that Tuba1a expression is induced coincident with or shortly after the terminal cell division that gives rise to neurons, and provides >95% of the α-tubulin mRNA in these cells [16].

While much of the RNA probing data suggest that Tuba1a expression is limited primarily

to neurons, the recent RNA-seq dataset contends that Tuba1a may be expressed in more cell

types than the earlier studies indicate [126]. As stated previously, numerous studies claim that

20 Tuba1a mRNA is absent in stem cell, glial, and neuron progenitor populations, limited only to early born neurons [14, 28, 141, 142]. However, the RNA-Seq data suggest that Tuba1a may in fact be expressed in numerous cell types found in the brain, with Tuba1a expression equally high in newly-formed oligodendrocytes as it is in neurons at P7. As immunohistochemistry double labeling studies suggest that oligodendrocytes in embryonic and adult mouse do not express

Tuba1a [28, 141, 142], the contradictory expression of Tuba1a in glial populations calls for additional studies to conclusively uncover what cell types contain Tuba1a.

After brain development, Tuba1a mRNA levels persist, albeit at lower levels, in the adult mouse brain. Which adult cell populations express Tuba1a is somewhat controversial, due to conflicting data. Bamji and Miller found that Tuba1a mRNA was panneuronal in the adult mouse brain, with the highest levels found in neuronal populations that have the potential for morphologic growth, such as neurons of the piriform cortex and hippocampus [27]. In contrast,

Coksaygan and colleagues only detected Tuba1a mRNA in some regions of the adult brain

[141]. How experimental differences might contribute to these discrepancies is not immediately clear.

While much of the Tuba1a expression data have been conducted in mouse, studies in rat models yield similar findings. Rat Tuba1a mRNA is highly enriched in the embryonic nervous system and is less abundant in adult brain [16]. Additionally, the rat data also support the idea that Tuba1a is induced in neurons undergoing neurite extension, such as in vivo at the cortical plate as well as in vitro in PC12 cells induced to differentiate with Nerve Growth Factor (NGF) treatment [16].

Interestingly, studies of the zebrafish tuba1a yield results that overlap with those from mammalian models, but with several important differences. Whereas zebrafish tuba1a is

21 dramatically induced in the developing nervous system and declines coincident with the

maturation, it is maintained at high levels in progenitor cells in the retinal periphery, lining the

brain ventricles, and around the central canal of the spinal cord [143]. When cultured in vitro, these tuba1a-expressing cells divide and give rise to new neurons, suggesting that zebrafish may express tuba1a in neuronal progenitors as well as newly-born neurons [143]. Consistent with this notion, morpholino-based knock down of zebrafish tuba1a suppresses central nervous system development and leads to fewer differentiating neurons [144]. Intriguingly, Ramachandran et al. generated transgenic fish to conditionally and permanently label tuba1a-expressing cells, and found transient tuba1a expression in neural progenitors, skeletal muscle, heart, and intestine progenitors [145]. This experiment suggests that tuba1a expression is not neuronally limited in

zebrafish.

These expression studies strongly suggest that TUBA1A is highly induced in neurons as

they undergo cellular morphology changes to generate cytoplasmic processes. This suggests a

simple model for why neurons, but not other cell types, use the conserved expression program

for TUBA1A—neurons need significantly more α-tubulin at the time when they are maturing to

form the long processes necessary to wire the nervous system.

Mechanisms Regulating TUBA1A Expression

While questions remain regarding which cell populations express TUBA1A, it is clear that

expression is carefully regulated during development and adulthood. What pathways control this

expression program for TUBA1A? A variety of factors and conditions that mediate TUBA1A

mRNA production have been identified, and growth factors involved in neuronal differentiation

play a prominent role.

22 Several studies have shown that growth factors are sufficient to stimulate Tuba1a expression. For example, PC12 cells from rats can be induced to differentiate into neurons with the addition of NGF, and consequently Tuba1a mRNA levels increase as the differentiated PC12 cells begin to actively extend neurites [16]. Similarly, NGF treatment of neonatal rats causes a 5-

10-fold increase of Tuba1a mRNA during a developmental period when Tuba1a levels normally

decrease [146]. In addition to NGF, Fibroblast Growth Factor (FGF) and Platelet-Derived

Growth Factor (PDGF) also stimulate Tuba1a expression. FGF was first shown to stimulate

Tuba1a expression in rat sensory neurons, where treatment with FGF-1 increases Tuba1a mRNA

levels by nearly ten-fold, and causes a concomitant increase in neurite outgrowth [147]. These

findings suggest that exogenous stimuli such as growth factors may be sufficient to induce

TUBA1A expression, at least in certain cell types.

The most detailed evidence of the regulatory pathways that act downstream of growth

factors to increase Tuba1a transcription comes from a study using mouse primary cultures of

E12–E13 cortical progenitors [148]. This study revealed a phosphorylation cascade initialized by

FGF and PDGF activation that ends in drastic upregulation of Tuba1a, and ultimately

neurogenesis. Inhibiting factors in this pathway, MEK or C/EBP, leads to a significant decrease

in Tuba1a expression and stops neurogenesis. Conversely, introducing an activated C/EBP

phosphomimetic enhances Tuba1a expression and promotes neurogenesis. Furthermore, this

study identified three C/EBP binding sites in the conserved TUBA1A promoter, and revealed that

direct C/EBP binding to these promoter regions greatly enhances Tuba1a transcription.

Interestingly, inhibiting the Tuba1a neurogenesis pathway causes the would-be neurons to take

on an astrocyte fate. This reveals that the same progenitors can be induced to become either

23 neurons or astrocytes, and that Tuba1a activation is an important step in promoting the neuronal fate.

While the studies described above implicate common neurogenesis growth factors in

Tuba1a expression, the specific requirements for Tuba1a activation remain largely unexplored.

However, these data provide an additional link between the presence of Tuba1a and neurogenesis, further promoting the idea that Tuba1a may be an important regulator of neuronal fate.

Tuba1a in Regeneration

In addition to the Tuba1a developmental expression pattern, Tuba1a expression is also elevated following nerve injury [31, 149-151]. Although Tuba1a mRNA expression is low in the adult, decreasing approximately 20 fold from developmental levels, its expression is once again rapidly induced following nerve injury [16, 30]. Its unique temporal expression pattern has established a clear role for Tuba1a as a growth-associated neuronal a-tubulin. The induction of a developmental a-tubulin, such as TUBA1A, during regeneration supports the idea that developing and regenerating neurons may utilize common mechanisms.

In contrast, evidence emerged suggesting that Tuba1a induction following nerve injury arises through a different mechanism than developmental induction. Several studies suggest that

Tuba1a expression is repressed by homeostasis in adult neurons, but if normal processes are disrupted, Tuba1a expression is induced. For example, Wu et al. suggest that Tuba1a gene expression is induced by the breakdown of axon trafficking because in their system Tuba1a could be induced by simply blocking fast axonal transport [33, 152]. Tuba1a repression could be mediated through intrinsic or extrinsic mechanisms that depend upon intact axonal trafficking.

24 Specifically, the neuron could be constantly monitoring its axonal integrity using intrinsic cues,

such as backup of protein cargoes that would occur within the axon upon injury, or extrinsic cues

such as constant uptake of growth factors like Ciliary Neurotrophic Factor (CNTF) from nearby

Schwann cells [30, 33]. Alternatively, it has been proposed that repression could be maintained

through physical contacts between neurons and neighboring cells [152]. However, current

evidence does not support this model, as studies have found that loss of contact with either the

basal lamina or the neuron’s target structure alone is insufficient to induce Tuba1a expression

[29, 30]. To date, the exact mechanisms by which Tuba1a gene expression is induced after nerve injury remain unclear, however it has become apparent that Tuba1a a-tubulin is of particular importance to growing neurons, regardless of age.

The expression of Tuba1a during regeneration after nerve injury provides an opportunity to study regenerative capacity in neurons. While peripheral nerves are able to regenerate after injury, most neurons of the central nervous system do not have this capacity [153]. What allows peripheral axons to regenerate, while central axons cannot? Many of the initial studies identifying Tuba1a expression in response to injury focused on the peripheral nervous system,

however, increases in Tuba1a expression have been noted following injury to central nerves as well [31]. This observation provided an opportunity to probe the differences in regenerative capacity between the central and peripheral nervous systems. Interestingly, one study found that gene expression was broadly similar between peripheral nerves that regenerate after injury and proximal central nerves that do not [31]. Importantly, in peripheral neurons, Tuba1a expression remained elevated until regeneration was complete. In central neurons, which did not regenerate,

Tuba1a expression remained elevated for as long as two weeks post injury, even after levels of other injury-related cytoskeletal proteins had returned to normal [31]. Other studies have

25 demonstrated that Tuba1a expression correlates with the timing of regeneration, and that in general Tuba1a will remain elevated until the axon is able to regenerate [29, 32]. Thus, these data show that specific central neurons have regenerative capacity, and that likely in these cases it is an unfavorable environment that prevents complete regeneration. Conversely, in a different population of central neurons, Tuba1a is never upregulated after injury [149]. This provides evidence of heterogeneity in regenerative capacity between different populations of central neurons, indicating that some neurons may lack the capacity to regenerate, regardless of whether the environment is conducive to regeneration [149]. Further, Tuba1a can become upregulated after injury, even in uninjured regions of the nervous system. For example, elevated Tuba1a expression was not limited solely to injured peripheral axons, but neighboring axons frequently also displayed increased Tuba1a mRNA [29, 32, 146]. Additionally, elevated Tuba1a expression was found in the red nucleus, a brainstem nucleus that sends axons to the spinal cord, following a spinal cord compression injury [150]. The studies outlined here show a selective upregulation of

Tuba1a in regenerating neurons. The properties of TUBA1A that make it particularly suited to these functions are as yet unknown. However, it is interesting to speculate that, in line with the tubulin code model, combinations of different tubulin isotypes could produce microtubules that are preferentially suited for specific functions.

Microtubules in Developing and Adult Neurons

Microtubules in Neural Development

As outlined in previous sections, the microtubule cytoskeleton is capable of facilitating a spectrum of cellular processes. Microtubules play numerous important roles during brain development, particularly in neurons, where the microtubule cytoskeleton has been intensely

26 studied. Once neuronal progenitors exit the cell cycle to become neurons, diverse microtubule- based maturation stages must occur for the neuron to correctly extend dynamic neurites, migrate to the proper position, form and guide long axons, set-up synapses, and sustain the diverse regions of the neuron. Microtubules are essential for these phases of neuronal maturation and are differentially regulated by numerous MAPs to perform diverse neuronal microtubule-based functions.

Microtubules interact with numerous MAPs during each stage of neuronal maturation to properly perform various cellular tasks. During neurite initiation, dynamic actin forms lamellipodia that become stabilized by invading microtubules [154]. Interestingly, the dynamic properties of microtubules at this stage may inform which neurite becomes the future axon.

Locally stabilizing microtubules in a particular neurite with stabilizing drugs results in that neurite becoming the axon [155]. Later, as neurites mature, microtubules become stabilized by

MAPs such as tau and MAP2 [156, 157]. The presence of tau helps identify the neurite as the axon, while MAP2 identifies a dendritic fate. At this stage, motors such as kinesin-1 and dynein are important in sorting and pushing microtubules to the end of the neurite [158].

As development proceeds, neuronal microtubules begin to exhibit a distinct polarity and decoration by MAPs depending on which type of process they inhabit. Microtubules within the axon become uniformly oriented with their plus-ends directed away from the soma, while dendritic microtubules retain a mixed polarity [159, 160]. Dynein has been implicated in establishing and maintaining this plus-end-out (away from the soma) orientation in axons [158].

Plus-end-out polarity is critical for axonal transport, discussed below. The quantity and diversity of MAPs in the neuronal environment, and the interplay between them makes understanding each MAPs’ role difficult. For example, XMAP215 directly regulates microtubule plus-end

27 dynamics in vitro, but in the growth cone it regulates the linkage of translocating microtubules to the F-actin network, thereby constraining microtubule growth velocity [161]. Future studies are needed to determine how microtubules are regulated throughout each stage of neuronal maturation, with attention given to how microtubule dynamics are regulated by the plethora of

MAPs present during each developmental stage.

Microtubule function is critical for neuronal migration, which is required to form the layers of the cortex and many other structures in the brain. During neuronal migration, microtubules extend into the leading process where they help to steer the protrusive growth cone at the end of the axon [162]. Microtubules also generate force to move the nucleus, in a process known as nucleokinesis. Here, microtubules extend back from the centrosome to form a cage- like network around the nucleus [163]. Dynein and its regulator LIS1 generate pulling forces to draw the nucleus toward the centrosome and move the centrosome toward the leading process

[164]. Consistent with the important roles of the microtubule network in neuronal migration, mutations in LIS1 and the microtubule regulator doublecortin/DCX are associated with migration disorders that give rise to brain malformations [165].

Adult Neuronal Microtubule Function

One of the most important functions of microtubules in adult neurons is to facilitate the efficient trafficking of organelles, proteins, mRNAs, and other cargoes across long distances in the axon. To appreciate the importance of efficient transport, consider that the distal regions of an axon can be as far as one meter away from critical protein and mRNA synthesis occurring in the soma. The unique microtubule polarity of the axon that is established early in development organizes transport in the anterograde and retrograde directions, with the help of motor proteins

28 such as kinesin and dynein that move towards the plus- or minus-ends, respectively. Kinesins, particularly kinesin-1 and 3 family members, carry cargoes towards the axon terminal by walking towards the plus ends of axonal microtubules [64]. Conversely, dynein carries cargoes back toward the soma by walking towards the minus ends [166]. The combination of uniform microtubule orientation and processive motility by directional molecular motors allows neurons to effectively transport cargoes across large distances.

In addition to facilitating intracellular transport, neuronal microtubules play an important role in synaptic transmission. Neurons are post-mitotic cells, but they still require mechanisms to respond to and interact with their environment. In particular, contact points between pre-synaptic termini and post-synaptic dendritic spines are known to change in response to neural activity, allowing neurons to prioritize productive connections and eliminate unnecessary ones. Dendritic spines undergo structural increases or decreases in size called potentiation and depression, respectively, following bursts of synaptic transmission. This phenomenon is called synaptic plasticity and is widely regarded to be the cellular correlate of learning and memory [167-169].

Dendritic spines are actin-rich structures which typically do not contain microtubules. However, unlike the relatively stable axonal microtubule population, microtubules within the dendritic shaft have been shown to retain dynamic properties in mature neurons [76, 170-173]. Work by

Kapitein et al. identified that dynamic microtubules transiently polymerized into dendritic spines, an occurrence that was suppressed under conditions of synaptic depression [5]. Further, depletion of microtubules, or pharmacological inhibition of microtubule polymerization prevented synaptic potentiation in vitro and in vivo [3, 4, 174, 175]. Collectively, these data implicate dynamic microtubules as key players in driving the structural changes that underlie synaptic plasticity.

29 Neurons require elaborate networks of microtubules to support their complex

morphologies, and to perform a variety of critical functions. In developing neurons, dynamic

microtubules are necessary to facilitate essential functions such as neuronal morphological

changes that underlie differentiation and migration [2, 176, 177]. Adult neurons, however,

require much more stable networks of microtubules, particularly in the axon. While specific

properties of microtubules have been observed to change between newly born and adult neurons,

the factors that govern this switch in microtubule function over time are not fully known. The

mechanisms by which microtubule function can be regulated to support diverse cellular tasks are

under active investigation, but evidence suggests that microtubule impairment has devastating

consequences for neuronal function. In the following section, I will discuss known roles for

microtubules in disease of the nervous system.

Microtubules in Nervous System Disease

Tubulinopathies Reveal Essential Role of TUBA1A in Brain Formation and Function

Recent studies investigating the genetic cause of brain malformation disorders have

revealed that heterozygous, missense mutations to TUBA1A and other neuronal tubulin isotypes

play an important role in brain development. First shown by Keays et al. in 2007, sequencing of

tubulin genes has proven fruitful in uncovering the source of genetic variation in numerous

patients exhibiting complex neurological and physical phenotypes with cortical malformations.

From these genotype-phenotype analyses, TUBA1A is recognized as vital for neurodevelopment based on the devastating effects observed in patients containing heterozygous, de novo TUBA1A

missense mutations [17-22, 178-181]. Brain malformation disorders caused by mutations to

TUBA1A and other neuronally expressed tubulin isotypes are collectively termed

30 “tubulinopathies”, and lead to severe cortical abnormalities, mental retardation, and commonly

epilepsy and paralysis [17-22, 178-181]. Patients containing TUBA1A mutations exhibit a wide

variety of cerebral cortex malformation phenotypes, including lissencephaly, pachygyria,

microlissencephaly, and polymicrogyria. While known genetic causes of these phenotypes hint at

which developmental processes may be disrupted, little is known about how the TUBA1A mutations disrupt microtubule functions, or even how these disruptions could cause the larger- scale cellular and tissue problems seen in patients. The wide variety of brain malformations observed in patients leads to the prediction that different missense mutations in TUBA1A may disrupt different phases of neuronal development. Disease-associated mutations to TUBA1A therefore provide a valuable opportunity to investigate the numerous neurodevelopmental stages that require TUBA1A, and how microtubules must be regulated for each stage to occur appropriately.

The identification of neurodevelopment disorder-associated TUBA1A mutations supports the hypothesis that this particular α-tubulin isotype is essential for neuronal maturation; however how can subtle changes to one α-tubulin protein lead to drastic changes in the formation of the brain? Expression studies of TUBA1A mRNA make it clear that Tuba1a is by far the most prevalent α-tubulin isotype in the embryonic nervous system, accounting for more than 95% of

α-tubulin mRNA [16]. Thus, mutations to TUBA1A could greatly affect the available pool of neuronal tubulin. Tubulin mutations can act to alter numerous tubulin/microtubule characteristics and functions, but most of the possible changes fit into three general categories: 1) disrupting tubulin folding and heterodimer formation, thereby depleting the pool of α-tubulin that is competent to assemble microtubules; 2) disrupting polymerization activity and/or microtubule dynamics regulation, which could either deplete the pool of assembly-competent α-tubulin or

31 alter dynamics once mutant heterodimers have formed the microtubule lattice; or 3) assembling into microtubules but altering microtubule function by disrupting interactions with MAPs and motors. Identifying which category TUBA1A mutations fit into will greatly increase our understanding of tubulinopathy disease progression. This task is not easy, however, as structure- function predictions for tubulin residues are rarely straightforward. Tubulin is a complex protein that interacts with numerous binding partners and undergoes long range conformational changes as part of its principle biochemical activity. Therefore, knowing the location of a change in the tubulin sequence does not necessarily give insight into its functional consequences. This underscores the need for functional studies that can test and refine structural predictions.

However, despite the growing list of tubulinopathy mutations, few studies provide insight into how individual mutations impact the tubulin protein and its function in vivo. Future studies must be performed to fill in the missing molecular, mechanistic steps between the known TUBA1A mutations and the final brain phenotype.

While few studies have specifically addressed how mutations that disrupt TUBA1A cause cortical malformations, understanding the known mechanisms can provide clues into how attributes of TUBA1A contribute to normal brain development. However, there is much more work to be done to elucidate the mechanistic role of TUBA1A. Several different aspects of

TUBA1A function could go awry to disrupt cellular processes and ultimately lead to abnormal brain development. For example, interference with neuronal migration can lead to lissencephaly, but not all the aspects of TUBA1A that are important for neuronal migration are known. For example, a mutation to TUBA1A could cause haploinsufficiency through many mechanisms, including disruptions to TUBA1A folding, heterodimer formation, or microtubule assembly.

This could disrupt neuronal migration due to a pool of incompetent tubulin dimers forming less

32 stable microtubules. Alternatively, specific TUBA1A mutants could appropriately assemble into

microtubules but then cause dominant changes that “poison” the microtubule network. The

dominant disruption could be achieved through changes to microtubule dynamics or disruption

of binding of MAPs involved in migration.

Mouse Models Provide Insight into neuronal microtubule function

TUBA1A variants are associated with severe cortical malformation phenotypes in humans

and comprise the majority of identified human tubulinopathy mutations [17-22, 178-181].

However, it is impossible to establish a mechanism of action for how TUBA1A variants disrupt

brain development based on clinical descriptions alone. Insight to mechanistic questions can be

gained from examining available mammalian models of Tuba1a mutations. Homozygous

Tuba1a deletion results in severe brain defects including disruption of cortical layering, enlargement of cerebral and third ventricles, and a reduction in the size of the basal ganglia

[182]. In contrast, no brain defects were reported in heterozygous Tuba1a knock out mice [182].

The same group also generated a truncation mutant allele which creates a premature stop at position 215 (Tuba1a-R215*)[182]. Homozygous Tuba1a-R215* mice have severe brain defects that are similar to homozygous Tuba1a deletion, but any consequences for brain development in heterozygous Tuba1a-R215* mice were not reported [182]. These findings suggest that while homozygous deletion of Tuba1a leads to severe defects in cortical migration, heterozygous depletion of Tuba1a may not disrupt cortical migration.

To date, two Tuba1a missense mutations have been generated through N-ethyl-N- nitrosourea (ENU)-induced mutagenesis in mice and investigated for their impact on neurodevelopment [180, 183]. The Tuba1a mutational studies in mice do not represent variants

33 identified in human tubulinopathy patients. One missense mutation causes a serine to glycine

substitution at residue 140, Tuba1a-S140G [180]. Tuba1a-S140G heterozygous mice have subtle

alterations to cortical layering and contain heterotopic neurons within the pyramidal

hippocampus, both of which are attributed to mild neuronal migration defects [180]. Importantly,

the impairment of neuronal migration in heterozygous Tuba1a-S140G mice is extremely mild when compared to the migration errors and resulting brain malformations seen in human patients with heterozygous TUBA1A mutations. Additionally, Tuba1a-S140G heterozygous mice display

an intriguing behavioral phenotype that consists of locomotor hyperactivity, impaired

hippocampal-dependent spatial working memory, and reduced anxiety-like behavior [180]. A

second allele, Tuba1a-N102D (Tuba1aND), results in an asparagine to aspartic acid substitution at residue 102. Tuba1aND homozygous mice have dramatic defects in brain development including abnormal cortical lamination and reduction in the size of olfactory bulbs, indicative of impaired neuronal migration [183]. In contrast, Tuba1aND heterozygous mice are viable, undergo normal cortical lamination, and do not exhibit any evidence of neuronal migration errors [183]. From a mechanistic standpoint, Tuba1a-R215*, -S140G and -N102D variants are all predicted to reduce the levels of functional Tuba1a, despite the observed difference in heterozygous phenotypes

[180, 182, 183]. The Tuba1a-R215* allele is predicted to generate a loss-of-function Tuba1a allele, due to the truncating nature of the mutation. Thus, it is notable that heterozygous Tuba1a-

R215* had few reported consequences for brain development, but more thorough morphological analysis of heterozygous Tuba1a-R215* brains is needed to determine whether all aspects of brain development occur normally in this model. The Tuba1a-S140G mutation impairs the ability of α-tubulin to bind GTP, which disrupts tubulin heterodimer formation and reduces the functional tubulin pool [180]. The Tuba1a-N102D substitution also destabilizes the heterodimer

34 and impairs incorporation of mutant Tuba1a into the microtubule lattice [183]. However, the divergent neurodevelopmental and behavioral phenotypes associated with the heterozygous

S140G and N102D mutant mice suggest that these variants may impact neuronal microtubule function via several differing mechanisms, not solely through loss-of-function (LOF). While

S140G and N102D are predicted to impair heterodimer formation, the degree to which each variant reduces the available tubulin pool within the brain is unclear. Beyond a pure LOF mechanism, phenotypic divergence between these variants could also arise if a significant amount of mutant Tuba1a protein populates the lattice, impairing microtubule function in distinct ways.

