Nervous System Compensation Following Tail Loss and Regeneration in the Leopard

Home , Neuromorphology

Gecko (Eublepharis macularius)

Stefanie Simone Bradley

A Thesis

presented to

the University of Guelph

In partial fulfillment of the requirements

for the degree of

Master of Science

Biomedical Science

Guelph, Ontario, Canada

ABSTRACT

NERVOUS SYSTEM COMPENSATION FOLLOWING TAIL LOSS AND REGENERATION IN THE LEOPARD GECKO (EUBLEPHARIS MACULARIUS)

Stefanie Bradley Advisor: University of Guelph, 2018 Dr. M.K. Vickaryous

Mass change is a physical phenomenon with important implications for biomechanics and locomotion. Here, we used the leopard gecko (Eublepharis macularius) to investigate the effect of a drastic change in mass following tail loss (autotomy), and subsequent regeneration of the tail. We assessed two components of the nervous system: tactile sensitivity, and Purkinje cell neuromorphology. Using Semmes-Weinstein monofilaments, we found regional differences in tactile sensitivity prior to autotomy. Following tail autotomy, the hindlimbs became significantly more sensitive, while the forelimbs did not. Golgi-Cox staining of Purkinje cells showed that tail autotomy had no significant effect on Purkinje cell structure. However, after 30 days of tail regeneration, there was evidence of dendritic remodeling corresponding to the interval where parallel fibers synapse with Purkinje cell dendrites. Together, these data provide support for short-term (transient) compensation of the peripheral nervous system, and long-term compensation of the central nervous system, in geckos following autotomy.

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ACKNOWLEDGEMENTS

Time to acknowledge some cool people!

First and foremost – Matt – for being an incredible P.I. A major reason I joined the lab was because of your friendly demeanor (as well as super cool science, of course). As an advisor, you’re especially hands-on, but also allow for independence in our research goals. I really appreciated your support when I wanted to branch into somewhat different territory for my project (i.e., poking geckos with nylon filaments). As a result of my time in the lab, I feel much more confident as a researcher and scientist. I also feel much more confident in assessing how

“peaty” different whiskeys are (mostly so I can shoot them anyways). Whatever life skills you’re bestowing on us, the net result of your leadership is a unique and motivating environment in the lab.

To Matt’s family - Jalene, for always being a gracious host, and always welcoming members of the lab with open arms (and amazing food). To honorary members of the lab, Isla and Piper, for their energy and gecko-centric artwork, which usually accompanies every lab milestone.

Next – to the Eublephosphere. To Sarah, Rebecca, and Kathy who have been there since the beginning of my Master’s. From practice talks, to conferences, to bar nights - the dynamic in our lab is fun, scientifically collaborative, and genuinely passionate. To newer members of the lab:

Laura and Yifan (also for helping with neuron tracings), and undergrad students as well.

To my advisory committee - Craig and Leah. It was motivating to have a committee that was genuinely interested in the work that was being done. I think I lucked out in that you guys were actively involved in my project, always approachable, and always helpful in providing valuable feedback and hands-on expertise. I’d also like to thank my examination committee, Dr.

MacLusky and Dr. LaMarre, for agreeing to participate in my defense on an end-of-semester

Friday afternoon (ideal for celebrating but not ideal for everyone else).

To Erika Howe from the Bent lab – my gargantuan monofilament data set may still remain a mystery if it had not been for you. I’m grateful for the many hours you spent helping me with statistical planning and analysis.

To my parents - for being supportive of their offspring pursuing science instead of business. To my mom for listening to me recite my seminars approximately one million times, and always being a support system. To my dad for actually asking questions at my defense. To my brother

Michael for being mildly interested in my research, and to my smaller brother Christopher for thinking stem cells and regenerative biology are the coolest things ever.

To my friends – for listening to me talk about thesis writing for many months, and for putting up with me continually re-scheduling my defense date (and then still taking the day off work to make it to my defense).