All four Tuba1a alleles in mice are reported as LOF, and if that is the primary mechanism of action much can be inferred about the requirement of Tuba1a in neurodevelopment from these models. When the aforementioned mutations are heterozygous, they lead to generally mild, if any, neurodevelopmental phenotypes. In contrast, all of the homozygous Tuba1a mutations are lethal and display cortical layering defects in addition to grossly malformed brains [180, 182,

183]. This suggests that one copy of Tuba1a is sufficient to support cortical migration, but that disruption of both copies has drastic consequences for brain development. Heterozygous brain morphology was not extensively investigated in these models, therefore it is unclear whether neuronal microtubules with reduced Tuba1a can support long-distance axon guidance or synaptic connections. While it is possible that the heterozygous mouse phenotype is less severe due to inherent species differences between human and mouse brain development, data from mouse models could suggest that heterozygous loss-of-function mutations in TUBA1A may not disrupt cortical migration to cause drastic brain morphology changes in humans. The striking absence of human TUBA1A variants that cause deletions, reading frame-shift, or truncations in patients with

35 severe brain malformations further supports the conclusion that mutations that have been

identified in patients with brain malformations may not disrupt brain development by a LOF

mechanism. Rather, the impact of heterozygous, de novo, LOF mutations to TUBA1A may be less severe, and thus be less examined in pediatric patients than heterozygous, de novo, dominant

mutations.

LOF in one copy of Tuba1a is sufficient to support cortical migration and survival in

rodent models [180, 182, 183], but the impact of Tuba1a haploinsufficiency on axon pathfinding,

synapse formation, and mature neuron function remains largely unexplored. Tuba1a comprises the majority of a-tubulin mRNA while neuronal microtubule networks are being built, and

Tuba1a protein has been shown to turn over slowly in the mammalian brain [127]. Thus, Tuba1a may be present in mature neuronal microtubules, and may influence microtubule function in the adult brain. In Chapter III, I will interrogate how Tuba1a haploinsufficiency impacts long-term cellular and behavioral function in the adult mammalian nervous system. Mature neurons depend heavily upon microtubules for intracellular transport and maintenance of synaptic connections, two aspects of neuronal function which are frequently disrupted in neurodegenerative disease

[34, 35, 37, 40, 184]. In the following section, I will discuss evidence that microtubules become dysregulated in neurodegenerative disease, and explore how microtubule deficits can impact mature neuronal function.

Microtubules in Neurodegeneration

Neurodegenerative diseases such as Alzheimer’s disease, Parkinson’s Disease, and

Amyotrophic Lateral Sclerosis (ALS), among others, affect a large portion of the aging population. In recent years, microtubule dysfunction has been recognized as a facet of many

36 neurodegenerative diseases. While it is unclear if microtubule dysfunction occurs as a cause or consequence of neurodegeneration, the observed microtubule dysfunction provides a potential new target for therapeutic intervention in these diseases. In several neurodegenerative diseases, microtubules are lost over time resulting in decreased microtubule density [40]. For example, an overall decrease in a-tubulin was found in the post-mortem brains of Alzheimer’s disease patients [185]. In addition to microtubule loss, normal microtubule stability becomes compromised in several neurodegenerative diseases [186, 187]. The mechanisms of microtubule dysfunction in neurodegeneration remain unclear, however aberrant protein activity in many neurodegenerative diseases could compromise microtubule integrity. For instance, abnormal phosphorylation of MAP Tau is a key feature of Alzheimer’s disease, and related neurodegenerative diseases known as tauopathies [188]. Abnormally phosphorylated Tau is unable to properly bind and stabilize microtubules, potentially allowing neuronal microtubules to become more prone to disassembly [189, 190]. Further, in Parkinson’s disease, abnormal redox activity leads to decreased acetylation of microtubules via augmented activity of the tubulin de- acetylase, Sirtuin-2 [191]. Similarly, in familial dysautonomia, the disease-causing mutation leads to downstream activation of a-tubulin de-acetylase HDAC6, leading to an overall decrease in acetylation of a-tubulin [192]. Although acetylation itself does not confer any stability to microtubules, decreased acetylation of microtubules could disrupt interactions between microtubules and other proteins [191, 193]. In addition, hereditary spastic paraplegia, a degenerative disorder that primarily affects motor function, involves mutation of the microtubule-severing protein spastin [187]. Together, these studies provide evidence that microtubule dysfunction is a common element of a wide array of neurodegenerative diseases with different pathologies.

37 Changes in microtubule dynamics occur in neurodegenerative disease. In what ways could changes in microtubule stability contribute to neurodegeneration? Microtubules facilitate axonal transport in adult neurons. Disruption of axonal transport is sufficient to cause neuronal degeneration [194, 195]. Furthermore, breakdowns in axonal transport have been documented in different neurodegenerative diseases [34, 184, 196]. Additionally, changes to microtubule dynamics and defects in axonal transport often occur together. For example, an increase in

2 microtubule turnover, as measured by H2O incorporation, preceded breakdowns in axonal trafficking in a model of ALS [197]. Stabilizing microtubules by pharmacological means effectively rescued axonal transport defects, suggesting that transport defects in a model of ALS occur as the result of changes to microtubule stability [197]. Further, it has been shown that treating cells with 1-methyl-4-phenylpiridinium (MPP+), a drug used to induce Parkinson’s

Disease-like pathology in neurons, leads to a selective loss of dynamic microtubules, and enrichment of stable microtubules, that also precedes a decline in axonal transport efficiency

[39]. These data provide evidence that changes in microtubule dynamics often precede breakdown of axonal transport in neurodegeneration, and that pharmacologic intervention to stabilize microtubules can prevent transport defects.

While microtubule dysfunction has been clearly implicated in a large number of neurodegenerative diseases, the question remains as to whether microtubule dysfunction can be considered a causative factor in these diseases. In recent years, evidence has emerged to support the idea that disruption of normal microtubule function can be sufficient to induce neurodegeneration. Most notably, exome-wide sequencing performed on individuals with familial ALS identified disease-associated mutations in the neuronal a-tubulin gene, TUBA4A

[198]. Mutations that disrupt TUBA4A both decreased the efficiency of tubulin incorporation

38 into the microtubule lattice, and also prevented efficient re-assembly of microtubules following depolymerization [198]. Importantly, this study provided evidence that mutations affecting microtubule function are capable of causing neurodegeneration in humans. Mutations that disrupt a member of the tubulin folding pathway, tubulin binding cofactor E (TBCE), can cause progressive peripheral motor neuronopathy in mice [199-202]. In this case, the mutant TBCE protein is not functional and is degraded, leading to progressive degeneration of motor neurons, and sensory neurons [203]. Degeneration in TBCE mutant mice relies on the inability of mutant

TBCE to maintain microtubule networks, particularly in motor neurons where TBCE is highly expressed [200]. However, the mechanism of action in this model has not been completely elucidated, and the possibility remains that TBCE could have other functional activities that have yet to be discovered. These studies provide compelling evidence that microtubule dysfunction can cause neurodegeneration.

Therapeutic intervention may target microtubules to prevent or slow neurodegeneration.

Microtubule-stabilizing drugs have shown promise in animal models of neurodegeneration [36,

204-206]. However, broadly stabilizing all microtubules could have unwanted consequences in humans [207]. Alternative microtubule targeting therapies are also being explored. These include drugs that target microtubule severing proteins, as well as therapies that add to the labile pool of microtubules [41, 96, 208]. Overall, the array of microtubule properties that are disrupted in different neurodegenerative diseases provide an exciting opportunity to tailor treatments to specific aspects of microtubule dysfunction in different disorders. Regardless of whether microtubule dysfunction is causative or consequential to degeneration, therapeutic intervention at the level of microtubule dysfunction could be advantageous for preserving neuronal health. It

39 will be interesting to see how such therapies continue to evolve as more information emerges

about the specific role of microtubules in neurodegenerative disease.

Conclusions

Microtubules are dynamic cytoskeletal polymers that fulfill diverse roles in different cell

types. Neuronal microtubules are crucial throughout all stages of brain development and support

the dramatic morphological changes required for differentiation, migration, neurite branching,

axon guidance, and synapse formation. In mature neurons, microtubules form organized

highways for intracellular cargo trafficking and promote the structural plasticity underlying

synaptic potentiation and depression. The mechanisms by which microtubules are regulated to

perform such diverse functions in neurons are under active investigation, but differences in

tubulin isotype expression, tubulin PTMs and MAP interactions act as a tubulin code to regulate

microtubule network function. TUBA1A is the major a-tubulin isotype expressed in the developing brain and is critical for brain development. Less is currently known about its role in mature neurons. Understanding how TUBA1A contributes to neuronal microtubule function will help elucidate its role in diseases of the nervous system. In the following chapters I will investigate requirements for TUBA1A in the developing and adult nervous system, and interrogate the role of TUBA1A in disease.

40 CHAPTER II

TUBA1A PLAYS AN ESSENTIAL ROLE IN AXON PATHFINDING

Introduction

Mammalian brain development is a complex process that requires precise coordination of

multiple cell types and extracellular cues to form a fully specified tissue. Despite many advances

in understanding the cellular and molecular players involved in brain development, there is still

much that remains unknown. Insights into the molecular pathways governing neurodevelopment

can be gained from studying genetic mutations that disrupt specific aspects of brain

development. Severe cortical and neurodevelopmental phenotypes associated with mutations that

disrupt tubulin genes, termed tubulinopathies, have recently been described in humans [17-19,

22]. Tubulinopathy mutations cause a spectrum of neurodevelopmental phenotypes, but

frequently involve cortical malformations such as lissencephaly, agenesis or hypoplasia of the

corpus callosum, and cerebellar hypoplasia [17, 18, 22, 209]. Recent studies of human

tubulinopathy mutations have revealed that each variant may impact different aspects of

microtubule function, such as protein folding, polymerization competency, and microtubule-

associated protein (MAP)-binding, among others [23-25, 210]. Tubulin mutations can therefore

be used to interrogate the requirement for specific aspects of microtubule function throughout

neurodevelopment.

Developing neurons must migrate to the correct location, extend axons to meet

sometimes distant binding partners and form functional synaptic connections. Throughout this

process, neurons undergo dramatic morphological changes that require coordinated interaction

between the cytoskeleton and the extracellular environment. In post-mitotic neurons, microtubule

polymers made of a/b-tubulin dimers, facilitate nucleokinesis and cellular migration, support

41 growth cone navigation, promote axon formation and form the tracks upon which intracellular

trafficking occurs [211-214]. The microtubule network needs to be precisely controlled to fulfill

diverse functions in neurons. Microtubule properties can be modulated through post-translational

modifications (PTMs), association with MAPs and through the particular tubulin genes, or

isotypes, that a cell expresses [215]. The human genome contains at least nine unique α- and ten

unique β-tubulin genes [7, 8]. The a-tubulin isotype encoded by the gene TUBA1A is abundant in

the brain, and is the most highly expressed a-tubulin in post-mitotic, developing neurons [14, 16,

28]. TUBA1A mutations are highly represented in cases of human tubulinopathies [15],

suggesting that TUBA1A plays an important role in neurodevelopment. However, due to the high degree of sequence conservation between a-tubulin genes, it has been historically challenging to study TUBA1A function in vivo, due to the limited availability of tools.

Mouse models harboring mutations to Tuba1a can be used as tools to interrogate the function of Tuba1a in the context of the neuronal milieu. As tubulin genes are often required for life and the nucleotide sequence between isotypes is conserved, generation of mutant mouse lines to study Tuba1a function in vivo has been challenging. To date, only a handful of Tuba1a mutant mouse lines have been generated, three by ENU-induced forward genetic screens and one by site-directed CRISPR gene editing [180, 182, 183]. We previously showed that the ENU-induced

Tuba1aN102D allele impaired microtubule function in both S. cerevisiae and mice [183].

Homozygous Tuba1aND mice had severely impaired brain development and were neonatal lethal,

similar to phenotypes seen in the Tuba1anull and Tuba1a-R215* mutant mice, and to what is clinically described in tubulinopathy patients [15, 182, 183]. In patients, and in homozygous

Tuba1aND, Tuba1aNull and Tuba1a-R215* mice, cortical migration and commissural formation are severely disrupted making it difficult to infer whether axon pathfinding is a direct

42 consequence of altered TUBA1A function or if it is secondary to abnormal cortical layering and

migration. Tuba1aND/+ heterozygous mutant mice have reduced Tuba1a function during brain

development, which was sufficient to support survival and cortical migration resulting in normal

cortical layers [183], but does not support formation of commissures. Therefore, Tuba1aND/+

heterozygous animals can provide insight into how Tuba1a contributes specifically to axon

pathfinding.

Here, we show that a reduction in developmental Tuba1a causes specific impairments in

axon guidance through large brain commissures. Heterozygous Tuba1aND/+ brains fail to form the corpus callosum, anterior and hippocampal commissures due to apparent deficits in axon pathfinding through the midline. Cultured neurons from Tuba1aND/+ and wild-type cortices

revealed that Tuba1aND/+ neurons have shorter neurites than wild-type and grow slower overall, due to alterations in microtubule polymerization dynamics. Further, we demonstrate that

Tuba1aND/+ neurons fail to localize at least one critical developmental MAP to the developing

growth cone. Collectively, our data present evidence for precise mechanisms by which loss of

Tuba1a causes axonal pathfinding deficits during development.

Materials and Methods

Animals

All animal research was performed in accordance with the Institutional Animal Care and Use

Committee at the University of Colorado School of Medicine. All mice used were maintained on

a 129S1/C57Bl6 genetic background. Mice were kept on a 12:12 light:dark cycle with ad libitum access to food and water. Tuba1aND and wild-type littermate mice were maintained on water

ND/+ supplemented with 0.2g/L MgSO4 to promote Tuba1a survival and ability to reproduce. For

43 timed mating male and female mice were introduced overnight and separated upon confirmation of mating, which was then considered embryonic day 0.5. Male and female mice were represented in all studies. All mice were genotyped by PCR amplification of tail DNA followed by Sanger sequencing to differentiate homozygous or heterozygous Tuba1aND/+ mice from wild- type.

Histology

Mice were anesthetized and trans-cardially perfused with 0.1M NaCl and 4% paraformaldehyde

(PFA) for histology. Tissues of interest were dissected and post-fixed in 4% PFA. Tissue sectioning was performed on a CM1520 cryostat (Leica, Wetzlar, Germany) and 30µm cryosections were obtained for all histology. For luxol fast blue staining, sections from brain were stained using a 0.1% luxol fast blue solution. For immunofluorescence studies PFA-fixed tissues or cells were blocked in phosphate-buffered saline (PBS) containing 5% goat serum or bovine serum albumin (BSA) with 0.3% Triton-X 100. Primary and secondary antibodies were diluted in PBS containing 1% BSA with 0.1% Triton-X 100.

Electron Microscopy

Mice used for electron microscopy were perfused with 0.1M NaCl and 2.5% glutaraldehyde 4%

PFA, after which the brain was dissected and post-fixed in 2.5% glutaraldehyde 4% PFA overnight at 4°C. Following post-fixation, brains were sent for further processing and imaging by the CU School of Medicine Electron Microscopy Core facility.

Plasmids and Reagents

The hexahisitidine (His6) epitope tag was inserted in the α-tubulin internal loop region [216-

218]. Codon optimization for rattus norvegicus (https://www.idtdna.com/codonopt) was used to generate the His6 sequence CATCACCACCATCATCAC, which was inserted into the coding

44 region of human TUBA1A from the Mammalian Genome Collection (clone

ID:LIFESEQ6302156) between residues I42 and G43. Gibson cloning was used to insert the

gBlock of the TUBA1A internally tagged with His6 (TUBA1A-int-His6) into pCIG2 plasmid

linearized with NruI and HindIII. TUBA1A-int-His6 incorporation was confirmed by sequencing

across the complete TUBA1A coding region. The GFP-MACF43 vector was shared by Dr. Laura

Anne Lowery (Boston College) and Dr. Casper Hoogenraad (Utrecht University). Myr-

TdTomato plasmid DNA was shared from Mark Gutierrez and Dr. Santos Franco (University of

Colorado).

Cell Culture and Nucleofection

Cos-7 cells (Thermo Fisher Scientific, Waltham, MA; ATCC® CRL-1651™) were cultured in a

37°C humidified incubator with 5% CO2 in DMEM (Gibco) supplemented with 10% fetal bovine serum (Gibco), 1X sodium pyruvate (Thermo), and 1X penn/strep (Thermo). Cos-7 cells were transfected with 2.5µg of hexahistidine (His6) tagged TUBA1A plasmid DNA using

Lipofectamine 3000 (Invitrogen). For Cos-7 proteasome inhibition assays, 5µm Lactacystin A

[219, 220]. was added to normal culture medium for 24 hours, the day following transfection with TUBA1A-His6 constructs. Dissociated neurons were cultured from male and female P0-P2 mouse or rat cortices. Brains were removed and placed into HBSS (Life Technologies) supplemented with 1M HEPES (Life Technologies) and 1mM kynurenic acid (Tocris

Bioscience, Bristol, UK). Meninges were removed and cortices were dissected and cut into approximately 1mm pieces. Cortical pieces were triturated to a single-cell suspension using glass

Pasteur pipettes. Cortical neurons were plated onto 35mm Poly-D-Lysine coated glass-bottom culture dishes at a density of 350,000 cells (Willco Wells, HBSt-3522). For nucelofected mouse and rat neurons, 4 µg of plasmid DNA was introduced to 4x106 neurons using an AMAXA

45 nucleofection kit (VPG-1001, VPG-1003; Lonza). AMAXA-nucleofected cells were plated in

35mm glass bottom imaging dishes. Neurons were maintained in a 37°C humidified incubator

with 5% CO2 in phenol-free Neurobasal-A medium (Life Technologies) supplemented with B-27

(Thermo Fisher Scientific, Waltham, MA), Penn/strep (Thermo), GlutaMax (Thermo), 5ng/mL b-FGF (Gibco), and Sodium Pyruvate (Thermo).

RNA isolation + RTPCR

RNA was isolated from Cos-7 cells, 48-hours post-transfection using TRIzol Reagent (Thermo;

15596026). RNA concentration and purity were determined using a spectrophotometer, then cDNA was synthesized using the RT2 First Strand Kit (Qiagen, Hilden, Germany; 330401). qRT-PCR reactions were prepared with SYBR Green RT-PCR Master mix (Thermo; S9194) and run with a CFX Connect Real-Time System (Bio-Rad). Samples were run in triplicate, results were analyzed in Excel. All qPCR data presented in this chapter was normalized to expression of

GFP, which was present on the same plasmid as TUBA1A-His6 constructs. Wild-type TUBA1A mRNA quantity was set to = 1 and TUBA1AND relative mRNA quantity was presented relative to wild-type. For all qRT-PCR experiments 3 biological replicates were used per genotype.

Neuron Immunocytochemistry

DIV 3 primary cortical neurons were washed with PBS and fixed with a fixation solution of 4%

PFA and 0.2% glutaraldehyde in PBS for 15 minutes at room temperature. For tubulin extraction, cells were washed with PBS followed by PHEM buffer, then soluble tubulin dimers were extracted using 0.1% triton with 10µM taxol and 0.1% DMSO in PHEM buffer. Extracted cells were fixed with 2% PFA and 0.05% glutaraldehyde in PBS for 10 minutes, washed with

PBS and then reduced in 0.1% NaBH4 in PBS for 7 minutes at room temperature. Cells were then washed with PBS and blocked in 3% BSA and 0.2% Triton in PBS for 20 minutes at room

46 temperature, with agitation. Immunostaining was performed using primary antibodies directed

against: 6X-Histidine (Invitrogen, 4A12E4 37-2900; 1:500), total α-tubulin (Sigma, DM1A

T6199), Acetylated Tubulin (Sigma, T7451; 1:1,000), Tyrosinated Tubulin (Chemicon,

MAB1864; 1:1,000), Map1b (Santa Cruz Biotech, sc-135978; 1:500), Map2 (Novus Biologicals,

NB300-213; 1:2,000). Primary antibodies were diluted in blocking buffer and incubated

overnight at 4ºC in a humidified chamber. After primary antibody staining, cells were washed

three times with PBS. Fluorescently-conjugated secondary antibodies were diluted 1:500 in

blocking buffer and incubated for 2 hours at room temperature, protected from light. Secondary

antibodies were from Life Technologies (Carlsbad, CA) all used at 1:500. Alexa Fluor 568-

conjugated Phalloidin (Thermo, A12380; 1:20) was added during secondary antibody incubation

for labeling of filamentous actin. Tissues or cells of interest were mounted onto glass microscope

slides and sealed with glass coverslips and aqueous mounting media containing DAPI (Vector

Laboratories, #H-1200) and imaged on a Zeiss 780 or 880 confocal microscope with a 40X or

63X oil objective.

Microtubule Dynamics and Neuron Growth Rates

Primary neurons from wild-type and Tuba1aND/+ neonatal mouse cortices were cultured as described above. Prior to plating, mouse cortical neurons were nucleofected with 4µg each of

GFP-MACF43 and Myr-TdTomato plasmid DNA. Following 1 day in culture, neurons were transferred to a 37ºC imaging chamber and time-lapse images of GFP-MACF43 comets and

Myr-TdTomato membrane label were acquired using a Zeiss 780 confocal microscope with 40X oil objective. Following acquisition of baseline images. Images were acquired every 2 seconds for 2 minutes. A total of four time-lapse videos were acquired per neuron, with a 10-minute waiting period in between imaging sessions. Data from all four GFP-MACF43 time points were

47 pooled by cell to generate a cell average and then grouped by genotype for further analysis.

Membrane-bound Myr-TdTomato images were acquired at the beginning and end of the imaging

period and were used to track neuronal growth. Kymographs of GFP-MACF43 comets were

generated to assess microtubule polymerization rates in ImageJ/FIJI (National Institutes of

Health).

Western Blotting

Protein was isolated from brains of P0-P2 mice by dounce homogenization and ultra-

centrifugation. Tubulin-enriched protein fractions with MAPs were isolated according to a

previously established protocol [221]. Cos-7 cell protein was extracted using a Tris-Triton lysis

buffer with protease inhibitor cocktail (Sigma). Protein concentrations were assessed using a

BCA assay (Thermo), and relative concentration was determined using a Synergy H1 microplate

reader (BioTek Instruments, Winooski, VT). 5µg of either whole brain lysate or tubulin-enriched

protein fraction was loaded per lane and run on 4-20% Mini-Protean TGX Stain-Free precast

gels (4568093; Bio-Rad Laboratories, Hercules, CA) at 150mV for 1hr. Prior to protein transfer,

Stain-Free gels were activated with UV light for 1 minute and imaged for total protein on gel

using a ChemiDoc MP imager (Bio-Rad). Proteins were transferred to PVDF blotting

membranes (Bio-Rad) in standard 25mM Tris-base, 192mM glycine, 15% methanol transfer

buffer, or transfer buffer optimized for high molecular-weight proteins (>200kDa) by the

addition of 0.025% SDS. Blots were transferred at 4°C and 75V for either 1 hour for standard molecular-weight proteins, or 3 hours for high molecular-weight proteins. Immediately following transfer, PVDF membranes were rinsed in TBST and imaged to quantify the total protein on blot.

Gels were also imaged post-transfer to assess transfer efficiency for each blot. Membranes were blocked in Tris-buffered Saline containing 0.1% Tween-20 (TBST) with 5% BSA for 1 hour and

48 incubated in primary antibody diluted in TBST containing 1% BSA overnight at 4°C. Primary

antibodies were: mouse anti 6X-Histidine (Invitrogen, 4A12E4; 1:500), chicken anti-GFP

(Invitrogen, A10262; 1:1,000), and mouse anti-Map1b (Santa Cruz, sc-135978; 1:500). Blots

were incubated in HRP-conjugated Goat-anti-mouse (1:5,000; Santa Cruz) secondary antibody

diluted in TBST containing 0.5% BSA with streptavidin-HRP (Bio-Rad, 1:10,000) for 1 hour at

room temperature. Blots were developed in ECL solution for 5 minutes at room temperature

(Bio-Rad) and imaged.

Experimental design and statistical analyses.

Band volume of all Western blots was analyzed using Image Lab software (Bio-Rad).

Fluorescence intensity measurements, area and morphological assessment, kymograph

generation, and quantification of EM images was performed using ImageJ/FIJI software (NIH)

and Excel (Microsoft). Statistical analyses were performed, and graphs were created using Prism

version 8.0 (GraphPad). Most graphs display all data points to accurately represent the variability

in each dataset, except in cases where such presentation obscured visibility. For all statistical

analyses, statistical significance was considered to be p < 0.05. Statistical analyses used in each

experiment are indicated in their respective figure legends. For all graphs mean ± SEM was reported unless otherwise noted. Normality of each dataset was assessed using a Shapiro-Wilk test. In datasets with two groups, parametric data was analyzed using a Student’s t-test, while non-parametric data was assessed by Mann-Whitney U analysis of medians. Multiple groups

were compared by one-way or two-way ANOVA and analyzed post hoc by either a Bonferroni or Kruskal-Wallis test for parametric and non-parametric data, respectively.

49 Results

TUBA1A is required for formation of midline commissural structures

TUBA1A is a major component of developing neuronal microtubules, and is critical for

proper brain development [15]. Human TUBA1A-tubulinopathy patients and homozygous

TUBA1A mutant mouse models both exhibit severe brain malformations. Tuba1aND/+

heterozygous mutant mice undergo normal cortical migration, display comparable brain weight

to wild-type littermates at birth, and survive to adulthood [183]. However, Tuba1aND/+ brains exhibit agenesis of the corpus callosum and abnormal formation of the anterior and hippocampal commissures (Fig. 2.1A). In wild-type mice, nascent callosal ‘pioneer’ axons extend through midline at E15.5, and early ‘follower’ axons begin extending at E17 in mice [222]. Evidence of abnormal callosal projections were apparent as early as P0 in Tuba1aND/+ brains, as seen by the early formation of aberrant axon bundles adjacent to the callosum, known as Probst bundles (Fig.

2.1A)[223]. In addition to agenesis of the corpus callosum, adult Tuba1aND/+ corpus callosum

was found to be significantly thinner than wild-type (Fig. 2.1B). Similarly, adult Tuba1aND/+

anterior commissures are smaller than that of wild-type littermates (Fig. 2.1C). In wild-type

mice, corpus callosum thickness and anterior commissure area both increased significantly

between P0 and adulthood, however normal postnatal expansion of these tracts was not evident

in Tuba1aND/+ mice (Fig. 2.1B, C). Tuba1aND/+ axons fail to organize into typical midline

commissural structures, indicating that half of the normal complement of Tuba1a during brain

development is not sufficient for commissural axon guidance.

50

Figure 2.1 TUBA1A is required for formation of midline commissural structures. A. Luxol fast blue-stained coronal brain sections from postnatal day 0 (P0; top) and Adult (middle-bottom) wild-type and Tuba1aND/+ mice. Images portray abnormal midline commissural formation in Tuba1aND/+ mouse brains, with labels highlighting the corpus callosum (CC), anterior commissures (AC), and hippocampal commissure (HC). Scale bars are 500µm. B. Scatter plot representing corpus callosum width at P0, 3- months, and 10-months-old. Arrows indicate Probst bundles. Asterisk in A. shows where measurements for B. were obtained. C. Scatter plot displaying anterior commissure area in P0, 3-month, and 10-month-old wild-type and Tuba1aND/+ brains. Wild-type and Tuba1aND/+ measurements compared by two-way ANOVA. * p<0.05; ** p<0.01; *** p<0.001; and **** p<0.0001.

Examination of sagittal brain sections taken directly at midline revealed dramatic

disorganization of corpus callosum axons in Tuba1aND/+ brains (Fig. 2.2A). Compared to wild-

type, Tuba1aND/+ midline commissural axons were largely absent, and the existing axons failed to organize into a tract with uniform orientation (Fig. 2.2A). Despite dramatic differences between wild-type and Tuba1aND/+ callosal axon organization, Tuba1aND/+ axons were highly colocalized

with immunolabeled myelin sheaths (Fig. 2.2A). To further assess the impact of Tuba1aND/+

51 substitution on callosal axon morphology and myelination, we performed electron microscopy

(EM) in both wild-type and Tuba1aND/+ corpus callosi. Due to the scarcity of axons present

directly at midline in the Tuba1aND/+ corpus callosum, we sampled a region of corpus callosum

2mm lateral to midline for both wild-type and Tuba1aND/+ animals (Fig. 2.2B). EM images revealed a striking difference in axon density between wild-type and Tuba1aND/+ corpus callosi

(Fig. 2.2B, D; p=0.03). Myelin thickness, measured by G-ratio, was similar between wild-type

and Tuba1aND/+ brains (Fig. 2.2C; p=0.34), as was axon diameter (Fig. 2.2E; p=0.14). There was a trend towards decreased myelination in Tuba1aND/+ animals (p=0.07), however this difference was not statistically significant (Fig. 2.2F). These data provide evidence that Tuba1aND/+ callosal axons do not correctly organize to form a commissure. Previously published data indicated that reduced developmental Tuba1a function is tolerable for cortical neuron migration [183], however our results indicate that reduced Tuba1a is not sufficient to support axon pathfinding.

52

Figure 2.2 Axon density is reduced by Tuba1aND/+ in midline corpus callosum. A. Sagittal brain sections at midline from wild-type (top) and Tuba1aND/+ (bottom) brains stained with neurofilament-heavy (NFT-H; magenta) to label axons, and proteolipid protein (PLP; green) to label myelin. Images were taken at 4X (left) and 20X (right) magnification with 500µm and 50µm scale bars, respectively. B. Electron microscopy (EM) images of sagittal wild-type (top) and Tuba1aND/+ (bottom) sections of corpus callosum 2mm adjacent to midline. Enlarged regions on right are denoted by boxes, scale bar is 1µm. C. Scatter plot of G-ratio measurements for wild-type and Tuba1aND/+ axons. Data points represent individual myelinated axons and are clustered by animal (N=3 mice, n=100 axons; p=0.34). D. Scatter plot representing axon density of wild-type and Tuba1aND/+ axons in analyzed EM images. Data points represent average values for each animal (n=6 regions containing same total area; p=0.03). E. Scatter plot of axon diameters in wild-type and Tuba1aND/+ corpus callosum by EM. Only those axons captured in cross section were assessed for diameter, as skewed axons provide inaccurate measurements. Data points represent individual axons and are clustered by animal (n=100; p=0.14). F. Scatter plot representing the average percent of myelinated axons per animal in wild-type and Tuba1aND/+ corpus callosum. 6 regions containing the same total area were assessed (n=6; p=0.07). Statistical differences between means of wild-type and Tuba1aND/+ datasets were assessed by t test, with * p<0.05.

Tuba1aND a-tubulin does not incorporate into neuronal microtubules

Reduced developmental Tuba1a is not adequate to support long-distance axon targeting,

but the molecular and cellular mechanisms by which Tuba1a alters neuronal microtubule

function are unknown. The Tuba1aND allele was previously demonstrated to cause loss of

microtubule function in yeast and mice [183, 224]. However, the mechanism by which Tuba1aND substitution leads to loss-of-function (LOF) is unclear. The neuronal microtubule network is

53 complex, containing many different tubulin isotype proteins, PTMs, and binding of MAPs, all of which can modify functional microtubule properties. Further, the high degree of sequence similarity between a-tubulin isotypes has precluded study of individual tubulin isotypes at the protein level in vivo. No a-tubulin isotype or species-specific antibodies exist, and prior attempts to tag neuronal microtubules with N- or C-terminal fusion proteins have had detrimental effects on protein function [225]. These challenges have made the direct visualization of specific tubulin isotypes or mutant tubulin proteins in neurons historically difficult. Thus, the ways in which

TUBA1A specifically contributes to neuronal microtubule protein function have been difficult to ascertain. To address the need for better tools to study TUBA1A protein, we generated a functional hexahistidine (His6)-tagged TUBA1A construct based on a previously identified internal loop in the α-tubulin protein for in vivo visualization and manipulation of microtubules

[217] (Fig. 2.3A). We inserted a His6 tag into an internal loop of TUBA1A between residues I42 and G43. This region of α-tubulin was previously shown to tolerate addition of amino acids without disrupting tubulin function [217], and previous groups added a His6 tag in this loop to affinity purify tubulin subunits for reconstituted systems [216, 218]. However, to our knowledge this internal His6 tag on α-tubulin has never been used for immunohistochemical assays to visualize the microtubule network in vivo. Ectopically expressed wild-type TUBA1A-His6 is incorporated into Cos-7 cell microtubules (Fig. 2.3B). Compared to traditional immunolabeling of cellular microtubules (DM1A), TUBA1A-His6 provides excellent labeling of microtubule polymers (Fig. 2.3B). His6-tagged mutant TUBA1A can be ectopically expressed in cells to evaluate the abundance and polymerization capability of mutant TUBA1A proteins. We ectopically expressed three TUBA1A variants in Cos-7 cells, TUBA1AND, TUBA1ATE a known polymerization incompetent mutant [226], and TUBA1AEA a highly polymer-stable mutant

54 [227](Fig. 2.3B). As predicted, TUBA1ATE-His6 protein was not highly detected in Cos-7 cell

microtubule polymers, while TUBA1AEA-His6 protein was abundantly incorporated into cellular microtubules (Fig. 2.3B). In contrast to wild-type, ectopic TUBA1AND-His6 protein was not

detected in Cos-7 cells (Fig. 2.3B). Western blotting of lysates from Cos-7 cells, 48-hours post-

transfection revealed that TUBA1AND-His6 protein was significantly reduced compared to wild- type TUBA1A-His6 (Fig. 2.3C, D; p=0.04), despite similar TUBA1A mRNA levels between wild-type and TUBA1AND transfected Cos-7 cells (Fig. 2.3E; p=0.97). To evaluate if Tuba1aND mutant protein is targeted for proteasomal degradation, we treated TUBA1A-His6 transfected

Cos-7 cells with 5µM proteasome inhibitor, Lactacystin A (LactA [219, 220]), for 24-hours and probed for His6 abundance by western blot (Fig. 2.3C). Treatment with LactA significantly increased wild-type TUBA1A-His6 protein compared to control-treated cells, but had no effect on

TUBA1AND-His6 protein abundance (Fig. 2.3D; p=0.83). These results indicate that TUBA1AND mutant protein is likely not targeted for degradation by the proteasome, but may reduce cellular

TUBA1A by other mechanisms.

55

Figure 2.3 Tuba1aND impairs incorporation into cellular microtubules and reduces a-tubulin protein abundance. A. Schematic of Tuba1a-His6 tag and experimental design. B. Images showing Cos-7 cells transfected with wild-type, Tuba1aND, Tuba1aTE (polymerization-incompetent mutant), and Tuba1aEA (highly polymer stable mutant) Tuba1a-His6 plasmid DNA. Cells were immunolabeled with a-His6 antibodies to reveal microtubule incorporation of wild-type and mutant Tuba1a-His6 protein. Tuba1a-His6 images on the left are compared to traditional immunolabeling of a-tubulin using DM1A antibody in untransfected Cos-7 cells on right. C. Western blot for His6 in Tuba1a-His6 transfected Cos-7 cell lysates. A subset of transfected cells were treated with 5µM Lactacystin A (LactA) for 24 hours to block proteasomal degradation. His6 signal was normalized to GFP, which was expressed from the same plasmid. D. Quantification of band density for His6 western blot shown in C. His6 band density was normalized to GFP-expressing cells in control-treated (left), and cells treated with 5µM LactA for 24-hours (right). Data were analyzed by t test. N=3 transfections; n=3 technical replicates; p=0.04 for all comparisons marked with asterisks; p=0.83 for control vs LactA-treated Tuba1aND. E. Bar graph representing TUBA1A mRNA expression in Cos-7 cells transfected with Tuba1a-His6. TUBA1A mRNA expression was normalized to GFP mRNA expression. Data were normalized to TUBA1A expression in wild-type (WT) Tuba1a-His6-transfected cells and represent 3 separate transfection experiments and 3 qRT-PCR replicates. Differences between groups were assessed by t test (p=0.97). All images were taken at 40X magnification, scale bars are 10µm. All graphs show mean of data ± SEM, *p<0.05.

We next investigated if TUBA1AND substitution impairs incorporation of TUBA1A protein into neuronal microtubule polymers (Fig. 2.4). Wild-type rat cortical neurons were nucleofected with TUBA1A-GFP, wild-type TUBA1A-His6, TUBA1AND-His6 and TUBA1ATE-

His6 DNA at day in vitro 0 (DIV 0; Fig. 2.4A). Following 2-days in culture (DIV 2), cells were fixed and a subset of neurons were permeabilized to remove soluble tubulin dimers (“tubulin extraction”). Extraction of soluble tubulin leaves behind only polymer-bound tubulin, enabling visualization of polymerization competence of ectopic tubulin proteins [228]. Neurons expressing TUBA1A-GFP exhibited abundant GFP signal in intact cells, but tubulin extraction revealed that GFP fusion impaired incorporation of TUBA1A into neuronal microtubule polymers (Fig. 2.4B). In contrast, wild-type TUBA1A-His6 protein was highly present in both intact and tubulin extracted neurons, demonstrating that His6-tagging does not impair 56 polymerization ability of TUBA1A in neurons (Fig. 2.4C). As predicted, polymerization

incompetent TUBA1ATE-His6 mutant protein was diffusely visible in intact neurons, but was absent from tubulin extracted neurons (Fig. 2.4C). TUBA1AND-His6 protein was detectable at

very low levels in unextracted neurons, but was not visible following tubulin extraction,

indicating that TUBA1AND impairs incorporation into neuronal microtubules (Fig. 2.4C). These experiments illustrate that TUBA1AND reduces abundance of TUBA1A protein upstream of proteasomal degradation and prevents incorporation of mutant TUBA1A into cellular microtubules.

57

Figure 2.4 TUBA1AND a-tubulin does not incorporate into neuronal microtubule polymers. A. Schematic of cortical neuron isolation and transfection. B. Cortical rat neurons at DIV 2 transfected with TUBA1A-GFP. Top panel shows neurons with intracellular environment intact (unextracted), containing soluble tubulin dimers and polymerized microtubules transfected to express a TUBA1A-GFP fusion protein. Bottom panel shows neurons with soluble tubulin dimers extracted, showing only GFP-labeled TUBA1A that is incorporated into microtubule polymer. C. Rat cortical neurons at DIV 2 transfected with wild-type (WT) TUBA1A-His6, TUBA1AND-His6 LOF mutation, and TUBA1ATE-His6 polymerization-incompetent mutant as a negative control. Top panels show composite image containing membrane-bound GFP (green) for confirmation of transfection, a-His6 (Magenta) and DAPI (blue) immunolabeling. Middle panels show unextracted and bottom panels show tubulin extracted DIV 2 neurons (described in B.), labeled with a-His6 antibodies to visualize ectopic TUBA1A-His proteins. All images were taken at 63X magnification, scale bar is 25µm.

Reduced Tuba1a alters microtubule dynamic properties in neurons

Dynamic instability is a defining feature of microtubule polymers that is heavily

influenced by a number of factors to tune microtubule network function in cells [215]. MAP-

binding, PTMs, and incorporation of different tubulin isotypes can all influence microtubule

dynamics [12, 13, 60, 229]. Tuba1aND tubulin is less abundant than wild-type and does not incorporate into neuronal microtubule polymers (Figs. 2.3, 2.4). As decreased Tuba1a

58 incorporation was previously shown to accelerate microtubule polymerization in in vitro reconstituted microtubules [13], we next assessed microtubule dynamics in developing

Tuba1aND/+ neurons. Wild-type and Tuba1aND/+ cortical neurons were transfected with GFP-

MACF43, a fluorescently tagged protein containing the SxIP motif bound by microtubule plus-

end binding proteins (EBs), allowing for visualization of microtubule plus-end polymerization in

live cells (Fig. 2.5A)[230]. Kymograph plots generated from time-lapse GFP-MACF43 images

were analyzed for velocity, duration, and distance of GFP-MACF43 polymerization events (Fig.

2.5B). Previous analysis of Tuba1aND/+ GFP-MACF43 data revealed a significant reduction in the number of microtubule plus-ends per micron of neurite, which is evident in the example kymographs (Fig. 2.5B; [224]). Kymograph analysis showed that Tuba1aND/+ microtubules had accelerated polymerization velocity (19.67±1.1µm/min) compared to wild-type at DIV 1

(17.36±0.43 µm/min; Fig. 2.5C; n=688 events; p=0.0008 by Mann-Whitney test), consistent with prior data from S. cerevisiae [183]. Additionally, Tuba1aND/+ polymerization events covered a larger distance (Fig. 2.5D; n=688 events, p<0.0001) and lasted a longer amount of time (Fig.

2.5E; n=688 events, p<0.0001) than polymerization events in wild-type neurons. The increased polymerization capacity of Tuba1aND/+ neuronal microtubules was surprising, but collectively

with our previous results these data indicate that Tuba1aND protein is not functional, and that the

diminished a-tubulin pool leads to altered microtubule dynamics and changes to microtubule

organization in developing neurons.

59

Figure 2.5 Reduced Tuba1a alters neuronal microtubule polymerization. A. Schematic of nucleofection and imaging experimental setup. Neurons were nucleofected with GFP-MACF43 plasmid DNA and imaged at DIV 1 for microtubule polymerization events in an axonal region of interest. B. Representative images showing GFP-MACF43 expressing wild-type (WT) and Tuba1aND/+ neurons at 40X magnification (left) and representative kymographs that were generated from GFP-MACF43 images (right). Scale bars are 10µm and 5µm on neuron images and kymographs, respectively. C. Bar graph representing the average polymerization velocity for anterograde microtubule polymerization events in WT and Tuba1aND/+ cortical neurons at DIV 1 (n=688 events; p=0.0008 by Mann-Whitney test). D. Bar graph representing polymerization distance in DIV 1 WT and Tuba1aND/+ cortical neurons (n=688 events, p<0.0001). E. Bar graph representing polymerization duration in DIV 1 WT and Tuba1aND/+ cortical neurons (n=688 events, p<0.0001). Bars represent the mean of each group and error bars represent SEM. Differences between groups were assessed by t test, unless otherwise noted.

Tuba1a dictates neurite extension and cytoskeletal organization in growth cone

To assess potential mechanisms by which reduced Tuba1a prevents commissural neurons from reaching their contralateral targets, we measured neurite growth rates in cultured primary cortical neurons from P0 wild-type and Tuba1aND/+ mice (Fig. 2.6A). We labeled primary wild-

60 type and Tuba1aND/+ cortical neurons with a membrane-bound Myr-TdTomato to visualize growth rates in live neurons (Fig. 2.6A). Analysis of membrane bound Myr-TdTomato images taken one hour apart revealed that Tuba1aND/+ neurons grew at a significantly slower rate than wild-type neurons (Fig. 2.6B; N=3 mice, n=13 neurons; p=0.03 by Mann-Whitney test) with

Tuba1aND/+ neurons growing an average of 4.67±1.15µm/hr compared to 12.60±3.12µm/hr in

wild-type neurons. Additionally, measurements of the longest neurite, designated a putative

‘axon’, at DIV 3 revealed that Tuba1aND/+ neurites were significantly shorter than that of wild- type (Fig. 2.6C; N=3 mice, n=124 neurons; p=0.02).Together, these data show that developing neurons with reduced Tuba1a have shorter neurites and grow at a slower rate than wild-type.