To the geckos and mice – for being super cute model organisms (besides the biting) and contributing to science.

To the biomedical department at UofG – for providing a warm sense of community, and always being friendly. I’d also like to acknowledge Starbucks for providing endless amounts of caffeine and a makeshift office for many months of thesis writing.

My Master’s project was definitely a collaborative effort. It allowed me to take science from a hobby/interest, to a field that I will most likely remain in for my career. If you’d asked me a few years ago where I’d be now, I don’t think I would ever have guessed this - so that’s pretty cool.

TABLE OF CONTENTS

ABSTRACT ...... ii

ACKNOWLEDGEMENTS ...... iii

TABLE OF CONTENTS ...... iv

LIST OF FIGURES ...... viii

LIST OF TABLES ...... xi

LIST OF APPENDICES ...... xiii

LIST OF ABBREVIATIONS ...... xiv

DECLARATION OF WORK PERFORMED ...... xv

CHAPTER 1: LITERATURE REVIEW ...... 1

1.1 Mass Change in Nature ...... 1

1.2 Somatosensation ...... 3

1.3 The Cerebellum ...... 7

1.4 Dendritic Morphology ...... 11

1.5 Appendage Loss and the Leopard Gecko ...... 15

CHAPTER 1 FIGURES...... 19

RATIONALE ...... 21

CHAPTER 2: SOMATOSENATION IN THE PNS: CHARACTERIZATION OF TACTILE SENSITIVITY FOLLOWING TAIL LOSS AND REGENERATION IN THE LEOPARD GECKO (EUBLEPHARIS MACULARIUS)...... 23 2.1 INTRODUCTION ...... 23

2.2 MATERIALS AND METHODS...... 28

2.2.1 Experimental Animals and Animal Care...... 28

2.2.2 Experimental Design ...... 29

2.2.3 Tail Autotomy ...... 29

2.2.4 Monofilament Testing ...... 30

2.2.5 Euthanasia, Tissue Collection, and Preparation...... 31

2.2.6. Statistical Analysis...... 32

2.3 RESULTS ...... 33

2.3.1 Experimental Geckos ...... 33

2.3.2 Monofilament Testing ...... 34

2.3.3 Regional Differences Prior to Autotomy ...... 35

2.3.4 Sensitization of Control Group ...... 35

2.3.5 Hindlimb Sensitivity with Tail Loss and Regeneration ...... 36

2.3.6 Forelimb Sensitivity with Tail Loss and Regeneration ...... 37

2.3.7 Tail Base Sensitivity with Tail Loss and Regeneration ...... 37

2.3.8 Tail Tip Sensitivity with Tail Loss and Regeneration ...... 38

2.4 DISCUSSION ...... 39

2.4.1 Regional differences in tactile sensitivity exist prior to autotomy ...... 40

2.4.2 Monofilament testing results in sensitization……………………………...... 42

2.4.3 Sensitization is attenuated in the forelimbs and exaggerated in the hindlimbs, following tail loss ...... 44

2.4.4 Tactile sensitivity is variable in the tail base, and unchanged in the tail tip, following tail loss ...... 46

2.4.5 Differential changes in tactile sensitivity may be attributed to selective weighting of cutaneous feedback in the CNS, or altered skin capillary permeability ...... 47

2.4.6 Future directions...... 49

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CHAPTER 2 TABLES ...... 50

CHAPTER 2 FIGURES ...... 52

CHAPTER 3: NEUROMORPHOLOGY IN THE CNS: CHARACTERIZATION OF PURKINJE CELL MORPHOLOGY FOLLOWING TAIL LOSS AND REGENERATION IN THE LEOPARD GECKO (EUBLEPHARIS MACULARIUS) ...... 64