61

Figure 2.6 Tuba1aND impairs axon outgrowth and alters growth cone cytoskeleton in cortical neurons. A. Schematic of experimental design and time-lapse images of membrane-labeled Myr-TdTomato neurons at DIV 1. Myr-TdTomato images were aquired 60 minutes apart to assess neuronal growth rate in wild-type (WT; top) and Tuba1aND/+ (bottom) cortical neurons. B. Scatter plot of growth rates in live cortical neurons from WT and Tuba1aND/+ mice at DIV 1. Data points represent individual neurons (N=3 mice, n=13 neurons; p=0.03 by Mann-Whitney test). C. Scatter plot of neurite length at DIV 3 in fixed WT and Tuba1aND/+ primary cortical neurons. For each cell, the longest neurite, designated a putative ‘axon’, was measured from the cell soma to the distal neurite tip using an acetylated tubulin marker. Data points represent individual neurons (N=3 mice, n=124 neurons; p=0.02). D. Images of DIV 3 WT (top) and Tuba1aND/+ (bottom) cortical neurons stained with antibodies directed against acetylated tubulin (green) and rhodamine phalloidin (F-actin, magenta). Composite images are shown (left) with single channel grayscale images for acetylated tubulin (middle) and F-actin (right). Scale bars are 10µm. E. Scatter plot of growth cone area at DIV 3 in WT and Tuba1aND/+ cortical neurons. Data points represent individual growth cones (N=3 mice, n=52 growth cones; p=0.26). F. Scatter plot of acetylated tubulin intensity within the growth cone of WT and Tuba1aND/+ cortical neurons at DIV 3. Data points represent individual growth cones (n=49 growth cones; p=0.89). G. F-actin intensity within growth cone of WT and Tuba1aND/+ cortical neurons at DIV 3. Data points represent individual growth cones (N=3 mice, n=49 growth cones; p=0.0014). H. Representative images of WT and Tuba1aND/+ cortical neuron growth cones showing distribution of acetylated tubulin (green) and F-actin (magenta). I. Line plot showing the average acetylated tubulin fluorescence intensity across a 20µm line scan through the central domain of the growth cone in DIV 3 WT and Tuba1aND/+ neurons. Differences between WT and Tuba1aND/+ growth cones were assessed by two-way ANOVA which showed a significant interaction between genotype and fluorescence intensity by position (n=20 growth cones; p<0.0001). J. Scatter plot representing the ratio of acetylated tubulin to F-actin intensity within the growth cones of DIV 3 WT and Tuba1aND/+ cortical neurons. Data points represent individual growth cones (N=3 mice, n=49 growth cones; p=0.0003 by Mann-Whitney test). For all plots, lines represent mean and error bars report SEM. Differences between WT and Tuba1aND/+ datasets were assessed by t test unless otherwise noted. *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.

To explore the precise mechanisms by which reduced Tuba1a contributes to slowed

outgrowth in Tuba1aND/+ neurons, we assessed the abundance of acetylated microtubules and filamentous actin (F-actin) in developing growth cones of wild-type and Tuba1aND/+ cortical neurons at DIV 3 (Fig. 2.6D). The growth cone is a dynamic developmental structure that uses the coordinated action of actin and microtubule cytoskeletons to drive neuronal outgrowth in

62 response to internal and external cues [1, 231]. Growth cone area was not changed in Tuba1aND/+

neurons (317.4±31.3µm2) compared to wild-type (380.9±30.4µm2; Fig. 2.6E; n=49 growth

cones; p=0.15). We examined the amount of acetylated tubulin, a PTM associated with stable

microtubules (Fig. 2.6F) and found there to be no difference in the overall fluorescence intensity

of acetylated tubulin in Tuba1aND/+ neurons compared to wild-type (n=49 growth cones; p=0.89).

In contrast, we observed a significant increase in F-actin intensity within the growth cones of

Tuba1aND/+ neurons compared to wild-type (Fig. 2.6G; n=49 growth cones; p=0.0014). Neuronal

microtubules splay out in the central, actin-dominated regions of the growth cone, but are

bundled towards the peripheral domains of the growth cone [162]. To assess the degree of

growth cone microtubule bundling, we next performed line scans across the widest point of DIV

3 growth cones ≥10µm (Fig. 2.6H). Line scans of acetylated tubulin through the growth cone

revealed differences in microtubule organization between wild-type and Tuba1aND/+ neurons

(Fig. 2.6I; n=39 growth cones; p<0.0001 between genotypes by two-way ANOVA). Specifically,

we observed peaks in fluorescence, indicating bundled microtubules, at the edges of the growth

cone in wild-type neurons, where acetylated tubulin was more diffuse and lacked obvious

organization in Tuba1aND/+ growth cones (Fig. 2.6I). Intriguingly, the ratio of acetylated

microtubules to F-actin in the growth cone was significantly reduced in Tuba1aND/+ neurons

compared to wild-type, indicating changes to the overall growth cone cytoskeletal environment

in Tuba1aND/+ neurons (Fig. 2.6J; n=49 growth cones; p=0.0003). Together, these data indicate that neurite growth rate is particularly sensitive to the amount of Tuba1a tubulin available, and impaired Tuba1a function leads to abnormal actin and microtubule architecture in the developing growth cone.

63

Figure 2.7 Neurons deficient in Tuba1a fail to localize Map1b to the developing growth cone. A. Western blots showing Map1b protein bound to tubulin in a tubulin polymer-enriched fraction from brain (top panel) and total Map1b protein in whole brain lysate (bottom panel) from wild-type (WT) and Tuba1aND/+ mice. Map1b bands are compared to presumed tubulin protein band (50kDa) from a stain-free gel. B. Scatter plot quantifying Map1b bound to tubulin in polymer-enriched tubulin fraction, normalized to the total protein present on blot using stain-free western blotting. p=0.03 C. Scatter plot representing total Map1b protein in brain lysate by western blot, normalized to the total protein on a stain-free blot. p=0.98 D. Scatter plot showing the subcellular distribution of Map1b protein in WT and Tuba1aND/+ cortical neurons at DIV 3. Data are represented as Map1b fluorescent signal in growth cone region divided by an axonal region proximal to the cell body. p=0.009. E. Representative images showing altered subcellular distribution of Map1b in Tuba1aND/+ (bottom) cortical neurons compared to WT (top) at DIV 3. Composite and individual channel grayscale images of MAP2 and Map1b immunocytochemistry are shown, i and ii indicate enlarged regions shown in insets. Scale bars are 10µm. Differences between groups were evaluated by t test. * p<0.05; ** p<0.01.

64 Tuba1aND neurons fail to localize critical developmental proteins to growth cone

MAPs play a crucial role in regulating neuronal microtubule function to support proper

neurodevelopment. Microtubule-associated protein 1b (Map1b) promotes axon extension and is

required for formation of the corpus callosum in mice [70, 71]. There was no significant deficit

in the association of Map1b with microtubules from Tuba1aND/+ brain lysates compared to wild- type, in fact Tuba1aND/+ lysates bound slightly more Map1b than wild-type (Fig. 2.7A, B;

p=0.03). Western blot analysis of whole brain lysates from wild-type and Tuba1aND/+ mouse

brains showed no difference in the total amount of Map1b protein (Fig. 2.7A,C; p=0.98). These

data indicate Tuba1aND does not impair Map1b’s interaction with neuronal microtubules. In

developing neurons, Map1b localizes strongly to the growth cone to promote axon growth and

facilitate microtubule response to guidance cues [56, 67, 70, 232, 233]. Tuba1aND/+ neurons contained Map1b protein, but exhibited very little Map1b fluorescence in the growth cone, compared to wild-type neurons (Fig. 2.7D, E; n=31 growth cones; p=0.009). These data provide evidence that while the abundance of Map1b protein is unchanged by Tuba1aND , reduced

Tuba1a does not allow for correct localization of Map1b to the growth cone. Failure of

Tuba1aND/+ neurons to localize Map1b to the developing growth cone provides a putative

mechanism by which developing axons may fail to respond to critical guidance cues, leading to

the commissural deficits observed in Tuba1aND/+ mice.

65

Figure 2.8 Mechanisms of Tuba1a-induced axonal pathfinding deficits. A. Schematic illustrating how reduced Tuba1a may impair ability of Tuba1aND/+ axons to reach key signaling points at the correct developmental time to support proper brain formation. Wild-type axons (WT; top) reaching the midline crossing point of corpus callosum at embryonic day (E) 18.5 are compared to the potentially stunted axonal growth in Tuba1aND/+ brains (bottom). The reduced density of microtubules and altered polymerization dynamics likely contribute to slower growth of developing Tuba1aND/+ commissural neurons. B. Schematics illustrating potential molecular mechanisms by which Tuba1a supports navigating axons. i. Inset from (A), showing WT growth cones navigating through midline corpus callosum. WT growth cones are rich with Map1b, which binds growth cone microtubules to guide cellular response to guidance molecules like Netrin. ii. Inset from (A) showing Tuba1aND/+ growth cones, which fail to localize Map1b and may be rendered unable to mount a cytoskeletal response to extracellular cues.

66 Multi-faceted functions of TUBA1A during neurodevelopment

Our results support two potential models by which TUBA1A regulates neuronal

microtubules to promote commissural axon pathfinding. Using our TUBA1A-His6 tool, we

illustrated that the Tuba1aND allele diminishes Tuba1a protein abundance in cells (Fig. 2.3).

Here, we demonstrate that diminished Tuba1a alters microtubule dynamics and impairs neuron growth in vitro (Figs. 2.5, 2.6). Thus, reduced TUBA1A could render neurons incapable of supplying the microtubule bulk needed to drive axon outgrowth forward at a specific rate (Fig.

2.8A). The timing of neurodevelopment is precisely regulated, and neurons which fail to reach targets at the correct time can miss crucial developmental signaling events. We additionally show that neurons with reduced Tuba1a fail to localize a critical developmental MAP, Map1b, to the growth cone (Fig. 2.7). Interactions between MAPs and microtubules play a major role in adapting microtubule function in response to a changing intra- and extra-cellular developmental environment [68, 71, 233]. Therefore, neurons with reduced TUBA1A may be rendered unable to respond to extracellular guidance cues, such as Netrin1, due to failed sub-cellular localization of MAPs and other critical cargoes (Fig. 2.8B). Our evidence supports a multi-faceted role for

TUBA1A during neurodevelopment, where it tunes microtubule dynamics and density to fuel growth and also provides stable tracks for rapid intracellular transport.

Discussion

Studying a-tubulin isotypes in vivo

TUBA1A has long been associated with neurodevelopment due to its spatial and temporal

expression as well as its role in tubulinopathy [14-16, 18-22, 27, 28], but it has been historically

difficult to study the contribution of a single a-tubulin isotype to microtubule network function

67 in vivo due to the limited availability of isotype-specific tools. Here, we present a novel tool for

studying TUBA1A protein in vivo without impacting native microtubule properties by

introducing a hexahistidine (His6) tag into a previously identified internal loop of TUBA1A (Fig.

2.3) [216-218]. In this chapter, we provide the first evidence that TUBA1A microtubules are

essential for regulating neuronal microtubule function to support long-distance axon targeting.

Ectopic expression of TUBA1A-His6 protein in cells revealed that Tuba1aND protein is approximately half as abundant as wild-type (Fig. 2.3D), despite similar amounts of mRNA expression in transfected cells (Fig. 2.3E). Based on this evidence, Tuba1aND substitution causes

targeted degradation of the mutant Tuba1a protein, but our results suggest that degradation by

the proteasome is unlikely (Fig. 2.3D). Newly synthesized a- and b-tubulin proteins enter a

complex tubulin folding pathway, where they interact with cytosolic chaperonins and tubulin-

binding cofactors to become folded and assembled into tubulin heterodimers [234]. As Tuba1aND

protein is diminished compared to wild-type, but is not proteasomally degraded, we predict that

mutant Tuba1aND protein may ineffectively cycle through the tubulin folding pathway. Further testing will be needed to evaluate the precise molecular mechanisms of reduced Tuba1aND protein, but it is evident that Tuba1a function is lost in the Tuba1aND model.

Neurons with reduced Tuba1a function had accelerated polymerization compared to wild- type, but exhibited deficits in neurite extension likely due to the decreased axonal microtubule density (Figs. 2.5, 2.6; [224]). Additionally, Tuba1a-deficient microtubules were not adequate to support growth cone localization of at least one critical developmental MAP (Fig. 2.7). We show that Tuba1a-rich microtubules promote axon outgrowth and pathfinding, and reduced Tuba1a was not sufficient to form forebrain commissures in Tuba1aND/+ mice (Figs. 2.1, 2.2).

Collectively, these data support the conclusion that reduced TUBA1A during neurodevelopment

68 is adequate for cortical neuron migration [183], but does not allow for sufficient microtubule

tracks to properly localize proteins to the growth cone for axon guidance. Thus, long range axon

guidance may be exquisitely sensitive to a-tubulin levels and microtubule structure.

Studying individual a-tubulin isotypes in neurons has been arduous as the high degree of

amino acid sequence similarity between a-tubulin isotypes prohibits generation of species-

specific antibodies and has made genetically targeting a single a-tubulin gene challenging. The abundance of clinically identified mutations to TUBA1A provide strong evidence that TUBA1A is a major player in both tubulinopathy and typical neurodevelopment, however the lack of available tools to study TUBA1A in vivo has prevented researchers from understanding precisely how TUBA1A contributes to neurodevelopment. As such, previous studies of tubulinopathy mutations have relied heavily on mRNA analysis and indirect methods of evaluating TUBA1A function. Here we introduce an important advancement in the study of TUBA1A protein in vivo,

by harnessing a previously-identified internal loop within TUBA1A that tolerates addition of

small epitope tags without impacting TUBA1A incorporation or dynamics [217]. Our internal

His6 TUBA1A construct marks an important advancement for the study of tubulinopathy and

neuronal a-tubulin as a whole. TUBA1A-His6 was readily expressed in both Cos-7 cells and neurons and was able to incorporate into microtubule polymers, unlike TUBA1A containing a

GFP fusion that prohibited incorporation into neuronal microtubules (Fig. 2.4). Additionally, we successfully used this epitope-tagged TUBA1A to model mutant tubulin behavior in vitro (Figs.

2.3, 2.4). Overall, this tool provides an important advancement in the study of a-tubulin protein in vivo, and makes interrogating the function of specific a-tubulin isotypes accessible to more researchers.

69 TUBA1A influences microtubule density, dynamics, and function in cells

The microtubule cytoskeleton supports a wide range of different cellular functions in

different cell types, ranging from facilitating chromosome segregation during mitosis to forming

dynamic and motile structures like the neuronal growth cone. Understanding how different cells

use the same basic building blocks to create vastly different microtubule-based structures is a

major question in microtubule biology. To date, several mechanisms have been identified

through which cells can regulate microtubule network properties and overall function. We show

that a mutation which reduced Tuba1a incorporation into cellular microtubules accelerates

microtubule polymerization rates (Fig. 2.5), consistent with what was previously reported for the

analogous mutation in yeast, Tub1ND [183]. This result is also supported by recent evidence showing that increased incorporation of Tuba1a tubulin slowed microtubule polymerization rates in in vitro reconstituted tubulin [13]. These results are fitting with the “tubulin code” model, which proposes that incorporation of different tubulin isotypes can modify microtubule network behavior [215, 235]. Importantly, previous work has shown that local changes to growth cone microtubule dynamics facilitate growth cone turning in response to extracellular cues [162, 212,

214, 231, 236, 237]. Thus, dysregulation of growth cone microtubule dynamics, as was observed in Tuba1aND/+ cortical neurons, could diminish the ability of developing neurons to appropriately interact with their environment. Together, these data support the conclusion that incorporation of

Tuba1a a-tubulin tunes neuronal microtubule polymerization rates to support neurodevelopmental processes.

While the abundance of acetylated tubulin was not significantly different between

Tuba1aND/+ and wild-type growth cones, the distribution of acetylated microtubules was different

by genotype (Fig. 2.6). Differences in distribution of acetylated tubulin within the growth cone

70 likely reflects altered microtubule organization, as we did not detect a change in the overall

abundance of growth cone acetylated microtubules by genotype (Fig. 2.6). Acetylation is a

microtubule PTM that is associated with stable microtubule populations and as such is sparse in

dynamic structures like growth cones [81, 193, 238-241]. However, microtubule acetylation can

be induced in growth cones following contact with extracellular matrix proteins and was shown

to promote cortical neuron migration in vivo and suppress axon branching in vitro, demonstrating

a clear role for this PTM in development [242-244]. Tubulin PTMs, like acetylation, have been

shown to impact MAP-binding affinity and function, providing a clear mechanism by which

changing the PTM landscape of microtubules could alter neuronal microtubule function [12, 96,

106, 235, 241, 245, 246]. Thus, any changes to the organization or distribution of acetylated

microtubules in the growth cone could impact the ability of developing neurons to appropriately

navigate their environment and establish correct synaptic targets.

Tuba1aND/+ growth cones showed a significant increase in F-actin signal compared to wild-type, causing an overall shift in the growth cone microtubule-actin balance (Fig. 2.6). It is

well established that interplay between the actin and microtubule cytoskeleton drives growth

cone movements in developing neurons [1, 2, 231, 247]. Growth cone microtubule

polymerization has been shown to induce F-actin assembly, and coordination of actin and

microtubules is regulated by interactions with MAPs to drive appropriate growth cone response

[248-251]. As actin and microtubules are tightly regulated within the growth cone, it is

reasonable to assume that mutations which disrupt microtubule function, like Tuba1aND, likely also impact the actin cytoskeleton of developing neurons. In Tuba1aND/+ neurons, the actin

cytoskeleton may occupy increased growth cone territory as potential compensation for

dysfunctional microtubules. Further, we showed that Tuba1aND/+ neurons do not effectively

71 localize at least one developmental MAP, Map1b, to the growth cone (Fig. 2.7). Map1b acts downstream of several important developmental signaling pathways to regulate function of both actin and microtubules within the growth cone, and dysregulation of this or other MAPs could therefore impact multiple cytoskeletal components [232, 233, 252]. The mechanisms by which

Tuba1aND induces changes to the growth cone actin cytoskeleton remain to be explored, but could reveal important insights on how microtubules and actin are coordinately regulated to support growth cone navigation.

Models of Tuba1a-dysfunction reveal potential roles in developmental signaling

Tuba1aND/+ neuronal microtubules were not sufficient to support growth cone localization of Map1b (Fig. 2.7). Microtubules are the tracks upon which intracellular cargo transport occurs in neurons. Here we present evidence that impaired Tuba1a function in neurons causes aberrant localization of Map1b (Fig. 2.7). Map1b mRNA is a known target of the mRNA transport protein

FMRP and is locally translated within developing neurons [253, 254]. As we previously demonstrated that intracellular transport is impaired in developing Tuba1aND/+ neurons [224], this provides a putative model by which reduced Tuba1a could lead to altered localization of developmental MAPs. Intracellular transport is a crucial function of neuronal microtubules throughout life, however microtubule-based transport has been shown to be essential during neurodevelopment [69, 213, 255-259]. Correct localization of developmental MAPs, mRNAs and organelles are crucial for cytoskeletal response to extracellular guidance cues [69, 258, 260-

262]. In particular, Map1b is required for neuronal response to the guidance cue, Netrin1, a key player in commissural formation [263-268]. We showed that neuronal microtubules with reduced

Tuba1a do not support neuronal growth to the same degree as wild-type microtubules, causing

72 shorter neurite length and slower growth rates in vitro (Fig. 2.6). The timing of developmental processes is crucial for effective signal transduction and supports the formation of appropriate synaptic contacts [269, 270]. Collectively, the data presented in this chapter support two potential models by which Tuba1aND neuronal microtubules fail to support proper neurodevelopment. The first model posits that neurons lacking functional TUBA1A do not have adequate intracellular transport to support localization of critical developmental proteins (Fig.

2.8). Inappropriate protein localization during critical points in axon extension and guidance could render neurons unable to respond to incoming guidance cues, as the machinery required to induce microtubule response to extracellular cues is absent. The second model proposes that the timing of axon extension during development is crucial for effective axon guidance. If neurons lacking TUBA1A are not reaching the correct location at the correct time, it is possible that neurons will fail to receive key developmental signals (Fig. 2.8). Importantly, these models are not mutually exclusive, as we demonstrated that TUBA1A is crucial for both developmental protein localization and neuron outgrowth. The extent to which these processes contribute to the overall deficit in commissural axon guidance remains to be explored in future studies.

Understanding the mechanisms by which microtubules contribute to discrete aspects of neurodevelopment is an active area of research. Human neurodevelopmental disorders that impact microtubule function, such as tubulinopathy, demonstrate that microtubules are critical for proper neurodevelopment to occur. Tubulinopathy patients exhibit severe, often lethal, brain malformations that frequently impact multiple neurodevelopmental processes, including neuronal survival, migration and axon extension [19, 20, 23-26, 142, 271]. The range of phenotypes exhibited by tubulinopathy patients have made it challenging for scientists to pinpoint specific aspects of neuronal function that are reliant on TUBA1A tubulin. In this way,

73 mutations such as the Tuba1aND variant whose severity can be tuned according to gene dosage can be used as important tools to interrogate the requirement for Tuba1a in discrete aspects of neurodevelopment. Importantly, though tubulinopathy patients exhibit a range of brain phenotypes, commissural abnormalities such as agenesis of the corpus callosum, are one of the most commonly reported features of this disease [15, 17-19, 22]. As cortical malformations and neuronal migration errors are also common features of tubulinopathy, it has thus far been unclear as to whether commissural deficits occur as a primary or secondary consequence of TUBA1A dysfunction. In this study, we provide evidence that neurons deficient in Tuba1a fail to properly navigate to meet contralateral binding partners. These data demonstrate that TUBA1A is required for forebrain commissural formation, independent of its role in neuronal survival or migration.

The exciting insights presented in this chapter expand upon the currently known role for

TUBA1A in neurodevelopment, and advance the study of tubulinopathy by presenting specific mechanisms by which TUBA1A microtubules support neurodevelopment.

74 CHAPTER III

REDUCED TUBA1A TUBULIN CAUSES DEFECTS IN TRAFFICKING AND

IMPAIRED ADULT MOTOR BEHAVIOR2

Introduction

Microtubules are the tracks upon which proteins, RNA, and organelles are trafficked.

Microtubule polymers are composed of a/b-tubulin heterodimers. In humans, nine a- and ten b-

tubulin isotypes are differentially expressed and could influence properties of specific cellular

microtubule networks [7, 8, 215]. In developing neurons, the most highly expressed a-tubulin is

encoded by Tuba1a [14, 16, 28, 125, 126]. Heterozygous mutations that disrupt human TUBA1A are associated with severe brain malformations, termed tubulinopathies, suggesting that TUBA1A is critical for brain development [15, 17, 20, 180]. The most common human TUBA1A variants dominantly disrupt microtubule function in neurons [23, 25]. No deletions, frameshift, or nonsense mutations have been identified in association with TUBA1A-tubulinopathy patients

[17-19]. We recently proposed TUBA1A mutations that impair brain development dominantly disrupt microtubule function, while heterozygous TUBA1A loss-of-function mutations are tolerable for brain development and are not clinically reported [210]. How TUBA1A contributes to adult neuron function has not been reported.

TUBA1A constitutes approximately 95% of neuronal a-tubulin mRNA developmentally

[16]. While Tuba1a expression is reduced overall in the adult brain, some regions including the hippocampus, cerebellum, basal ganglia, and certain regions of the cerebral cortex retain Tuba1a

expression in adulthood [27]. Over time, neuronal microtubule networks acquire PTMs and

2 Portions of this chapter are published with permission from our previously published article: Buscaglia, G., et al., Reduced TUBA1A tubulin causes defects in trafficking and impaired adult motor behavior. eNeuro, 2020 [216].

75 associations with MAPs that stabilize neuronal microtubules [73, 94, 102, 103, 246, 272].

TUBA1A proteins may incorporate into microtubules during neurodevelopment and remain

within the microtubule network of adult neurons. Indeed, Tuba1a, and other tubulins, have a

half-life on the order of weeks to months in rodents [127]. TUBA1A may be important in the

adult neuronal microtubule network. However, mechanistic understanding of TUBA1A function

in vivo has been limited by the high degree of sequence similarity between a-tubulin isotypes, as isotype-specific a-tubulin antibodies are not available.

To overcome the technical limitations to studying the specific impact of Tuba1a in adult

neurons, we use the Tuba1aND mutant mouse which harbors an asparagine to aspartic acid substitution at amino acid residue 102 [183]. Homozygous Tuba1aND/ND mice are neonatal lethal

and have severe brain malformations consistent with homozygous Tuba1anull and Tuba1a-R215*

mutant mice [182] and similar to phenotypes observed in human patients with heterozygous

TUBA1A mutations [15, 183]. The corresponding ND substitution in the primary a-tubulin in

yeast decreased a-tubulin protein levels, altered microtubule dynamics, reduced incorporation of

the mutant a-tubulin into the lattice, and impaired mitotic division [183]. My results from

Chapter II indicate that the Tuba1aND allele reduced Tuba1a protein and impaired incorporation

into cellular microtubules. Together, these data demonstrate that Tuba1aND substitution results in

less stable microtubules. In contrast to the severe impact of Tuba1aND/ND homozygous substitution on neuronal development, heterozygous Tuba1aND/+ mice survive to adulthood

[183]. The Tuba1aND/+ heterozygous mice can be used to interrogate the requirement for Tuba1a neuronal microtubules within the mature brain.

Here, we show that reduced function of Tuba1a, a tubulin isotype that has historically been characterized only in a developmental context, impacts the ability to maintain synapses

76 over time and ultimately results in an adult-onset movement disorder. Microtubule tracks are not

adequately assembled when there is a deficit in Tuba1a tubulin early in development. Insufficient

microtubule tracks result in organelle trafficking deficits that are apparent in Tuba1aND/+

heterozygous neurons early in development. Despite microtubule track abnormalities and

resulting changes in trafficking, Tuba1aND/+ microtubules are sufficient to support survival and

morphologically normal neuromuscular junction synapses. However, neuromuscular junction

synapse morphology and animal behavior deteriorate in an age-related manner in Tuba1aND/+

animals, without evidence of neuronal cell death or degeneration. Together these data indicate

that both developing and adult neurons require functional TUBA1A, and suggest that the

‘developmental’ tubulin TUBA1A is required for mature neuronal function.

Materials and Methods

Mice

All animal research was performed in accordance with the Institutional Animal Care and Use

Committee at the University of Colorado School of Medicine. The Tuba1aND mouse model was generated via an ENU-induced mutagenesis forward genetic screen and is described in more detail in Hanson et al. (2016). All mice used were maintained on a 129S1/C57Bl6 genetic background. Mice were kept on a 12:12 light:dark cycle with ad libitum access to food and water. Tuba1aND and wild-type littermate mice were maintained on water supplemented with

ND/+ 0.2g/L MgSO4 to promote Tuba1a survival and ability to reproduce. Male and female mice were represented in all studies. All mice were genotyped by PCR amplification of tail DNA followed by Sanger sequencing to differentiate homozygous or heterozygous Tuba1aND/+ mice

from wild-type.

77 Behavioral Phenotyping

Mice were assessed for abnormal motor function between 2- and 10-months of age. Mice were allowed to acclimate to the testing room for 30 minutes prior to behavioral assessment. Gait analysis testing was performed in an enclosed raised chamber with a strip of paper lining the bottom. Fore and hind paws were painted differing colors just prior to testing, after which mice were placed at the chamber entrance and allowed to run through. Distances between footprints taken from a running gait were analyzed in ImageJ (ImageJ, National Institutes of Health).

Rotarod performance was assessed using a Rota-Rod Treadmill (Med Associates, Inc, Fairfax,

VT) apparatus with one mouse per lane and a maximum of four mice tested simultaneously.

Mice received no prior training on the rotarod task. The rod initially began rotating at 3 RPM and increased until reaching a terminal rotational speed of 30 RPM after 3 minutes. The time until each mouse fell off or lost its grip such that it could not remain upright on the rod was measured.

A cutoff time of 4 minutes was established for any mice that remained on the rod after peak rotational speed was reached. Mice performed three subsequent trials on the rotarod per testing session, with a 15-minute rest period in between trials. Grip strength was tested in mouse forelimbs using a grip strength meter (Stoelting Co., Wood Dale, IL). Three subsequent forelimb grip strengths were recorded per testing session.

Mouse hind-paw sensory function was tested between 3- and 6-months. For both Von

Frey and Hargreaves analysis individual mice were contained in 8cm x 8cm x 17cm chambers and were allowed to acclimate to the testing chamber for 30 minutes. Mice were only tested when resting on all four paws in a non-grooming awake state. Von Frey analysis of mechanical stimulus sensation was performed in left and right hind-paws using Touch Test Sensory Probes

(Stoelting Co.). Von Frey performance score was analyzed using the SUDO method established

78 in [273]. Hargreaves analysis of thermal pain sensation was assessed using a paw thermal

stimulator instrument (University of California, San Diego). Time until withdrawal from thermal

stimuli was recorded for each animal in both left and right hind paws. Three subsequent trials

were performed for each hind paw per testing session.

Histology

Mice were anesthetized and trans-cardially perfused with 0.1M NaCl and 4% paraformaldehyde

(PFA) for histology. Tissues of interest were dissected and post-fixed in 4% PFA. Serial tissue

sectioning was performed on a CM1520 cryostat (Leica, Wetzlar, Germany) and 30µm serial cryosections were obtained for all histology experiments. For all histological analyses, three regions were chosen for quantification which spanned the anterior-posterior axis, and the same regions were assessed for each animal. For Nissl staining, sections from the lumbar spinal cord were stained using 0.1% cresyl violet. For immunofluorescence studies PFA-fixed tissues were blocked in phosphate-buffered saline (PBS) containing 5% bovine serum albumin (BSA) with

0.3% Triton-X 100. Primary and secondary antibodies were diluted in PBS containing 1% BSA with 0.1% Triton-X 100. For a-Bungarotoxin labeling of neuromuscular junction synapses,

PFA-fixed extensor digitorum longus, flexor digitorum brevis, and soleus muscles were dissected from the hindlimbs of postnatal day 30 (P30) and adult mice. Muscle fibers were teased apart using forceps and mounted onto positively charged glass microscope slides. Muscle fibers were blocked in a blocking buffer containing 3% BSA, 5% goat serum and 0.5% Triton-X 100 then stained with primary antibodies diluted in blocking buffer at 4°C overnight. Secondary antibodies and rhodamine conjugated a-Bungarotoxin (a generous gift from Dr. John Caldwell) were added for two hours at room temperature. Primary antibodies were as follows: Mouse anti-

Calbindin D-28k (Swant AgCB10abs; 1:250), Rabbit anti-ER81 (BioLegend 840401; 1:5,000),

79 Mouse anti SMI32 (Biolegend 801702; 1:5,000), Rabbit anti-Synaptophysin (Invitrogen MA5-

14532; 1:200). Fluorescently conjugated secondary antibodies were Life Technologies

(Carlsbad, CA) all used at 1:500. For neuromuscular junction analysis, Z-stack images were obtained at 63X magnification with a slice size of 0.5µm with 10 total slices covering 4.5µm total z-distance.

Electron Microscopy

Mice used for electron microscopy were perfused with 0.1M NaCl and 2.5% glutaraldehyde 4%

PFA, after which the spinal column was dissected and post-fixed in 2.5% glutaraldehyde 4%

PFA overnight at 4°C. Following post-fixation spinal cords were dissected from the spinal column, and a 2mm region of the cervical spinal cord between C2-C4 was dissected and sent for further processing and imaging by the CU School of Medicine Electron Microscopy Core facility. Myelin thickness, axon density, and axon diameter were measured from electron micrographs in the lumbar spinal cord. G-ratio was calculated using the formula G = ri/Ri where

ri = radius of axon and Ri = radius of axon and myelin. At least three sections per animal were

analyzed, measuring 100 axons per animal. The same total area was quantified between wild-

type and Tuba1aND/+ mice.

Cell Culture and Transfection

Dissociated neurons were cultured from P0-P2 mouse cortices. Brains were removed and placed into HBSS (Life Technologies) supplemented with 1M HEPES (Life Technologies) and 1mM kynurenic acid (Tocris Bioscience, Bristol, UK). Meninges were removed and cortices were dissected and cut into approximately 1mm pieces. Cortical pieces were triturated to a single-cell suspension using glass Pasteur pipettes. Cortical neurons were plated onto 35mm Poly-D-Lysine coated glass-bottom culture dishes at a density of 350,000 cells/35mm. Neurons were maintained

80 in a 37°C humidified incubator with 5% CO2 in phenol-free Neurobasal-A medium (Life

Technologies) supplemented with B-27 (Thermo), Penn/strep (Thermo), GlutaMax (Thermo),

5ng/mL b-FGF (Gibco), and Sodium Pyruvate (Thermo). For GFP-MACF43 puncta analysis,

neurons were nucleofected with 2µg of plasmid DNA using a mouse neuron nucleofector kit

(Lonza, Cologne, Germany) and Nucleofector™ 2b Device (Lonza), then plated normally.

GFP-MACF43 and Organelle Transport Assays

Primary cortical neurons nucleofected with GFP-MACF43 plasmid DNA were imaged at DIV1

to estimate the number of microtubule plus-ends per µm of axon. Cells were placed in a 37°C

imaging chamber and imaged using a 40X oil objective on a Zeiss 780 confocal microscope

using ZEN imaging software (Zeiss). GFP-MACF43 images were acquired every 2 seconds for a

total duration of two minutes. Multiple two-minute videos were acquired for each neuron

imaged, and data were used to generate a cell average, which was reported. Lysosomal and

Mitochondrial trafficking was assessed in cortical neurons at day in vitro 7 (DIV 7). Cortical

neurons were incubated in either LysoTracker Red DND-99 or MitoTracker Red FM

(Invitrogen) dye for 20 minutes at 37°C after which normal culture media was replaced and cells were transferred to a 37°C imaging chamber and imaged on either a Zeiss 780 confocal

(MitoTracker) or a Nikon Ti-E microscope equipped with a 1.3 NA 40× CFI60 Plan Fluor objective, piezo electric stage (Physik Instrumente; Auburn, MA), spinning disk confocal scanner unit (CSU10; Yokogawa), 488-nm and 561-nm lasers (Agilent Technologies; Santa

Clara, CA), and an EMCCD camera (iXon Ultra 897; Andor Technology; Belfast, UK) using

NIS Elements software (Nikon). Cells were imaged every 1 second or every 2 seconds for lysosomes and mitochondria, respectively.

81 Western Blotting

Protein was isolated from brains of P0-P2 and adult mice by dounce homogenization and ultra- centrifugation, using a modified version of the protocol described by Vallee et al. [221]. Protein concentrations were assessed using a BCA assay (Thermo), and relative concentration was determined using a Synergy H1 microplate reader (BioTek Instruments, Winooski, VT). For tubulin western blotting 0.25µg of brain lysate was loaded per lane, for all other blots 10µg of tissue lysate was used. Protein was run on 4-20% gradient acrylamide gels (Bio-Rad

Laboratories, Hercules, CA) at 150mV for 1hr. Proteins were transferred to PVDF blotting membranes (Bio-Rad) in transfer buffer containing 25mM Tris-base, 192mM glycine and 15% methanol for 1 hour at 75V at 4°C. Gels were dyed with Coomassie Blue (Bio-Rad) post-transfer to assess transfer efficiency for each blot. Membranes were blocked in Tris-buffered Saline containing 0.1% Tween-20 (TBST) with 5% BSA for 1 hour and incubated in primary antibody overnight at 4°C. Primary antibodies were diluted in TBST containing 1% BSA at 4°C overnight. Blots were incubated in secondary antibodies diluted in TBST containing 0.5% BSA with streptavidin-HRP (Bio-Rad, 1:10,000) for 1 hour at room temperature. Primary antibodies for western blotting were as follows: Mouse anti-DM1A (Sigma-Aldrich T6199; 1:10,000),

Rabbit anti-GAPDH (Cell Signaling 14C10; 1:1,000), Mouse anti-Acetylated tubulin (Sigma-

Aldrich T7451; 1:1,000), Mouse anti Tyrosinated tubulin (Sigma T9028; 1:5,000), Rabbit anti

Detyrosinated tubulin (Abcam ab48389; 1:1,000), Mouse anti Polyglutamylated tubulin (GT335,

Adipogen AG-20B-0020; 1:2,000), Mouse anti SMI32 (Biolegend 801702; 1:500). Secondary antibodies used were HRP-conjugated and diluted 1:5,000 (Santa Cruz Biotechnology, Dallas,

TX). Blots were developed in ECL solution (Bio-Rad) and imaged using a ChemiDoc MP imager (Bio-Rad).

82 RNA isolation + RTPCR

RNA was isolated from flash-frozen mouse brains dissected from P0 or adult mice using an SV

Total RNA Isolation system (Promega, Madison, WI). RNA concentration and purity were determined using a spectrophotometer, then cDNA was synthesized using the RT2 First Strand

Kit (Qiagen, Hilden, Germany). RT-PCR reactions were prepared with SYBR Green RT-PCR

Master mix (Thermo) and run with a CFX Connect Real-Time System (Bio-Rad). Samples were run in triplicate, results were analyzed in Excel. All qPCR data presented in this chapter was normalized to expression of the housekeeping gene Cyclin A. Tuba1aND/+ relative mRNA quantity was normalized to wild-type in cases where mRNA expression was compared by genotype. In cases where tubulin genes were compared to one another, we calculated relative quantity of mRNA (RQ) using the equation RQ = E ^-∆CT, where E is the primer efficiency, calculated from running a dilution curve of each primer set. For all qRT-PCR experiments 3 biological replicates were used per genotype. Primers were as follows: TUBA1A

(F:ATTATGAGGAGGTTGGTGTG, R:TGTTGGAACACAATAAACATC);

TUBA1B(F:CTCATTGCGTTACTTACCTC, R:TCACGCATGATAGCAACG);

TUBA1C(F:GGTATATAAGCCCTGTCCTG, R:CTCACGCATATTTAGTCCTTG);

TUBA4A(F:ACTGTAATCGATGAGATCCG, R:AATACTAGGAAGCCCTGAAG).

Experimental design and statistical analyses.

All experiments utilized experimental design statistics for random data. Band volume of all

Western blots was analyzed using Image Lab software (Bio-Rad). Organelle transport and GFP-

MACF43 comet analysis, kymograph generation, and assessment of EM images was performed using publicly available plugins for ImageJ/FIJI software (National Instituted of Health).

Maximum intensity projection images from en face neuromuscular junction synapses were

83 analyzed for synaptic area, pre- and post- synaptic marker intensities using custom MATLAB

software (MathWorks 2018). Cell counting was performed either using ImageJ/FIJI or

MATLAB software. Statistical analyses were performed, and graphs were created using Prism

version 8.0 (GraphPad). Most graphs display all data points to accurately represent the variability

in each dataset, except in cases where such presentation obscured the conclusion. For all

statistical analyses, means were considered to be significantly different if p < 0.05. Statistical

analyses used in each experiment are indicated in their respective figure legends. For all graphs

mean ± SEM was reported unless otherwise noted. Normality of each dataset was assessed using a Shapiro-Wilk test. In datasets with two groups, parametric data was analyzed using a Student’s t-test, while non-parametric data was assessed by Mann-Whitney U analysis of medians.

Multiple groups were compared by one-way or two-way ANOVA and analyzed post hoc by

either a Bonferroni or Kruskal-Wallis test for parametric and non-parametric data, respectively.

84 Results

Tuba1a is a major component of adult a-tubulin mRNA

During embryonic development, Tuba1a constitutes approximately 95% of all a-tubulin

mRNA in the mouse brain [16]. By comparison, postnatal Tuba1a expression is dramatically

decreased, however certain brain regions including the neocortex and hippocampus retain

Tuba1a expression into adulthood [27]. Many studies have focused on the role of Tuba1a during

brain development, but it is unclear how much Tuba1a contributes to the a-tubulin pool in adult

neurons of the brain and spinal cord. Although TUBA1A is ubiquitously expressed in post-mitotic

neurons, the relative abundance of different a-tubulin isotypes in specific neuronal subtypes over

time has not been well characterized. To understand how the neuronal a-tubulin isotype blend

changes over time, we assessed mRNA for Tuba1a, Tuba1b, Tuba1c and Tuba4a in the cortex

and spinal cord of P0 and adult wild-type mice (Fig. 3.1A-C; N=3). Unfortunately, levels of

Tuba1c mRNA were too low to be reliably detected in our assays, thus we have omitted those

data. Although the relative quantity of mRNA for each isotype differed between the cortex and

spinal cord, we found that the ratio of a-tubulin isotypes was comparable between spinal cord

and cortex, in both newborn and adult animals (Fig. 3.1D; N=3). Intriguingly, although Tuba1a

expression is downregulated between development and adulthood, we found that in wild-type

adult mice, Tuba1a comprises 32% of all cortical and 47% of all spinal cord a-tubulin mRNA

(Fig. 3.1D). These data demonstrate that although Tuba1a mRNA abundance decreases

postnatally, Tuba1a is the most abundant a-tubulin isotype in adult spinal cord and the second- most abundant a-tubulin in adult cortex. Because Tuba1a constitutes a large percentage of the total a-tubulin in both cortical and spinal neurons over time, we predict that Tuba1a plays an important role in the adult neuronal microtubule network.

85 Tuba1aND is a loss-of-function mutation in mice

While it is known that Tuba1a contributes significantly to development, it has been difficult to determine how this isotype contributes to adult neuronal function due to the lack of available tools to study Tuba1a protein. My results from Chapter II and previous data in yeast

[183] suggested that heterozygous Tuba1aND/+ mice may have reduced Tuba1a function. To determine if the Tuba1aND substitution affects levels of a-tubulin in mice, we isolated protein from wild-type and Tuba1aND/+ brains at P0-2. We did not include homozygous Tuba1aND/ND mouse brain lysates due to their perinatal lethality. Quantitative western blots revealed an approximate 50% decrease in a-tubulin protein in heterozygous Tuba1aND/+ mouse brains compared with wild-type at birth (Fig. 3.2A, B; N=4, p<0.0001) consistent with the molecular impact of the ND substitution in yeast reported in Hanson et al. (2016) and in cells (Chapter II).

This reduction in a-tubulin protein was not due to a change in brain size, as weights between wild-type and Tuba1aND/+ brains were comparable (Supplemental Fig. 3.1); N=10, p=0.68).

Adult Tuba1aND/+ a-tubulin protein levels were comparable to wild-type (Fig. 3.2A, B; N=6,

p=0.38). The observed reduction in developmental a-tubulin indicates that the availability of a-

tubulin protein is decreased in Tuba1aND/+ neurons during a critical period for neuronal

microtubule network establishment. As total a-tubulin protein levels normalized in the brains of

adult Tuba1aND/+ mice, we assessed brain tubulin post-translational modifications (PTMs) because PTMs influence binding interactions with microtubule associated proteins (MAPs) and overall microtubule function. Western blots of whole brain lysates from adult wild-type and

Tuba1aND/+ mice revealed no genotypic difference in the abundance of acetylated (N=5; p=0.66), tyrosinated (N=3; p=0.52), detyrosinated (N=5; p=0.81), or polyglutamylated (N=3; p=0.31) a- tubulin relative to the total a-tubulin (Fig. 3.2C-D). Therefore, we conclude that Tuba1aND

86 substitution reduces the availability of a-tubulin protein developmentally, but does not impact the PTM landscape of adult brain microtubules.

Figure 3.1 Changes in a-tubulin isotype expression in cortex and spinal cord between postnatal day 0 (P0) and Adult. A. Bar graphs representing relative quantity of Tuba1a mRNA in P0 (left) and adult (right) mouse cortex and spinal cord. B. Bar graph representing relative quantity of Tuba1b mRNA in cortex and spinal cord of P0 (left) and adult (right) mice. C. Bar graph representing relative quantity of Tuba4a mRNA in cortex and spinal cord of P0 (left) and adult (right) mice. D. Bar graphs representing the a-tubulin isotype mRNA expression data from A-C as a percentage of the total a-tubulin isotype composition. Three animals per genotype were analyzed for each time point, with three technical replicates per animal. Data are plotted as relative mRNA quantity normalized to housekeeping gene Cyclin A.

87

Figure 3.2 a-tubulin protein is decreased in P0 Tuba1aND/+ brains. A. Western blots representing a-tubulin abundance in P0 (left) and adult (right) Tuba1aND/+ and wild-type mouse brains. B. Bar graph quantifying a-tubulin protein in P0 and adult brain tissue lysates relative to GAPDH (N=3 mice, p<0.0001 by Two-Way ANOVA). Quantifications are representative of at least three separate experiments. C. Western blots showing the abundance of a-tubulin post- translational modifications (PTMs) acetylation (top left), tyrosination (top right), detyrosination (center left), and polyglutamylation (center right), and total a-tubulin (bottom) in adult wild-type and Tuba1aND/+ whole brain lysates. D. Bar graph quantification of western blot data shown in C. PTM band volume was normalized to a-tubulin (p>0.05 for all by t test). E, F. mRNA expression of brain a-tubulin isotypes in Tuba1aND/+ and wild-type mice at P0. Data are plotted as fold change in Tuba1aND/+ relative to wild-type (E.) or as a percentage of the total a-tubulin isotype composition (F.) (p>0.05 for all by two-way ANOVA). G, H. Bar graph representing mRNA expression of brain a- tubulin isotypes in adult Tuba1aND/+ and wild-type mice. Data are plotted as fold change in Tuba1aND/+ relative to wild-type (G.) or as a percentage of the total a-tubulin isotype composition (H.) (p>0.05 for all by two-way ANOVA). All western blot data represents three replicate blots with lysates from at least three animals per genotype.