3.1 INTRODUCTION ...... 64

3.2 MATERIALS AND METHODS...... 69

3.2.1 Experimental Animals and Animal Care...... 69

3.2.2 Experimental Design ...... 70

3.2.3 Tail Autotomy ...... 71

3.2.4 Euthanasia, Tissue Collection, and Preparation ...... 71

3.2.5 Golgi-Cox Impregnation ...... 72

3.2.6 Golgi-Cox Sectioning ...... 72

3.2.7 Golgi-Cox Section Processing ...... 73

3.2.8 Golgi-Cox Mounting and Dehydrating Brain Sections...... 73

3.2.9 Microscopy and Imaging Stained Neurons...... 74

3.2.10 Histology: Hematoxylin and Eosin...... 74

3.2.11 Immunofluorescence...... 75

3.2.12 Sholl and Branch Analyses……………………………………………………….76

3.2.13 Statistical analysis...... 77

3.3 RESULTS ...... 77

3.3.1 Experimental Geckos...... 77

3.3.2 Organization of the Cerebellar cortex ...... 78

3.3.3 Purkinje Cell Visualization ...... 79

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3.3.4 Species Differences in Purkinje Cell Neuromorphology...... 79

3.3.5 Purkinje Cell Neuromorphology with Tail Loss and Regeneration...... 80

3.4 DISCUSSION ...... 82

3.4.1 A phylogenetic trend is apparent in the dendritic complexity of Purkinje cells…...83

3.4.2 Tail loss and regeneration does not significantly affect Purkinje cell

neuromorphology ...... 84

3.4.3 Purkinje cells show dendritic remodeling at specific intervals following tail

regeneration...... 85

3.4.4 Purkinje cell morphology does not change shortly after tail loss……………….... 88

3.4.5 Future directions...... 89

CHAPTER 3 TABLES ...... 91

CHAPTER 3 FIGURES ...... 92

CHAPTER 4: CONCLUDING STATEMENTS AND FUTURE DIRECTIONS ...... 103

4.1 Is post-autotomy compensation occurring in the PNS of the leopard gecko?...... 103

4.2 Is post-autotomy compensation occurring in the CNS of the leopard gecko?...... 105

4.3 Integration of the CNS and PNS in the leopard gecko ...... 106

CHAPTER 4 TABLES …………………………………………………………………....… 109

REFERENCES ...... 110

APPENDIX I: EXPERIMENTAL GROUPS...... 133

APPENDIX II: DETAILED HISTOCHEMICAL AND IMMUNOHISTOCHEMICAL PROTOCOLS AND RECIPES...... 136

APPENDIX III: RAW MONOFILAMENT DATA...... 140

APPENDIX IV: TWO-DIMENSIONAL NEURON TRACINGS ...... 143

APPENDIX V: STATISTICAL OUTPUT ...... 144

LIST OF FIGURES

CHAPTER ONE

Figure 1.1

A schematic representation of central and peripheral nervous system integration with a focus on the coordination of movement

Figure 1.2

Synaptic connections of the cerebellar cortices

CHAPTER TWO

Figure 2.1

Schematic illustration of experimental design for monofilament assay

Figure 2.2

Locations of the sites of interest for monofilament testing

Figure 2.3

Monofilament testing chamber and a representative Semmes-Weinstein monofilament

Figure 2.4

Ascending stepwise monofilament method

Figure 2.5

Regional sensitivity differences at baseline

Figure 2.6

Changes in hindlimb sensitivity with respect to baseline following tail loss and tail regeneration

Figure 2.7

Changes in forelimb sensitivity with respect to baseline following tail loss and tail regeneration

Figure 2.8

Changes in tail base sensitivity with respect to baseline following tail loss and tail regeneration

Figure 2.9

Changes in tail tip sensitivity with respect to baseline following tail loss and tail regeneration