88 Mice have nine a-tubulin genes and mammalian neurons express at least four distinct a- tubulin genes [14, 16, 27, 126]. Mutations that disrupt the major isoform, Tuba1a, could potentially induce compensatory changes to the expression of other a-tubulin mRNAs. We found that mRNA expression of Tuba1a and other brain a-tubulin isotypes was not significantly changed in heterozygous Tuba1aND/+ P0 brain lysates (Fig. 3.2E; p>0.9 for all comparisons by two-way ANOVA). Additionally, the relative ratio of each a-tubulin isotype mRNA was unchanged in Tuba1aND/+ P0 brain lysates, with Tuba1a constituting approximately 98% of all a-tubulin mRNA in both wild-type and Tuba1aND/+ brains (Fig. 3.2F; N=3, p=0.92). We conclude from these results that alternative a-tubulin isotypes are not upregulated to compensate for the loss of Tuba1a protein in the developing brain. We examined the abundance of a-tubulin isotype mRNAs in the cortex of adult wild-type and Tuba1aND/+ mice and found that Tuba1a remains one of the predominant a-tubulins for both genotypes (Fig. 3.2G; N=3; p>0.7 for all by two-way ANOVA). The modest, but statistically insignificant increase in Tuba1a mRNA in the adult Tuba1aND/+ cortex slightly shifted the ratio of a-tubulin isotype mRNAs in the adult cortex, with an increased percentage of Tuba1a a-tubulin mRNA compared to the other isotypes (Fig.

3.2H; p<0.0001 in Tuba1aND/+ cortex compared to wild-type by two-way ANOVA). Together,

these data demonstrate that Tuba1aND/+ transiently depletes a-tubulin protein during a critical developmental window, without inducing compensatory upregulation of other a-tubulin isotypes.

89 Reduced Tuba1a function results in fewer microtubule tracks in neurites

To determine how reduced Tuba1a impacts the development of a neuronal microtubule

network, we used primary neurons from wild-type and Tuba1aND/+ P0-P2 cortices to examine microtubule organization in vitro. Wild-type and Tuba1aND/+ primary cortical neurons were nucleofected with membrane-bound Myr-TdTomato and GFP-MACF43, which binds to polymerizing microtubule plus-End Binding proteins (EBs) without affecting microtubule function [173]. Kymograph plots generated from GFP-MACF43 videos allowed us to interrogate the number of growing microtubule plus-ends per micrometer of neurite at DIV 1 (Fig. 3.3A-C).

We found that Tuba1aND/+ neurites have approximately half the number of GFP-MACF43 puncta

per micrometer of neurite as wild-type neurites (2.610±0.20 growing microtubule plus-ends in

Tuba1aND/+ vs 4.636±0.34 in wild-type, Fig 3.3D; n=24 neurites; p<0.0001). These data suggest the approximate 50% reduction in a-tubulin protein observed in developmental Tuba1aND/+ mice

causes functional loss of microtubule track density within developing Tuba1aND/+ neurons.

Tuba1a is required for functional intracellular transport

One critical function of neuronal microtubules is to facilitate intracellular trafficking of proteins, mRNAs, and organelles between the cell body and distal neurites. Thus, we explored the possibility that reduced Tuba1a function and a reduced density of microtubule tracks impairs microtubule-based intracellular transport. To evaluate how loss of Tuba1a impacts the dynamics of microtubule-based transport, we examined transport of lysosomes in cultured primary neurons from P0-P2 wild-type and Tuba1aND/+ cortices.

90

Figure 3.3 Reduced TUBA1A function results in fewer microtubule tracks in neurites. A. Schematic of experimental design, illustrating GFP-MACF43 puncta analysis and kymograph generation. B. Representative images of DIV 1 wild-type and Tuba1aND/+ neurons, visualized using a membrane-bound Myr-TdTomato. Scale bars are 10µm. C. Representative kymograph plots generated from GFP-MACF43 images in wild-type (left) and Tuba1aND/+ (right) neurons. Scale bars are 5µm. D. Bar graph representing the number of GFP-MACF43 puncta per µm of neurite in DIV 1 wild-type and Tuba1aND/+ cortical neurons. Bars illustrate mean density of GFP-MACF43 puncta per kymograph and error bars indicate SEM (n=24 neurons, p<0.0001). Statistical differences between groups were assessed by t test.

91 Lysosomes facilitate enzymatic degradation of intracellular components and are highly

motile organelles that are transported bidirectionally on microtubules through all compartments

of the neuron, making them an ideal organelle in which we could examine microtubule-based

transport efficiency. Time-lapse images of lysosome movement in wild-type or Tuba1aND/+

cortical neurons were used to generate kymograph plots of lysosome movement over time within

single wild-type and Tuba1aND/+ axons and dendrites (Fig. 3.4A, B), from which we could

measure velocity, dynamics of movement, and total distance traveled. From these analyses we

found that during the 3-minute imaging period, approximately 70% of wild-type lysosomes were

moving, while the remainder were stationary for the duration of the video (Fig. 3.4C; N=3 mice,

n=25 neurons). A significantly higher number of Tuba1aND/+ lysosomes were stationary compared to wild-type, with only 46% of Tuba1aND/+ lysosomes moving during the 3-minute

window (Fig. 3.4C; p=0.001). Of those lysosomes that were moving, the velocity of each

movement was unchanged between wild-type and Tuba1aND/+ neurons, with average velocity of

0.56±0.02µm/s and 0.53±0.02µm/s for wild-type and Tuba1aND/+ neurons, respectively (Fig.

3.4D; n=635 lysosomes, p=0.27). During active transport, cargoes exhibit phases of movement and pausing, thus we wanted to evaluate the dynamics of lysosome movement by measuring pause duration for moving lysosomes. There was no difference in pause duration of moving lysosomes in Tuba1aND/+ neurons, which paused on average 17.51±1.0s, compared to a

16.19±0.81s pause duration in wild-type neurons (Fig. 3.4E; n=337 events, p=0.56). Finally, we observed a significant decrease in the total distance traveled by lysosomes over the 3-minute imaging window, from 24.77±1.84µm in wild-type neurons to 17.19±1.66µm in Tuba1aND/+

neurons (Fig. 3.4F; n=100 neurons, p=0.003). These data demonstrate that Tuba1aND/+

92 substitution causes a deficit in neuronal intracellular transport by which Tuba1aND/+ lysosomes do not travel as far as wild-type within the same time window.

To further understand the impact of reduced Tuba1a function on neuronal microtubule- based transport, we examined mitochondrial trafficking in cultured primary Tuba1aND/+ neurons compared to wild-type. Transport of mitochondria and lysosomes is carried out by a different set of kinesins [194, 274-276]. Therefore, examining transport of multiple cargoes provides mechanistic insight into the intracellular trafficking disruption in Tuba1aND/+ neurons. The

analyses described above were repeated in DIV 7 wild-type and Tuba1aND/+ cortical neurons that had mitochondria labeled (Supplemental Fig. 3.2). Over the course of a 2-minute imaging period, we observed that the number of moving mitochondria was similar between wild-type (36%) and

Tuba1aND/+ (40%) neurons, and the remainder were stationary for the duration of the video (Fig.

3.4G; N=3 mice, n=30 neurons, p=0.38). Importantly, the relative number of stationary mitochondria was more than double the number of stationary lysosomes in wild-type neurons

(64.78±2.9% and 30.01±5.0%, respectively; Fig. 3.4C, 3.4G). Similar to what was observed for lysosomal trafficking, we found that the velocity of moving mitochondria was unchanged between wild-type (0.45±0.02µm/s) and Tuba1aND/+ (0.43±0.02µm/s) neurons (Fig. 3.4H; n=301 events, p=0.51). These mitochondrial movements were interrupted by differential periods of pause, with a significant increase in Tuba1aND/+ mitochondrial pausing compared to wild-type, with average pause durations of 28.36±1.6s and 24±1.4s, respectively (Fig. 3.4I; n=251 events, p=0.02). The total distance traveled by motile mitochondria during the 2-minute imaging window was found to be significantly decreased in Tuba1aND/+ neurons, with Tuba1aND/+ mitochondria traveling 9.46±1.3µm on average compared to 15.8±1.7µm in wild-type neurons

(Fig. 3.4J; n=301 events, p=0.01). Combined, these analyses indicate that a Tuba1a deficit

93 impairs transport of multiple organelle types in cortical neurons, resulting in a significant reduction in the total distance of organelle movement.

Figure 3.4 Tuba1aND/+ neurons have deficits in organelle transport caused by increased stationary cargoes. A. Still images from time-lapse microscopy of DIV 3 cortical neurons labeled with LysoTracker dye to mark lysosomes in wild-type and Tuba1aND/+ neurons. B. Representative kymograph plots of lysosome movement over time within select neurites for wild-type (top) and Tuba1aND/+ (bottom). C. Scatter plot representing the percent of stationary lysosomes per kymograph in wild-type and Tuba1aND/+ neurons. Data points represent individual neurons (N=3 mice, n=25 neurons, p=0.001). D. Scatter plot representing lysosome velocities in µm/s for wild-type and Tuba1aND/+ neurons. Data points represent individual lysosomes (n=635 lysosomes, p=0.27) E. Scatter plot representing pause duration (s), for moving lysosomes in wild-type and Tuba1aND/+ neurons. Data points represent individual pause events (n=338 events, p=0.56). F. Scatter plot representing total distance traveled (µm) by moving lysosomes in wild-type and Tuba1aND/+ neurons. Data points represent individual lysosomes (n=87, p=0.003). G. Scatter plot representing the percentage of stationary mitochondria in wild-type and Tuba1aND/+ neurons. Data points represent individual neurons (N=3 mice, n=30 neurons, p=0.34 by). H. Scatter plot the velocity of short mitochondrial movements in wild-type and Tuba1aND/+ neurons. Data points represent individual mitochondria (N=3 mice, n=301 mitochondria, p=0.45). I. Scatter plot representing pause duration (s), for moving mitochondria in wild-type and Tuba1aND/+ neurons. Data points represent individual pause events (n=417events, p=0.79). J. Scatter plot representing total distance traveled (µm) by moving mitochondria in wild-type and Tuba1aND/+ neurons. Data points represent individual mitochondria (n=86, p=0.01). Cortical neurons from three animals per genotype were analyzed. Lysosomes were imaged once every second for three minutes, mitochondria were imaged once every two seconds for two minutes. Statistical differences between groups were assessed by t test, **p<0.01; ***p<0.001.

94 Diminished Tuba1a causes late-onset behavioral deficits

Having established that neurons from young Tuba1aND mutant animals exhibit deficits in microtubule density and organelle trafficking, we next asked whether these lead to impairments at the level of behavior. The Tuba1aND mutant allele was identified due to locomotor deficits apparent in homozygous mutant embryos. Tuba1aND/ND homozygous substitution induced severe neurodevelopmental abnormalities that impaired motor neuron function and caused perinatal lethality [183]. In contrast, Tuba1aND/+ embryos survive to adulthood and were indistinguishable from wild-type siblings at the age of weaning (P21). Tuba1aND/+ embryos exhibit normal pre- and post- synaptic motor development within the diaphragm and respond to touch stimulation

[183]. However, as Tuba1aND/+ mice aged, we noticed differences in locomotion that were quantified with gait analysis. While the gait of 2-month-old Tuba1aND/+ mice is not significantly different from wild-type (N=2), by 3-months of age, Tuba1aND/+ mice develop an ataxic gait, as evidenced by a wider rear stance (Fig. 3.5A, B; N=5, p=0.02). Tuba1aND/+ mice developed impaired motor coordination by 5-months of age, as assessed by rotarod performance (Fig. 3.5C;

N=10, p=0.03). After the initial onset of behavioral phenotypes, Tuba1aND/+ mice performed worse than wild-type at all time-points examined (p<0.05 for all). Heterozygous Tuba1aND/+ mice

did not display deficits in motor learning as assessed by the difference in rotarod performance

between trials (Fig. 3.5D; N=10, wild-type R2=0.98, Tuba1aND/+ R2=0.89, p=0.31 by linear regression).

95

Figure 3.5 Tuba1aND/+ mice exhibit adult-onset motor and sensory behavioral deficits. A. Images depicting gait abnormalities in Tuba1aND/+ adult mice compared to wild-type. Still images were taken from video of Tuba1aND/+ and wild-type mice mid-stride. B. Line graph quantifying changes in rear stance width at 2-months (p=0.61), 3-months (p=0.02), 5-months (p=0.04), 8-months (p=0.03), and 10-months of age (p<0.0001) in wild-type and Tuba1aND/+ mice. C. Line graph representing latency to fall on rotarod task at 2-months (p=0.31), 3-months (p=0.82), 5-months (p=0.03), 8-months (p=0.004), and 10-months of age (p=0.04) in wild- type and Tuba1aND/+ mice. Points depict the mean stance width or latency to fall by genotype with SEM. The same mice were analyzed throughout, and a two-way ANOVA was used to assess statistical significance between groups. Number of mice per genotype is as follows: N=2 at 2 months, N=5 at 3 months, N=8 at 5 months, N=12 at 8 months, N=8 at 10 months. D. Line graph representing performance over three subsequent rotarod trials in a single testing session at 5-months of age for Tuba1aND/+ compared to wild-type. Slopes of the lines for wild-type and Tuba1aND/+ were compared by linear regression and were not found to be significantly different (p=0.32, wild-type R squared=0.98, Tuba1aND/+ R squared=0.89). E. Scatter plot of grip strength assessed in the forelimbs of female 5-month-old wild-type and Tuba1aND/+ mice (N=7, p=0.19). F. Bar graph of total body weight for 8-10-month old Tuba1aND/+ and wild-type female mice (N=7, p=0.24). G. Bar graph of SUDO scores for Von Frey mechanical sensation testing in 3-6-month old wild-type and Tuba1aND/+ mice (N=7, p=0.04). H. Bar graph of time to withdraw from thermal stimulus in Hargreaves behavioral analysis on 3-month old wild-type and Tuba1aND/+ mice (N=7, p=0.19). Sex as a variable did not significantly impact rear stance width or rotarod performance at any time point examined (N=7 females, N=5 males; p>0.05 for all comparisons of male vs. female). Sex was also not found to significantly influence sensory behavior (N=7 males, N=2 females; p>0.05 for all comparisons).*p<0.05, **p<0.01, ****p<0.0001.

96 Locomotor impairment of these mice impeded more rigorous testing of cognitive function.

Importantly, forelimb function, and all other parameters of gait were similar to that measured in

wild-type animals (Supplemental Fig. 3.3). Body weights of Tuba1aND/+ mice were comparable

to that of control mice (Fig. 3.5F; N=7, p=0.24), and forelimb muscular strength was not

impaired (Fig. 3.5E; N=7, p=0.19). These results indicate that neither altered muscle content nor

impaired gross muscle function are responsible for the observed motor deficits, indicating

Tuba1aND-induced neuronal dysfunction is a likely cause. The hindlimb-specificity of this

behavioral phenotype further suggests that neurons with extremely long axons, such as those that

innervate the hindlimbs, may be exquisitely sensitive to perturbations of Tuba1a.

Tuba1a is expressed in all post-mitotic neurons [27, 277], therefore we predicted that

behavioral impairments induced by Tuba1aND/+ substitution would not be restricted to motor neurons. To assess whether Tuba1aND/+ adult mice had deficits in sensory behavior, we examined

hind-paw sensory function using two separate assays in mature mice. Response to hind-paw

stimulation with Von Frey filaments revealed an increased threshold for retraction in Tuba1aND/+

compared to wild-type mice, indicating reduced sensation of mechanical stimuli (Fig. 3.5G; N=7,

p=0.04). Additionally, adult Tuba1aND/+ animals displayed a modest, but not statistically

significant decrease in thermal pain sensation of the hind paws, as measured by a Hargreaves

assay (Fig. 3.5H; N=7, p=0.19). These results support our prediction that Tuba1aND/+-induced behavioral deficits are not restricted to the motor system, but rather appear to preferentially impact the far-reaching neurons that innervate the hindlimbs. The degenerative behavioral phenotypes in Tuba1aND/+ mice indicate that Tuba1a deficiency has long-lasting consequences

for neuronal function.

97 Tuba1a deficit impairs hindlimb behavior without causing cell death or axon degeneration

Complex motor behaviors involve multiple types of motor neurons coordinated through

multiple brain regions including the cortex, cerebellum, and spinal cord. We examined survival

of neurons that are important for coordinated movement in each of these systems in Tuba1aND/+

and wild-type mice. Cortical lamination occurs normally in Tuba1aND/+ mice, and there was no evidence of heterotopic neurons in Tuba1aND/+ brains (Fig. 3.6A; N=3). To assess cortical motor neuron number, we used the transcription factor ER81 as a marker of layer V cortical neurons that project to the spinal cord [278]. We found no difference in the number of ER81+ cortical motor neurons in layer V of the motor cortex and no evidence of apoptosis before or after the onset of quantifiable motor deficits in Tuba1aND/+ and wild-type (Fig. 3.6A, B, Supplemental

Fig. 3.4; p=0.1). We quantified the number of Purkinje neurons, the primary output cells of the cerebellum, and found no difference in Purkinje cell number between genotypes at either time point (Fig. 3.6A, C; N=3, p=0.12). Quantification of motor neurons in serial sections through the lumbar spinal cord, which contains neurons that innervate the rear limbs, revealed that there was no alteration to the number of spinal motor neurons in Tuba1aND/+ animals compared to wild-

type (Fig. 3.6A, D; N=3, p=0.2). In electron micrographs of mature spinal cords from

Tuba1aND/+ and wild-type animals (Fig. 3.7A; N=2 mice), we found no difference in myelin

thickness by G-ratio (Fig. 3.7B; n=100 axons, p=0.42), axon density (Fig. 3.7C; p=0.72), or axon

diameter (Fig. 3.7D, E; p=0.63) at 3-months or 10-months of age. Reduced neuron survival, loss

of myelination, and axon degeneration are reported in several movement disorders [34, 184, 186,

279-281]. However, these results indicate that reduced Tuba1a function does not impact the

establishment or maintenance of spinal motor neuron density, axon caliber, or myelination.

98

Figure 3.6 Tuba1aND/+ does not impact neuronal cell body survival. A. Top row: Coronal sections of motor cortex immunolabeled with Calbindin (green), ER81 (magenta), and DAPI (blue) in wild-type and Tuba1aND/+ mice at 3-months (left) and 10-months of age (right) at 20X magnification. Center row: Sagittal sections of cerebellum labeled with Calbindin (green) and DAPI (blue) in wild-type and Tuba1aND/+ mice at 3-months (left) and 10-months of age (right) at 20X magnification. Bottom row: Nissl stained coronal sections of the lumbar spinal cord in wild-type and Tuba1aND/+ mice at 3-months (left) and 10-months of age (right) at 4X magnification. B. Scatter plot representing the number of ER81+ layer V neurons per image in motor cortex at 3- and 10-months (N=3 mice, p=0.19 and p=0.78, respectively). C. Scatter plot representing the number of Calbindin+ Purkinje neurons per image in the cerebellum at 3- and 10-months (N=3, p=0.94 and p=0.35, respectively). D. Scatter plot representing the number of Nissl+ ventral horn motor neurons per image in the spinal cord at 3- and 10-months (N=3, p=0.66 and p=0.99, respectively). Motor neurons were identified morphologically. Data points represent ROIs with horizontal line depicting mean with SEM. Data points were nested by animal and analyzed by two-way ANOVA.

We examined the possibility that reduced Tuba1a function causes axon degeneration. To

evaluate axon integrity in the Tuba1aND/+ model, we quantified non-phosphorylated neurofilament (SMI32) that is highly abundant in degenerating axons [282-284]. We sampled several regions of interest from cortex and observed no increase in SMI32 immunoreactivity in cortical sections from Tuba1aND/+ mice compared to wild-type (Fig. 3.7F, G; N=4, p=0.84). We also quantified SMI32 abundance in Tuba1aND/+ and wild-type whole brain lysates and detected no difference in SMI32 protein between the two groups (Fig. 3.7H, I; N=3, NFT-M p=0.61,

99 NFT-H p=0.70). Overall, these data support the conclusion that reduction of Tuba1a causes adult-onset behavioral deficits without obvious detriment to neuronal survival or morphology.

Figure 3.7. No deficits in axon morphology, myelination, or survival in Tuba1aND/+ mice. A. Electron micrographs of spinal cord axons in cross-section for adult wild type (WT; left) and Tuba1aND/+ mice (right). Six images per animal and two animals per genotype were assessed for EM analyses. B. Scatter plot displaying myelin thickness measure (G-ratio) for two wild-type and two Tuba1aND/+ mice (N=2 mice, n=100 axons, p=0.42). C. Scatter plot of axon density quantified in electron micrograph images for adult wild-type and Tuba1aND/+ mice (N=2 mice, n=6 fields, p=0.72). D. Scatter plot of axon diameter quantified in electron micrograph images for adult wild-type and Tuba1aND/+ mice (N=2 mice, n=100 axons, p=0.63). E. Histogram representing the distribution of axon diameters from data in (D). F. Anti-SMI-32 immunofluorescence images in cortex of adult wild-type (left) and Tuba1aND/+ mice (right). 40X magnification, scale bar is 10 µm. G. Bar graph representing SMI-32 intensity in a cortical region of interest for wild-type and Tuba1aND/+ mice. Data points represent regions of interest, with three animals per genotype assessed (p=0.77). H. Western blot for SMI-32 in whole brain lysates from adult wild-type and Tuba1aND/+ mice. Bands for heavy (200kDa), medium (160kDa), and light (125kDa) chain non-phosphorylated neurofilament shown (top). GAPDH used for normalization (37kDa; bottom). Data points represent technical replicates with three biological replicates performed. I. Bar graphs representing quantification of SMI- 32 medium (right, p=0.61) and heavy (left, p=0.70) expression in whole brain lysates, normalized to GAPDH.

100 Neuromuscular junction synapses are diminished by prolonged microtubule dysfunction

Finally, we examined whether Tuba1aND/+ impacts synaptic structure at the neuromuscular junction (NMJ). NMJ function is critical to motor output, as it allows for the conversion of electrochemical signaling in neurons to muscle contraction and movement. To assess NMJ synaptic morphology, we labeled acetylcholine receptors (AChRs) with fluorescently conjugated a-Bungarotoxin in extensor digitorum longus (EDL) muscles of both juvenile and adult Tuba1aND/+ and wild-type hindlimbs. In rodents, NMJ synapses begin forming embryonically, and complete the final stages of synapse maturation, including synaptic pruning and stabilization, between 2-3 weeks postnatal [285]. Juvenile Tuba1aND/+ NMJ synapses were

evaluated at 1-month of age, prior to the onset of quantifiable behavioral deficits, but following

the completion of most NMJ development. Wild-type and Tuba1aND/+ synapses labeled for post-

synaptic AChRs as well as pre-synaptic vesicle marker synaptophysin were evaluated for

synaptic area and the relative density of both pre- and post-synaptic machinery (Fig. 3.8A).

Juvenile Tuba1aND/+ EDL NMJ synapses were indistinguishable from wild-type in synaptic area

(Fig. 3.8B; n=69 synapses, p=0.3), density of pre- and post-synaptic components (Fig. 3.8D, C;

N=3 mice, n=39 synapses, p>0.5 for both), and the ratio of pre- to post-synaptic machinery (Fig.

3.8E; n=39 synapses, p=0.74). The same analyses were performed for EDL synapses in adult (~1 year) wild-type and Tuba1aND/+ mice. A dramatic reduction in synaptic branching complexity in

the Tuba1aND/+ EDL was immediately apparent, illustrated by the example images (Fig. 3.8A).

Further, in adult Tuba1aND/+ NMJs, synaptic area was found to be significantly reduced compared to wild-type (Fig. 3.8F; N=3 mice, n=45 synapses, p<0.0001). Additionally, adult

Tuba1aND/+ synapses contained a higher density of post-synaptic AChRs than wild-type (Fig.

3.8G; p<0.0001). The increased AChR density in the adult EDL was also accompanied by an

101 increase in pre-synaptic synaptophysin (Fig. 3.8H), but the ratio of pre- to post-synaptic machinery was modestly decreased in Tuba1aND/+ NMJs compared to wild-type, indicating that

Tuba1aND/+ NMJs had lower pre-synaptic vesicle density relative to the post-synaptic receptor density (Fig. 3.8I; p<0.0001 and p=0.03, respectively). Collectively these data demonstrate that reduced Tuba1a diminishes NMJ size and alters NMJ synapse morphology and functional properties in an age-related manner.

102

Figure 3.8 Neuromuscular junction synapses deteriorate over time in Tuba1aND/+ mice. A. Teased EDL muscle fibers labeled with a-Bungarotoxin (cyan) and synaptophysin (magenta) for 1-month-old (top) and 1-year-old (bottom) wild-type and Tuba1aND/+ mice. Muscle fibers from three mice per genotype at each time point were assessed. B. Scatter plot of EDL synaptic area measured from a-Bungarotoxin labeling in juvenile wild-type and Tuba1aND/+ mice (N=3 mice, n=69 synapses, p=0.43). C. Scatter plot of post- synaptic acetylcholine receptor (AChR) density measured as synaptic a-Bungarotoxin intensity divided by synaptic area in juvenile mice (N=3 mice, n=39 synapses, p=0.89). D. Scatter plot of pre-synaptic vesicle density measured as synaptic synaptophysin intensity divided by synaptic area in juvenile mice (N=3 mice, n=39 synapses, p=0.54). E. Scatter plot of the ratio of synaptophysin to AChR (pre- to post-synaptic) intensity in juvenile wild-type and Tuba1aND/+ mice (N=3 mice, n=39 synapses, p=0.74). F. Scatter plot of EDL synaptic area measured from a-Bungarotoxin labeling in adult wild-type and Tuba1aND/+ mice (N=3 mice, n=45 synapses, p<0.0001). G. Scatter plot of post-synaptic AChR density measured as synaptic a-Bungarotoxin intensity divided by synaptic area in adult mice (N=3 mice, n=45 synapses, p<0.0001). H. Scatter plot of pre-synaptic vesicle density measured as synaptic synaptophysin intensity divided by synaptic area in adult mice (N=3, n=45, p<0.0001). I. Scatter plot of the ratio of synaptophysin to AChR (pre- to post-synaptic) intensity in adult wild-type and Tuba1aND/+ mice (N=3 mice, n=45 synapses, p=0.03). Data points on graph represent individual synapses. Data were nested by animal and analyzed for statistical significance between genotypes by a two- way ANOVA. ****p<0.0001.

103 Discussion

Tuba1a is critical for developing and mature neurons

Here, we show that significant reduction in developmental total a-tubulin resulting from

a LOF mutation in Tuba1a (Tuba1aND/+) causes an adult onset movement disorder. We show that assembly of the microtubule network is altered resulting in disrupted organelle trafficking early in development in cultured Tuba1aND/+ cortical neurons, suggesting that neuronal microtubule

networks are not set up correctly when there is not adequate a-tubulin available (Figs 3.1-3.4).

While these aberrant neuronal microtubule networks are sufficient to establish juvenile NMJ

synapses similarly to wild-type, Tuba1aND/+ NMJ synapses deteriorate over time (Fig. 3.8),

potentially contributing to adult-onset motor deficits. This work provides the first evidence that

the ‘developmental’ tubulin TUBA1A is necessary for adult neuronal function and suggests that

mature neurons require TUBA1A for proper function throughout their lifetime.

Tuba1a is Required for Intracellular Transport

The microtubule cytoskeleton forms the tracks upon which motor proteins bind to

facilitate intracellular transport. Mitochondria and lysosomes are both transported by dynein

motor proteins in the retrograde direction, but differ in the kinesin motors that primarily regulate

their anterograde movement, with lysosome transport dominated by kinesin-2 and mitochondrial

transport occurring primarily by kinesin-1 [274, 275, 286]. Thus, examining transport dynamics

for these two different organelles allowed us to distinguish between compromised interactions

with specific motor proteins or universal effects of a compromised microtubule network. Neither

lysosome nor mitochondria velocity was affected by depletion of Tuba1a function suggesting

that motors interact with the microtubule substrate normally (Figs. 3.4D, H). Importantly, we did

104 not see differences in amounts of PTMs that are known to affect the activity of motors on their

microtubule substrates (Fig. 3.2C-D). Thus, this is the first demonstration that reducing available

Tuba1a affects organelle trafficking in living neurons.

Mitochondria and lysosomes move with similar average velocities in wild-type neurons,

however the dynamics of movement for these two organelle types vary greatly. We observed a

larger proportion of stationary mitochondria (60%) than stationary lysosomes (30%) in wild-type

neurons (Figs. 3.4C, G). The proportion of stationary mitochondria in our cortical cultures was

consistent with what has been previously reported in neurons [286-288]. The variation in

transport dynamics between these two organelle types could potentially explain the discrepancy

in trafficking deficits we observed between lysosomes and mitochondria in Tuba1aND/+ neurons.

Lysosomes are highly motile organelles; therefore, a lysosomal stalling deficit was readily detected in Tuba1aND/+ neurons (Fig. 3.4C). However, such a deficit was not apparent in the relatively stationary mitochondrial population (Fig. 3.4G). The subtle deficit in pause duration that was observed in mitochondria was not detectable in lysosomes, which had much shorter pause durations on average (Figs. 3.4E, I). Despite the discrepancies in how these abnormalities were detected between different organelles, the overall distance traveled by mitochondria and lysosomes was consistently reduced in Tuba1aND/+ neurons compared to wild-type (Figs. 3.4F,

J). The increase in pausing or stalling behavior (Figs. 3.4C, I), combined with the reduced a- tubulin protein during development, and normal neuronal survival (Figs. 3.2, 3.6, 3.7) suggests that reduced availability of Tuba1a is detrimental to adult neuronal function, but not neuronal survival.

105 Neuronal Tuba1a Establishes Microtubule Network Function

The abundance of a-tubulin protein during brain development is reduced in Tuba1aND/+ brains (Fig. 3.2). This deficit in a-tubulin reduced the number of growing microtubule plus-ends per micrometer of neurite in developing Tuba1aND/+ neurons, indicating the neuronal microtubule

network contains a lower density of microtubules in Tuba1aND/+mice (Fig. 3.3). We did not

observe differences in neuronal survival, axon caliber, myelination, or markers for axon

degeneration in Tuba1aND/+ compared to wild-type siblings (Figs. 3.6 and 3.7) suggesting that reduced Tuba1a impacts neuron function rather than neuron survival or axon morphology.

Collectively, these results could suggest that adult Tuba1a expression may be important for maintenance of microtubule tracks in neurons and mutant TUBA1AND microtubules are not

sufficient for that requirement. We favor a second model, that the developmental TUBA1A

deficit does not allow a neuron to correctly establish microtubule networks of the appropriate

density, and these defects cannot be sufficiently repaired to maintain functional NMJ synapses in

adulthood despite adequate levels of adult a-tubulin protein (Figs. 3.2, 3.3, 3.8).

Intracellular transport deficits in Tuba1aND/+ neurons provide a putative mechanism by which reduced Tuba1a impairs neuronal function to cause adult-onset behavioral deficits.

However, as cortical neurons are not representative of the adult condition, follow-up studies are

needed to determine if trafficking deficits persist into adulthood in the Tuba1aND/+ model.

Because developmental a-tubulin protein is reduced in Tuba1aND/+ brains without detectable compensation from any other a-tubulin isotypes (Fig. 3.2), we propose a model in which early a-tubulin deficits cause formation of inadequate neuronal microtubule tracks that cannot be rescued by subsequent restoration of a-tubulin protein. Over time, the consequences of early tubulin deficiency could become more pronounced and result in impairment of synaptic function

106 and animal behavior. Microtubules in neurons are not singly nucleated from the centrosome, but

rather exist in short segments that tile the length of the axons and dendrites [289]. When moving

cargoes reach the end of a microtubule segment, the motor protein pauses to switch tracks [290].

Thus, the observed increase in organelle pausing behavior paired with developmental decreases

in a-tubulin protein and microtubule density could indicate that motor proteins in Tuba1aND/+ neurons encounter microtubule ends more frequently, due to the presence of potential gaps in the tiled microtubule network. Alternatively, while no compensation for loss of Tuba1a was observed at the mRNA level, decreased a-tubulin protein would likely shift the a-tubulin isotype blend at the protein level. It has been previously shown that incorporation of different tubulin isotypes alters microtubule properties, whether directly or through effects on MAP binding [10-

12, 215]. Although a-tubulin protein levels are similar to wild-type in the adult Tuba1aND/+ brain, it is unclear whether adult Tuba1aND/+ microtubule polymers contain normal a-tubulin isotype compositions.

Tuba1a Contributes to Function of Mature Neurons

Microtubules and intracellular transport are important for synaptic signaling and maintenance [4, 291-293]. Neurons are morphologically complex cells, with processes often extending long distances from the soma, and are thus highly dependent on functional intracellular transport. Transport deficits, such as those observed in the Tuba1aND/+ model could impair both delivery and clearance of cargoes if persistent over time, leading to neuronal dysfunction. Functional intracellular transport is essential for delivery of mitochondria and synaptic components to the pre-synapse [294]. Thus, the transport deficits caused by Tuba1aND/+ substitution could alter synaptic transmission and thereby cause deterioration of synapse

107 morphology, potentially explaining the observed NMJ deficits (Fig. 3.8). The increased density

of both pre- and post-synaptic machinery that was observed in the Tuba1aND/+ NMJ (Fig. 3.8G,

H) suggests a possible impairment in clearance of vesicles from the pre-synapse and receptors

from the post-synapse. Moreover, the reduction in NMJ synaptic area (Fig. 3.8F) and branching

complexity indicates that the increased density of AChRs and pre-synaptic vesicles may be a

form of compensation for improperly functioning synapses. This hypothesis is further supported

by the observed dysregulation of pre- to post-synaptic component ratios in adult Tuba1aND/+ synapses (Fig. 3.8I). NMJ synapses are generally quite stable over time, but are known to undergo morphological changes in response to processes that alter synaptic transmission, such as diseases of the neuromuscular system and aging [281, 295-297]. Decreased NMJ size explains impaired motor behaviors in Tuba1aND/+ mice (Fig. 3.5A-C). However, given that Tuba1a is

expressed in all neurons, and Tuba1aND/+ mice also exhibited degenerative sensory deficits (Fig.

3.5G, H), we do not expect that the NMJ is the only synapse type affected by deficiency in

Tuba1a, but this idea remains to be tested. The prominence of hindlimb-specific behavioral

phenotypes in the Tuba1aND/+ mouse model suggests that axon length may predict sensitivity to

tubulin-deficiency.

Microtubules in Neurodegeneration: Illuminating New Roles for TUBA1A

Microtubule dysfunction, including intracellular trafficking defects, have been described

in nearly every neurodegenerative disease of the nervous system, and can precede other signs of

neuronal dysfunction [34, 39, 41, 184, 298]. Although there is abundant evidence of microtubule

dysfunction in neurodegenerative disease, it has been difficult to pinpoint how or if the

microtubule cytoskeleton contributes to disease etiology. Microtubule dynamics, PTMs, and

108 MAP interactions become dysregulated in neurodegenerative diseases such as Alzheimer’s and

Parkinson’s disease, and ALS, however it has remained unclear as to whether cytoskeletal

dysfunction is causative of disease or merely correlated with its progression [36, 39-41, 95].

Mutations that disrupt the neuronal a-tubulin TUBA4A have been identified familial and

sporadic ALS [198, 299-301]. The mechanisms by which TUBA4A mutations may cause

neurodegeneration remain unclear. Using the Tuba1aND/+ model, we have demonstrated that LOF in a major neuronal a-tubulin is sufficient to induce adult-onset behavioral degeneration.

Further, we have shown that Tuba1aND/+ does not induce neuronal death or degeneration, but

rather causes synaptic abnormalities, potentially by impairing intracellular transport. We provide

direct evidence that deficits in microtubule function can cause degenerative neuronal pathology

at the molecular, cellular and behavioral level.

109 Supplemental Figures

Supplemental Figure 3.1 Tuba1aND/+ does not alter brain weight at birth. A. Scatter plot of brain weight for Tuba1aND/+ and wild-type mice at P0-P2 (N=10 mice, p=0.68 by t test). Weights were recorded from frozen, dissected brains.

Supplemental Figure 3.2 Tuba1aND/+ impairs mitochondrial transport by increasing pause duration. A. Still images from time-lapse microscopy of DIV 3 cortical neurons labeled with MitoTracker dye to mark mitochondria in wild-type and Tuba1aND/+ neurons. B. Insets show representative kymograph plots of mitochondrial movement over time within select neurites for wild-type (left) and Tuba1aND/+(right).

Supplemental Figure 3.3 Tuba1aND/+ does not impact forelimb gait and specifically impacts rear stance width. A. Scatter plot of front stance width in 5-month-old wild-type and Tuba1aND/+ mice. B, C. Scatter plot of forelimb (B) and hindlimb (C) stride length in 5-month-old wild-type and Tuba1aND/+ mice (N=8 mice, p>0.05 for all by t test).

110

Supplemental Figure 3.4 No evidence of apoptosis in Tuba1aND/+ cortex in young or old mice. A. TUNEL staining (green) with DAPI (blue) in wild-type (left) and Tuba1aND/+ (center) cortex, with DNase-treated positive control cortex (right). Sections from 3-month-old (top) and 10-month-old (bottom) animals are shown. No evidence of increased apoptosis by genotype was detected at either time point

111 CHAPTER IV

CONCLUSIONS3

Introduction

The results presented in this dissertation illustrate precise requirements for neuronal

TUBA1A during brain development and reveal a novel role for TUBA1A in mature neuron function. My results using a Tuba1a loss-of-function (LOF) mouse model (Tuba1aND) showed that TUBA1A haploinsufficiency did not replicate human tubulinopathies, and instead caused adult-onset neurological symptoms. First, I showed that a 50% reduction in Tuba1a was sufficient to support cortical migration, but did not support long-range commissural axon targeting (Chapter II). Reduced abundance of Tuba1a decreased axonal microtubule density

(Chapter III) and altered microtubule polymerization rates (Chapter II), collectively leading to impaired neuron growth in vitro. Tuba1a-deficient microtubules did not support normal axonal transport (Chapter III), and as such failed to normally regulate the growth cone cytoskeleton

(Chapter II). These data illustrate that developmental TUBA1A is critical for axon outgrowth and

regulates the growth cone cytoskeleton, but is not essential for cortical migration. Next, I

explored the requirement for TUBA1A in the adult mammalian brain. I found that Tuba1a is a

major component of adult neuronal a-tubulin mRNA (Chapter III), supporting a role for

TUBA1A outside of brain development. Reduced function in Tuba1a caused neuromuscular

junction (NMJ) synapses to deteriorate with age and resulted in adult-onset motor and sensory

deficits in Tuba1aND/+ mice (Chapter III). I provide direct evidence that microtubule dysfunction can be sufficient to induce age-related behavioral deficits, in the absence of neuronal

3 Portions of this chapter are published with permission from our previously published article: Aiken, J., et al., Tubulin mutations in brain development disorders: Why haploinsufficiency does not explain TUBA1A tubulinopathies. Cytoskeleton (Hoboken), 2019 [210].

112 degeneration in mice. My results define a novel role for TUBA1A in adult neurons, where it contributes to long-term maintenance of adult synapses and influences overall mature neuron function. Overall, the work presented in this dissertation refines understanding of how TUBA1A contributes to neurodevelopment, and identifies the long-lasting impact of TUBA1A on neuronal microtubule function.

TUBA1A in the Developing and Mature Brain

TUBA1A is a major component of developing and adult neuronal microtubules

From its discovery, TUBA1A has been widely regarded as a developmental tubulin isotype. This characterization of TUBA1A is largely based on two factors: 1) mRNA expression of TUBA1A peaks during embryonic brain development [14, 16, 27] and 2) human TUBA1A mutations cause cortical malformations and other neurodevelopmental phenotypes, termed tubulinopathies [18-22]. Tubulinopathy-associated mutations to TUBA1A cause severe brain defects and are frequently lethal either embryonically or in early childhood [18-22]. Due to the spectrum of neurodevelopmental phenotypes associated with mutations to TUBA1A, it has thus far been difficult to ascertain molecular and cellular requirements for TUBA1A in specific aspects of brain development. Further, TUBA1A tubulinopathies provide little insight to the function of TUBA1A in adult neurons. Several studies have investigated the impact of wild-type and mutant TUBA1A in vitro [13, 23-25, 302], but the sequence conservation between a-tubulin genes has historically made study of individual a-tubulin isotypes challenging in vivo. Thus, many questions remain about the role that TUBA1A plays in developing and adult neurons.

As discussed in Chapter I, TUBA1A reaches peak expression during embryogenesis in rodents. During brain development, Tuba1a constitutes 95% of all a-tubulin mRNA in the brain,

113 indicating that it is found abundantly in neuronal microtubules at this time [16]. My results,

discussed in Chapter III, show that Tuba1a comprises 95% of brain a-tubulin and 98% of cortical a-tubulin mRNA at birth in wild-type mice (Fig. 3.1). Thus, Tuba1a makes up nearly the entire pool of a-tubulin in developing neurons. Further, I showed that the Tuba1aND LOF allele

reduced total brain a-tubulin protein by half at birth in Tuba1aND/+ heterozygous mutant mice

compared to wild-type (Fig. 3.2). These data clearly show that Tuba1a is the major a-tubulin

isotype in the developing brain, in terms of both mRNA and protein abundance.

Tuba1a expression reduces approximately 20 fold between P3 and P23 in mice [16].

Despite this dramatic decrease, lower levels of Tuba1a expression persist in the adult brain, and

neurogenic regions like the cortex and hippocampus retain high levels of TUBA1A expression

into adulthood [27]. Additionally, recent data from both whole brain and isolated synaptosomes

showed that Tuba1a protein has a half-life greater than 7 days in rodents [127]. In Chapter III, I

show that Tuba1a comprises 32% of all a-tubulin mRNA in the 1-year-old adult rodent cortex

and 47% of all a-tubulin mRNA in the adult spinal cord (Fig. 3.1). Together, these data show

that Tuba1a constitutes nearly all the a-tubulin available to developing neurons, is not rapidly

turned over, and remains among the highest expressed a-tubulin isotypes in adult neurons. While

the a-tubulin isotype blend certainly changes dramatically between developing and adult

neurons, TUBA1A is a major component of neuronal microtubules throughout life.

TUBA1A Lays the Groundwork for life-long neuronal microtubule function

Continued incorporation of TUBA1A protein into adult neuronal microtubules likely

influences overall microtubule network properties and contributes to specialization of neuronal

microtubule function. Increased Tuba1a incorporation into microtubules was shown to slow

114 polymerization in vitro [13]. In Chapter II, my results show that the Tuba1aND LOF allele

reduces incorporation of TUBA1A into neuronal microtubules, resulting in accelerated

microtubule polymerization in cultured neurons (Figs. 2.3, 2.4). In Chapter III, I further show

that reducing neuronal Tuba1a, using the LOF Tuba1aND allele, reduced the density of

microtubules/µm of axon and impaired microtubule-based intracellular transport (Figs. 3.3, 3.4).

From these results, we can infer that TUBA1A helps to establish microtubule density, dictates

polymerization dynamics, and is necessary for axonal transport in developing neurons.

Of particular interest is the evident role for TUBA1A in facilitating intracellular transport.

Cortical neurons with reduced Tuba1a had impaired transport of both lysosomes and

mitochondria, caused by increased cargo stalling (Fig. 3.4). As developmental a-tubulin protein

is reduced by 50% in this model, we predict that Tuba1aND/+ neurons do not have enough microtubule “bricks” to build functional “roads” for transport. Microtubules in neurons are tiled in overlapping segments which collectively span the entire neuron [290]. We propose that reduced microtubule density in Tuba1aND/+ neurons leads to more gaps in the microtubule road,

which act as “potholes” causing motor proteins to stall and requiring them to change tracks.

Additional a-tubulin isotypes, such as TUBA4A, become upregulated in adult mammalian

neurons [198]. The increased expression of other a-tubulin isotypes indicates that adult neurons

contain a blend of a-tubulin isotypes rather than the TUBA1A-dominant condition seen in

developing neurons. In Chapter III I show that adult Tuba1aND/+ brain a-tubulin protein is

restored to wild-type levels, demonstrating that Tuba1a LOF causes an a-tubulin deficit during

brain development, but adult neurons have alternative a-tubulin isotypes available. Thus, adult

Tuba1aND/+ neurons have ample microtubule building blocks to repair the potholes in faulty

microtubule tracks. Despite the availability of adult a-tubulin protein, Tuba1aND LOF mutant

115 neurons have impaired intracellular transport, and ultimately fail to maintain proper synaptic connections, leading to behavioral deficits (Figs. 3.5, 3.6). Collectively, these data indicate that

TUBA1A a-tubulin is essential for establishing functional microtubule tracks within the developing neuron. The adult-onset synaptic and behavioral phenotypes in our Tuba1a LOF model demonstrate that poorly laid developing microtubule tracks cannot be adequately repaired to support neuron function later in life. We liken this situation to a road with potholes: attempts can be made to repair the potholes as they become problematic for traffic, but the road will never be as smooth as if it were built correctly in the first place. This model remains to be tested, but it is intriguing to consider that TUBA1A may be uniquely important for establishing neuronal microtubule architecture developmentally, that will be used by the neuron for its entire lifetime.