CHAPTER THREE

Figure 3.1

Schematic illustration of experimental design for Golgi-Cox Procedure

Figure 3.2

Gross anatomy of the gecko brain

Figure 3.3

Histology of the gecko cerebellum

Figure 3.4

Cortical layers of the gecko cerebellum

Figure 3.5

Golgi-Cox stained cerebella of the gecko and mouse

Figure 3.6

Two-dimensional tracings of gecko and mouse Purkinje cells

Figure 3.7

Morphological analysis of gecko Purkinje cell dendrites

Figure 3.8

Branch analyses of gecko Purkinje cell dendrites

Figure 3.9

Schematic illustration mapping Purkinje cell neuromorphology onto a phylogeny of vertebrates

APPENDIX IV

Figure A1

Two-dimensional Purkinje cell tracings

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LIST OF TABLES

CHAPTER TWO

Table 2.1

Monofilament buckling force values

Table 2.2

Absolute and normalized monofilament threshold values relative to baseline

CHAPTER THREE

Table 3.1

Quantification of mouse and gecko Purkinje cell neuromorphological characteristics

CHAPTER FOUR

Table 4.1

Properties of tail loss and tail regeneration in the leopard gecko

APPENDIX I

Table A1

Experimental group data at start of experiment

Table A2

Experimental group data at euthanasia

Table A3

Autotomy data, tail loss group

Table A4

Autotomy data, tail regeneration group

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APPENDIX III

Table A5

Raw monofilament data for control group

Table A6

Raw monofilament data for tail regeneration group

Table A7

Raw monofilament data for tail loss group

APPENDIX IV

Table A8

Number of neurons (‘n’) averaged for each gecko and mouse

APPENDIX V

Table A9

Three-way ANOVA output for all monofilament data (normalized to baselines)

Table A10

Two-way ANOVA output for Sholl analyses of Purkinje cell intersections and diameter

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LIST OF APPENDICES

APPENDIX I

Experimental groups and timelines

APPENDIX II

Detailed histochemical and immunohistochemical protocols and recipes

APPENDIX III

Monofilament output

APPENDIX IV

Two-dimensional neuron tracings

APPENDIX V

Statistical output

LIST OF ABBREVIATIONS

˚C degrees Celsius

CNS central nervous system

CoM center of mass

DAPI 4’,6-diamidino-2-phenylindole dH2O deionized water

GABA gamma-Aminobutyric acid

GRF ground reaction force

H&E hematoxylin and eosin

IF immunofluorescence

LSD least significant difference

MF monofilament

NBF neutral buffered formalin

PBS phosphate buffered saline

PNS peripheral nervous system

RT room temperature

SVL snout-vent length

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DECLARATION OF WORK PERFORMED

I declare that I performed all work demonstrated in this thesis with the exception of the following items: Dr. Matt Vickaryous assisted with gecko autotomy. Kathy Jacyniak, Sarah

Donato, and Rebecca McDonald assisted with perfusions of geckos for histology and immunofluorescence. Dr. Craig Bailey assisted with mouse euthanasia and Golgi-Cox protocol.

Yifan Liu assisted with some control gecko and mouse Purkinje cell tracings. Erika Howe assisted with monofilament statistical planning and analysis.

CHAPTER 1: LITERATURE REVIEW

1.1 Mass Change In Nature Body mass is a key determinant of organismal structure and morphology, ecology, physiology, and locomotor abilities (Farley and Ferris, 1998; Ogamba et al., 2016; Jagnandan and Higham, 2018). During evolution and development, gradual changes in body mass impact virtually every organ system, resulting in profound alteration of tissue composition, the organization of gas exchange surfaces, and patterns of innervation (Farebrother et al., 1974;

Gomez et al., 2007; Jagnandan and Higham, 2018). However, not all changes in mass are gradual. Many species experience rapid changes, occurring on a scale of seconds or less.

Furthermore, these changes may be transient (such as bearing an external load) or permanent

(such as amputation of a limb). Regardless of their origin, a rapid change in body mass can have a significant effect on how the organism interacts with its environment. One key aspect of the interaction is how the overall mass of the organism is distributed, or its center of mass (CoM).

Altered CoM occurs when animals bear extrinsic loads, such as a horse supporting its rider, or when there is an intrinsic change resulting from a natural process such as gravidity, egg laying (oviposition), or appendage loss (Weishaupt et al., 2004; Jagnandan and Higham, 2018).

Mass change confined to a specific area of the body can significantly alter the distribution of weight within an organism, thus changing vertical force distribution and the contributions of each limb during locomotion (Besancon et al., 2004; Browning et al., 2007; Bockstahler et al.,

2016). Quadruped or biped motion causes fluctuating patterns in CoM, and therefore to maintain stability and locomotor performance; compensatory changes in gait, posture, and balance are required (Farley and Ferris, 1998; Zehr and Stein, 1999). This is seen in humans during later stages of pregnancy, where an increase in anterior mass (and subsequent anterior shift in CoM)

results in kinematic changes at the knee, hip, pelvis, and trunk to reduce the effect of the added load (Ogamba et al., 2016). In another example, when bearing extrinsic loads box turtles compensate by modifying their gait to take shorter strides (Marvin and Lutterschmidt, 1997).

Changes in posture also result in altered CoM (Jagnandan et al., 2014). While these adjustments are adaptive to intrinsic and extrinsic CoM changes, they are also employed when responding to obstacles in the external environment, such as uneven ground (Riemann and Lephart, 2002), and are essential for long-term survival and fitness (Irschick and Garland, 2001). These adjustments are facilitated by neuromuscular shifts – modulated by the central and peripheral nervous system; an integrative process of neural signaling to modify gait and locomotion (Chiel et al., 1997;

Baloh et al., 1998; Hoogland et al., 2015) (Figure 1.1).

Historically, the nervous system was thought to be non-dynamic (Butz et al., 2009).

However, it is now well understood that the nervous system of vertebrates is capable of plastic remodeling to compensate for intrinsic (such as CoM) and extrinsic (environmental) changes

(Irschick and Garland, 2001; Navarro et al., 2005; Butz et al., 2009). This is an integrative process, combining changes in sensory input and shifting neural signaling to regulate neuromuscular output (Riemann and Lephart, 2002; Jagnandan and Higham, 2018). Motor plasticity involves efferent signals going from the central nervous system (CNS) to the peripheral nervous system (PNS) (Irschick et al., 2001; Navarro et al., 2005), and is modulated in part by reciprocal sensory feedback providing information about the extrinsic environment (Chiel et al.,

1997; Navarro et al., Mugge et al., 2009; Bent and Lowry, 2013). Sensory feedback incorporates inputs from various sources, including somatosensory (proprioceptive, joint, and cutaneous), visual, and vestibular afferents, typically in combination (Riemann and Lephart, 2002; Mugge et al., 2009). For example, environmental obstacles may be evaluated using visual and tactile

(mechanoreceptor) detection, resulting in corrections to motor output to prevent destabilization

(Riemann and Lephart, 2002; Roudaut et al., 2012; Requarth and Sawtell, 2014). Achieving and maintaining biomechanical stability involves an integrated interaction between both neuromuscular and musculoskeletal systems across multiple body regions (Farley and Ferris,

1998; Jagnandan and Higham, 2018). This motor-sensory loop is essential to maintaining motor plasticity and behavioral adaptation, and involves a range of molecular, structural, and functional changes in the nervous system (Shumway-Cook and Horak, 1986, Wang and Lin, 2008;

Takakusaki, 2017). Here we focus on two main components: the somatosensory system of the

PNS and the cerebellum of the CNS.