The precise mechanisms by which TUBA1A influences neuronal microtubule function remain to be tested, but my data support two possible hypotheses. 1) TUBA1A is highly expressed under its endogenous promoters, and could provide the “burst” of tubulin expression that neurons require to build complex microtubule structures during periods of intense growth or repair. The conserved nature of a-tubulin isotypes makes it unclear whether TUBA1A is functionally different from similar isotypes, like TUBA1B, or if TUBA1A supports growth by supplying neurons with abundant a-tubulin. My results show that the Tuba1aND LOF mutation reduced total a-tubulin protein by half in the developing brain, indicating that neurons lacking

Tuba1a may not have sufficient microtubules to support structurally-demanding feats like long- distance axon extension (Fig. 3.2). Compensation from other neuronally-expressed a-tubulin isotypes was not induced in the Tuba1aND LOF model, making it impossible to discern whether other a-tubulin isotypes could functionally replace Tuba1a in brain development (Fig. 3.2). A non-neuronal a-tubulin isotype, Tuba8, was not able to rescue neuronal migration deficits in

116 Tuba1a-S140G loss-of-function mutant mice [26] indicating that Tuba1a may play a unique role

in developing microtubules. 2) TUBA1A could confer specialized properties to neuronal

microtubule networks that promote establishment and maintenance of neuronal microtubule

function. As discussed in Chapter I, the tubulin code hypothesis proposes that genetically-

encoded differences between tubulin isotypes and PTMs to tubulin act together to modify

function of microtubules [119, 215]. TUBA1A can be tyrosinated/detyrosinated and

polyglutamylated on its CTT, potentially impacting microtubule dynamic properties and

interactions with MAPs [12, 78, 80, 94]. Polyglutamylation, in particular, only occurs on

TUBA1A and TUBA1B, and influences spastin-mediated microtubule severing, MAP-binding

affinity, and microtubule-based transport in neurons [91, 96, 246, 303]. Tubulin PTMs were not

significantly altered by reduced Tuba1a in adult Tuba1aND/+ brains (Fig. 3.2), but developing neuronal growth cones had altered distribution of acetylated microtubules (Fig. 2.6). Further examination is needed to determine if reduced TUBA1A significantly impacts the neuronal tubulin PTM landscape, but loss of TUBA1A could alter microtubule PTMs, impacting crucial protein interactions for neuronal development. Additionally, Tuba1a was shown to strongly impact microtubule dynamics, with increased Tuba1a incorporation associated with decreased microtubule polymerization velocity in vitro (Chapter II)[13]. Changes to microtubule polymerization directly impact the function of microtubule-based structures. In particular, microtubule dynamics have been shown to be involved in growth cone navigation, axon specification, synaptic remodeling and disease states like neurodegeneration [3, 35, 155, 162].

Importantly, these hypotheses are not mutually exclusive and likely both contribute to why

TUBA1A is uniquely important for neuron function.

117 Potential Roles for TUBA1A in Mature Neuronal Microtubules

My results show that Tuba1a is a major component of adult neuronal microtubule mRNA and illustrate that reduced Tuba1a impairs adult neuronal function in mice. These data provide the first evidence that Tuba1a influences adult microtubule function in vivo, but the specific requirements for TUBA1A in adult neurons remain to be explored. My data suggest that Tuba1a, and neuronal microtubules in general, participate in active maintenance of synaptic connections throughout life. Despite appearing morphologically normal at 1-month-old, 1-year-old

Tuba1aND/+ NMJs had diminished synaptic area and abnormal vesicle/receptor densities (Fig.

3.8). NMJ synapses are generally stable, but can undergo structural changes in response to increased or decreased stimulation [285, 295]. As such, the reduced synaptic area in adult

Tuba1aND/+ mice could indicate improper or loss of neuronal innervation, weak neurotransmission, or an inability of Tuba1aND/+ synapses to support structural plasticity.

Previous studies showed that dynamic microtubules participate in synaptic plasticity, potentially by facilitating structural changes to synapses [3-5]. Further, Tuba1a was shown to be enriched in mature rodent synapses [127]. My results illustrate that Tuba1a influences mature synapse morphology, poising TUBA1A to have an intriguing and novel role in regulating mature synapse function. Future studies will be needed to evaluate if TUBA1A-rich microtubules directly participate in structural plasticity of adult synapses, or if reduced TUBA1A impairs synaptic maintenance by other mechanisms.

In addition to altered synaptic area, Tuba1aND/+ mice also exhibited abnormal pre- and post-synaptic protein densities (Fig. 3.8). The accumulation of pre-synaptic vesicles and post- synaptic receptors in Tuba1aND/+ synapses suggests that reduced Tuba1a could impair synaptic protein delivery and clearance. Interactions between the microtubule cytoskeleton and the post-

118 synaptic membrane are necessary for targeted delivery of critical synaptic proteins, such as

acetylcholine receptors (AChRs) [304]. Post-synaptic AChR density at the NMJ is dictated by

the rate of membrane insertion and removal of AChRs. Turnover of membrane AChRs requires

“capture” of dynamic microtubule ends by agrin, a heparansulfate proteoglycan that is a

structural organizer in motor nerve terminals [305]. Specifically, microtubules become stabilized

at agrin-induced clusters of AChRs to facilitate focal delivery and removal of AChR subunits

[305]. My results show that reduced Tuba1a diminished axonal microtubule density (Fig. 3.3), thus it is possible that adult Tuba1aND/+ motor neurons do not contain enough microtubules to facilitate synaptic protein turnover. Further testing is needed to evaluate this hypothesis, but it illustrates one potential mechanism by which microtubule dysfunction could impact long-term synaptic health.

As discussed in Chapter I, mature neurons depend upon microtubules to facilitate intracellular transport of proteins, mRNAs, organelles and wastes. Breakdowns in intracellular transport, as the result of microtubule deficiency, would be detrimental to neuronal function. My results showed that developing cortical Tuba1aND/+ neurons have axonal transport deficits (Fig.

3.4). While it remains to be tested if trafficking deficits persist in adult Tuba1aND/+ neurons,

prolonged deficits in intracellular transport and clearance provide a compelling mechanism by

which microtubules could influence long-term synaptic function. Additionally, my results tie

intracellular transport deficits and microtubule dysfunction to adult-onset behavioral deficits.

Axonal transport deficits have been identified as a feature of numerous neurodegenerative

diseases, including Alzheimer’s disease, Parkinson’s disease, ALS, and Huntington’s disease,

among others [34, 39, 184, 196, 279, 280, 306]. It has thus far been difficult to establish if

intracellular transport deficits are a consequence of an already diseased neuronal environment, or

119 if microtubule impairments can induce neuronal dysfunction and degeneration. My results

illustrate that a microtubule deficit that impaired axonal transport was sufficient to alter nervous

system function, at the cellular and behavioral level (Chapter III). Additional studies are needed

to more fully explore the impact of the microtubule cytoskeleton on disease pathogenesis in the

nervous system, but my results indicate that faulty microtubules can be detrimental to neuron

function.

Multiple aspects of microtubule function are known to be altered in neurodegenerative

disease. Changes to microtubule dynamics, microtubule density, and intracellular transport are

all associated with neurodegeneration, but their role in the etiology of disease remains unclear.

Mutations to the microtubule severing enzyme, spastin, cause Hereditary Spastic Paraplegia in

humans, and caused acceleration of microtubule dynamics in neurons [187, 307, 308]. Dynamic

microtubules were depleted from neurons in an in vitro model of Parkinson’s disease, prior to the onset of other molecular pathology [39]. Regulation of dynamic and stable microtubule populations is clearly important for neuronal function, but it is unclear precisely how altered microtubule dynamics play into disease. In contrast, axonal transport deficits have been firmly established as a feature of nearly every neurodegenerative disease [34, 184]. Accumulation of cargoes is associated with axonal swellings and likely contributes to neuronal dysfunction in

Alzheimer’s, Parkinson’s, ALS, and multiple sclerosis [184, 280, 298]. Although neuronal microtubules are highly associated with neurodegenerative cellular pathology, the specific a- and b-tubulin isotypes that are impacted by disease are largely unknown. My results provide evidence that Tuba1a is likely to be abundant in adult neuronal microtubules, and show that

reduced Tuba1a function is sufficient to impair synaptic morphology and cause behavioral

deficits in adult mice (Chapter III). These data suggest that TUBA1A is poised to participate in

120 adult neuron function, and illustrates that reduced TUBA1A replicates aspects of

neurodegenerative disease. These findings raise the possibility that diverse genetic and

environmental causes of adult nervous system disease may share convergent microtubule-based

mechanisms of cellular and molecular dysfunction.

Microtubule fulfill diverse functions in neurons, and as such there are numerous ways in

which microtubule deficits could impact neuronal function. To understand how microtubules can

become dysregulated in neurological disease, we must first understand their contributions to

normal neuronal function. Here I provide direct evidence that neuronal microtubules, in

particular TUBA1A, are required for neuron growth, axon targeting, synaptic maintenance/remodeling, and intracellular transport (Chapters II and III). Using the Tuba1aND allele, I explored how reducing TUBA1A impacted neuronal development and mature neuron function. However, it remains unclear whether microtubule LOF contributes significantly to the pathology of neurological diseases, such as neurodegeneration or tubulinopathy, or if gain-of- function mutations are larger contributors to disease. For many years the predominant hypothesis of tubulinopathy pathogenesis has been haploinsufficiency. In the following sections, I explore how tubulin LOF impacts nervous system function, and provide evidence that dominant, function-disrupting tubulin mutations are likely larger contributors to disease.

Haploinsufficiency vs. Dominant Negative Mechanisms of Disease

Other Tubulin Isotypes Provide Insight into Mechanism of Disease

Insight into the consequence of tubulin deficiency during brain development can be

ascertained by examining mutations in other tubulin genes. The strongest evidence that dominant

missense mutations in tubulin lead to stronger brain development phenotypes than isotype loss

121 comes from studies of the b-tubulin isotype Tubb3 in mouse models. Human TUBB3 mutations have been associated with a spectrum of developmental brain disorders and progressive neuropathy, and share many clinical features with TUBA1A-associated tubulinopathy [209].

Greater flexibility afforded by the increased sequence divergence of β-tubulin isotypes has allowed researchers to generate Tubb3 knock-in mouse models of patient TUBB3 mutations as well as generate a full Tubb3 knock-out mouse [118, 309, 310]. Tubb3-R262C homozygous knock-in mice exhibit lower Tubb3 protein levels, reduced heterodimer levels, altered microtubule stability, and decreased interactions with the kinesin motor Kif21a, resulting in axon guidance defects in vivo [309]. Deficits in guidance of commissural, corticospinal, and optic nerve axons are commonly associated with TUBB3 mutations in humans. The precise mechanisms by which TUBB3, a β-tubulin that is ubiquitously expressed in neurons, causes axon guidance deficits only in certain brain regions has been unclear. Recently, Tubb3-R262C was shown to prohibit interaction between TUBB3 and the receptor DCC, which is critical for axon guidance by Netrin signaling [311]. This evidence further indicates that the Tubb3-R262C variant acts dominantly to impair axon guidance by impeding microtubule function downstream of key developmental signaling pathways. These data collectively suggest that the Tubb3-R262C mutant may act to disrupt tubulin function through a combination of defects, including reduced heterodimer (also predicted for the Tuba1a mutant mice), changes to microtubule dynamics, and disruptions to specific MAP-binding and signaling interactions. When the disease-associated mutations in TUBB3 were modeled in yeast β-tubulin, all mutations (R62Q, A302T, R380C,

R262C, R262H, E410K, D417H, and D417N) were capable of polymerizing into microtubules, where they disrupted microtubule dynamics [309]. Additionally, a subset of the mutants was

122 shown to potently disrupt kinesin motors in yeast, mirroring the R262C mouse Kif21a phenotype

[309].

In contrast to the knock-in Tubb3 mutant mouse model, the Tubb3 null mouse exhibited completely normal development and behavior, with only a slight defect in mature axon regeneration [118]. Here, with a systemic loss of the single tubulin isotype, it was possible to study how losing Tubb3 impacts cellular β-tubulin isotype expression. Intriguingly, equivalent total β-tubulin mRNA and protein levels were observed in Tubb3 null mice [118]. Complete

deletion of Tubb3, long thought to be a critical neuronal β-tubulin isotype, was readily

compensated for via upregulation of the remaining neuronal β-tubulin isotypes. This result shows

that neurons are capable of upregulating other β-tubulins to compensate for the loss of Tubb3 and

did not rely on a single isotype to replace the needed β-tubulin. The mechanism by which the cell

senses the tubulin pool and compensates by regulating isotype expression is an outstanding

question in the field. This study also demonstrates directly that neurons can indeed tolerate the

loss of a neuronally expressed β-tubulin isotype without measurable defects in

neurodevelopment. This finding is intriguing in light of the severe consequences of heterozygous

TUBB3 and TUBA1A mutations both in humans and mice, and calls to question whether

complete loss of one copy could be compensated for by upregulating other tubulin isotypes, or if

there is a critical distinction between partial and full loss-of-function.

While most of the disease-associated mutations to α-tubulin occur in TUBA1A, a minor,

neuronally expressed α-tubulin isotype, TUBA4A, has also been linked to neuronal disease.

Interestingly, TUBA4A variants are not associated with cortical malformations, but have been

identified in humans with adult-onset neurodegenerative diseases such as Fronto-Temporal

Dementia (FTD) and familial and sporadic forms of ALS [198, 300, 301, 312, 313]. This delayed

123 onset of disease is likely due to TUBA4A’s expression level dramatically increasing 50-fold post-

development [198]. Three familial ALS missense mutations, TUBA4A-A383T and TUBA4A-

R320C/H, dominantly disrupt microtubule dynamics and stability in cells, supporting the

dominant-negative hypothesis for disease-causing tubulin mutations [198]. Unlike the

universally missense TUBA1A patient mutations, however, the cohort of ALS-associated

TUBA4A variants also encompasses several truncation mutations, which likely act as loss-of-

function alleles. Specifically, while the majority of ALS or FTD-associated variants are missense, the cohort also includes one nonsense mutation, one donor splice-site variant, and one frame-shift variant [198, 300, 313]. The truncated TUBA4A-W407X protein was unable to form tubulin dimers or incorporate effectively into microtubules in vitro, demonstrating a clear LOF mechanism [198]. While these mutants are predicted to cause truncated, nonfunctional TUBA4A protein, it is unclear whether they can also act in a dominant negative manner or whether reduced availability of TUBA4A is sufficient to cause neurodegenerative defects. It is important to note that expression profiles of TUBA1A and TUBA4A vary significantly and peak expression does not overlap, with TUBA1A representing the majority of α-tubulin developmentally while

TUBA4A expression increases dramatically with age [16, 198].

Examining how tubulin mutations impact neuronal function throughout life provides mechanistic insight into the functional requirements for tubulin isotypes and reveals tubulin-

MAP interactions that govern intracellular processes. The variants described above illustrate how tubulin loss-of-function and gain-of-function can play out within the context of the nervous system. Haploinsufficiency and dominant negative mechanisms likely both contribute to tubulin- related neurological disease. Elucidating how microtubules become dysregulated in disease states can provide important mechanistic insights that advance scientific understanding of

124 disease pathology. My work with the Tuba1aND LOF mouse model highlights requirements for

TUBA1A microtubules throughout nervous system development, and provides direct evidence

that TUBA1A haploinsufficiency does not cause severe brain malformations, but has a long-

lasting impact on neuronal function.

Tuba1aND Mouse Model Illustrates Consequences of TUBA1A LOF

Tuba1aND is a true LOF allele

Examining the requirements for TUBA1A in the developing and mature nervous system has been historically difficult due to the conserved nature of tubulin isotypes. Although many

TUBA1A mutations are associated with human tubulinopathy, only a handful of studies have examined molecular mechanisms by which these variants disrupt brain development. In this work, I used the Tuba1aND allele to interrogate the consequences of Tuba1a deficiency in the

developing and mature nervous system. The Tuba1aN102D allele was identified through a forward-

genetic ENU-induced mutagenesis screen in mice [183]. The N102D variant was predicted to

weaken heterodimer interaction by computational modeling and impaired microtubule-dependent

mitotic division in yeast, predicting it to be a LOF allele. Despite indications that Tuba1aND

likely impaired Tuba1a function, the lack of tools to examine Tuba1a protein made it difficult to

determine how the N102D mutant protein behaved in vivo.

My results elucidated molecular mechanisms by which Tuba1aND substitution causes

LOF in several ways. First, I demonstrated that Tuba1aND/+ heterozygous mice have an

approximate 50% reduction in total a-tubulin protein, at a developmental timepoint when

Tuba1a constitutes >98% of all a-tubulin mRNA in the brain (Fig. 3.1). Though Tuba1a protein

specifically could not be evaluated by western blot, loss of half of all brain a-tubulin in a

125 heterozygous model is consistent with LOF. Next, to specifically interrogate Tuba1a tubulin in the Tuba1aND model, I utilized a novel TUBA1A-His6 tagging system developed by Dr. Jayne

Aiken (University of Colorado Anschutz). His-tagged wild-type TUBA1A polymerized into cellular microtubules and was abundantly visible in all compartments of the neuron, including distal structures like the growth cone (Fig. 2.4). In contrast, His-tagged Tuba1aND tubulin was not detectable in either Cos-7 cells or in rat cortical neurons (Fig. 2.3, 2.4). Mutant Tuba1aND protein was not incorporated into cellular microtubules, despite similar mRNA expression of

TUBA1AND and wild-type TUBA1A in cells, further supporting a LOF-role for this variant.

Finally, I inhibited proteasome function in cells expressing Tuba1aND mutant protein to evaluate

potential mechanisms of degradation. My results indicate that Tuba1aND mutant protein is not

targeted for degradation by the proteasome, suggesting that mutant Tuba1aND protein may

become trapped in the tubulin folding pathway (Fig. 2.3). Collectively, my results conclusively

show that Tuba1aND is a LOF allele which leads to dramatic loss of TUBA1A protein from

neurons.

As the Tuba1aND allele causes complete LOF in Tuba1a, Tuba1aND/+ heterozygous

mutant mice can be used to investigate the consequences of Tuba1a haploinsufficiency for

neurodevelopment. Importantly, other brain-expressed a-tubulin isotypes were not upregulated

to compensated for reduced Tuba1a in Tuba1aND mice (Fig. 3.2). This marks a significant

distinction between the Tuba1aND allele and the previously described Tubb3 null mouse. Tubb3 null mice showed increased expression of other b-tubulin isotypes, causing the total amount of b-tubulin protein to remain consistent between Tubb3 null mouse brains and controls [118]. In contrast, Tuba1aND/+ mice had half as much brain a-tubulin protein as controls at birth (Fig. 3.2).

126 For this reason, we assert that the Tuba1aND/+ heterozygous mouse model represents a true condition of Tuba1a haploinsufficiency during the critical period of brain development.

Developmental TUBA1A Haploinsufficiency does not replicate tubulinopathy

Having established that Tuba1aND/+ mice experience developmental Tuba1a haploinsufficiency, much can be inferred about the requirements for Tuba1a during brain development from my results. Cortical malformations, like lissencephaly, are a common feature of human tubulinopathies, and can be caused by cortical neuron migration errors [19-22, 25, 180,

181]. Initial models of disease presumed that patients harboring heterozygous TUBA1A mutations did not have sufficient brain a-tubulin to support complex neurodevelopmental processes like cortical migration. As discussed in Chapter I, cortical neuron migration is a taxing feat that is highly dependent on microtubule-MAP interactions to generate motile force. My results directly contest the haploinsufficiency hypothesis of tubulinopathy by illustrating that developmental Tuba1a haploinsufficiency was sufficient to support normal cortical migration in

Tuba1aND/+ mice (Fig. 3.6). However, reduced developmental Tuba1a was detrimental to long- range axon pathfinding and precluded formation of forebrain commissures (Figs. 2.1, 2.2).

Human tubulinopathy patients commonly exhibit abnormal formation of brain commissures [15,

17-22, 178, 179, 181], but it has as yet been unknown whether commissural deficits occur as the result of tubulin dysfunction directly or if they are secondary to other brain malformations.

Pathfinding axons may be exquisitely sensitive to microtubule deficits, as they rely heavily on microtubules to drive neuronal growth and to dynamically respond to the extracellular environment to navigate towards correct synaptic targets. Commissural axons, in particular, extend across vast distances to reach contralateral synaptic targets and must correctly respond to

127 numerous guidance cues along their route, requiring precise control of microtubule function. My

results indicate that cortical migration and axon guidance/commissural formation differentially

require TUBA1A. Specifically, my evidence shows that TUBA1A is essential for commissural

axon pathfinding, but full TUBA1A function is not required for cortical migration (Chapters II

and III). These results highlight the intriguing possibility that dominant microtubule “poisoning”

variants and reduced tubulin function may both play a role in the disease pathology in human

tubulinopathies. Overall, my work with the Tuba1aND model illustrates that TUBA1A

haploinsufficiency does not replicate human tubulinopathy pathology.

Concluding Remarks

This work provides the first direct evidence that TUBA1A haploinsufficiency does not

explain the pathology of human tubulinopathies. If haploinsufficiency was a major contributor to

the neurodevelopmental defects observed in TUBA1A-associated tubulinopathy, one would expect

nonsense or frameshift mutations, which lead to truncated, nonfunctional protein, to be identified

in diagnosed patients. As discussed previously, all of the currently identified TUBA1A

tubulinopathy patients harbor missense mutations in which one amino acid is altered. The absence

of deletions, frameshifts, and truncations combined with viability of heterozygous null mouse

mutants in Tuba1a suggest that heterozygous tubulin LOF may be sufficient to support cortical migration and gross brain development. My results with the Tuba1aND mouse model further

illustrate that cortical migration, the primary feature of human tubulinopathy, occurs normally

under conditions of TUBA1A haploinsufficiency (Chapter III). Drawing from these observations

and the data presented above on tubulin mutation mouse models, we propose that TUBA1A disease-

associated mutants act in a dominant fashion, in which they are able to join the polymerization-

128 competent pool of tubulin heterodimers and then “poison” the microtubule networks from within.

This mechanism of action could potentially escape detection and upregulation of alternative isotypes and thus wreak havoc in the microtubule network in a dominant manner. A dominant negative mechanistic model presents the exciting possibility that tubulinopathy associated mutations could be used to reveal molecular interactions with microtubules that are important for brain development.

The work described in this dissertation also highlights a novel role for TUBA1A in the mature nervous system. Human tubulinopathy mutations are often lethal early in life, and as such have only been associated with neurodevelopmental phenotypes. The severity of clinical neurodevelopmental phenotypes in tubulinopathy patients makes it impossible to determine how or if tubulinopathy-associated variants would impact mature neuron function. As such, the

Tuba1aND mouse model is an excellent tool to interrogate the requirements for TUBA1A in developing and mature neurons. My results suggest that mature neurons require TUBA1A, and microtubules in general, for maintenance of synaptic morphology and adult neuronal function.

Specifically, I provide direct evidence that microtubule dysfunction is sufficient to cause adult- onset behavioral deficits, without incurring neuronal death (Chapter III). Microtubule dysfunction has been identified as a feature of numerous neurodegenerative diseases in humans, and my results support the idea that microtubule dysfunction can potentially drive pathology in these diseases.

Collectively, these findings expand upon the known roles of TUBA1A in neurodevelopment, and illustrate that TUBA1A is uniquely poised to be an important tubulin isotype for neuronal function throughout life.

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