1.2 Somatosensation Adaptability in posture-gait control, body orientation, and verticality, is achieved using afferent multi-sensory sources, including vestibular, visual and somatosensory inputs (Meyer et al, 2004; Carver et al., 2006; Horak, 2010). In bipeds and quadrupeds, the most commonly employed sensory input for the control of balance is somatosensory information from the extremities in contact with the external environment (Shumway-Cook and Horak, 1986; Quai et al., 2004). The somatosensory system provides tactile feedback, and is associated with the conscious perception of touch, pressure, pain, temperature, position, movement, and vibration

(Reed-Geaghan and Maricich, 2011; Roudaut et al., 2012; Bent and Lowrey, 2013). These sensations are detected by a variety of receptors, including mechano-, proprio-, thermo-, and nociceptors, which then relay signals to the CNS (Navarro et al, 2005; Reed-Geaghan and

Maricich, 2011).

Cutaneous mechanoreceptors innervate the skin and relay information about touch and pressure through sensory afferents (or nerves) to the somatosensory cortex of the brain (Johnson,

2001; Reed-Geaghan and Maricich, 2011). The term “cutaneous afferent” is often used to denote the combination of the mechanoreceptor ending and the sensory nerve (Ergen and Ulkar, 2007).

In terms of structure, mechanoreceptors are terminations of sensory neurons, with their cell somas located either in the trigeminal ganglia (innervating the head) or dorsal root ganglia adjacent to the vertebral column (innervating the body) (Johnson, 2001). Mechanoreceptors may be slowly (i.e., they respond for full duration of stimulation) or rapidly adapting (i.e. they only fire at the beginning and end of stimulation), and may be encapsulated (i.e., dendrites contained within a capsule) or unencapsulated (free nerve endings) (Knibestol and Vallbo, 1970). Human tactile mechanoreceptors include: Merkel’s disks (slowly adapting, unencapsulated), Meissner’s corpuscles (rapidly adapting, encapsulated), Ruffini endings (slowly adapting, encapsulated), and

Pacinian Corpuscles (rapidly adapting, encapsulated) – all found at different depths within the skin (Macefield, 2005). Mechanoreceptors can be further categorized based on their receptive field (stimulus region) characteristics, and ability to respond to different mechanical stimuli

(Knibestol and Vallbo, 1970). For example, the human fingertip has a high density of mechanoreceptors with small receptive fields, and thus can detect more precise detail than areas containing receptors with large receptive fields (Macefield, 2005). Sensation is initiated via mechanical deformation of the skin, as a result of stretch, vibration, or pressure (Johnson, 2001).

This deformation then stimulates stretch-sensitive ion channels in the mechanoreceptor endings to open, causing the membrane to depolarize and generate action potentials (Reed-Geaghan and

Maricich, 2011). The firing capacity of mechanoreceptors are contingent on the skin deforming and transmitting force to the mechanoreceptor endings (Knibelstol and Vallbo, 1970). There are two aspects to tactile sensation; sensitivity, the ability to perceive sensation, and acuity; which is

the ability to distinguish detail between two spatially distributed points (McPoil and Cornwall,

2006).

A quantitative method of measuring tactile sensitivity is through the administration of a graded series of aesthesiometers known as Von Frey hairs or Semmes-Weinstein monofilaments

(hereafter, ‘monofilaments’). This technique was first used in the 1960’s to measure sensory loss in patients with brain injuries (Semmes et al., 1960). It involves applying nylon monofilaments of varying diameters (and hence compression strengths) perpendicular to the skin (Lambert et al.,

2009). Reaction to the monofilament by the subject is used to establish a tactile threshold (Field et al., 1999). Monofilament administration is non-invasive, and can be used for both nociceptive and non-nociceptive clinical evaluations in humans and animals (Field et al., 1999; Bradman et al., 2015). A calibrated series of monofilaments includes a set of multiple instruments, each of which each buckles at a specific force (measured in grams), when applied perpendicular to the cutaneous surface (Lambert et al., 2009). The aim is to determine the lowest buckling force that can be perceived by a participant. For humans, contact of the monofilament is an anticipatory stimulus, whereas for animals this contact is unexpected (Bradman et al., 2015). Humans, as reporting participants, communicate their interpretation of sensation, which then provides a perceptual threshold. For non-human subjects, reactions are purely instinctive, and their reactions

(such as foot/paw withdrawal) must be interpreted and recorded as a behavioral measurement by an external observer (Field et al., 1999; Bradman et al., 2015). After establishing a naïve baseline threshold, comparative changes in cutaneous sensitivity following a biological perturbation may be quantified (Field et al., 1997). For instance, in humans, tactile sensitivity has been demonstrated as a function of age and disease state - geriatric patients have higher tactile thresholds (i.e., lower sensitivity) than young adults (Thornbury and Mistretta, 1981), while

diabetic patients have higher tactile thresholds than healthy controls (McBride and Mistretta,

1982). Moreover, tactile sensitivity also varies across areas of the body, with the tactile threshold of the human foot sole being higher than the human palm (Bent and Lowrey, 2013).

While the somatosensory system is highly conserved across vertebrates, there are species-specific variations as a result of adaptations to different ecologies (Schneider et al.,

2016). For many species, including reptiles and various invertebrates, sensory receptors in the skin are known by a variety of terms including “sensilla” (singular: sensillum), “corpuscles”,

“tubercles”, and “papillae” (Hiller, 1978; Bauer and Russell, 1998; Crowe-Riddell et al., 2016).

Although they primarily function as mechanoreceptors, there is evidence to suggest that sensilla are likely multi-functional, participating as chemoreceptors, thermoreceptors, hygroreceptors, and even hydroreceptors (Hiller, 1976, 1978; Ananjeva et al., 1991). Among squamates (snakes and lizards) sensilla are found within depressions on the surface of scales (Hiller, 1978; Bauer and Russell, 1988). At least in snakes, innervation is achieved mostly through free nerve endings

(Siminoff and Kruger, 1968). Crocodiles and alligators have unencapsulated free nerve endings, lanceolate endings (associated with microscopic bristles), Merkel-like disks, and Pacinian corpuscles (Siminoff and Kruger, 1968). It is hypothesized that differences in mechanoreceptor types and distribution may be due to different modes of movement used, in roaming and hunting

(Johnson, 2001; Crowe-Riddell et al., 2016; Schneider et al., 2016).

Cutaneous tactile feedback is essential for modulating central pattern generator activity, which influences gait and balance (Meyer et al., 2004; McPoil and Cornwall, 2006). As a result, understanding tactile thresholds of load-bearing areas, such as the feet, contributes to a deeper understanding of biomechanics (Magnusson et al., 1990; Meyer et al., 2004). Mechanoreceptors in the feet establish a sense of body placement with respect to the ground by playing a role in the

reflexive modulation of descending motor commands to lower limbs (Zehr and Stain, 1999). In humans, reduced cutaneous feedback of the foot sole is directly correlated with postural instability (Wang and Lin, 2008), and has been shown to shift compensatory motions in the ankles and hips in humans (Meyer et al., 2004). These findings demonstrate that mechanoreceptors in the feet play an important role in governing postural responses, and underscore their role as an important contributor to dynamic balance (Nurse and Nigg, 1999;

Meyer et al., 2004; Strzalkowski et al., 2015).

1.3 The Cerebellum

The cerebellum is traditionally associated with fine motor control, the second-to-second use of sensory information or a motor command to produce an appropriate movement based on the body and the environment (Anderson, 1993; Baumann et al., 2015). In addition, the cerebellum plays a role in detecting motor errors and calculates internal models of the motor apparatus, essentially predicting and providing the mechanics (e.g., position and speed) of movement (Requarth and Sawtell, 2014; Veloz et al., 2015). This is essential for improvised connections of posture and gait, shaping dynamics in multi-limb coordination (Baloh et al.,

1998). Damage to the cerebellum commonly results in ataxia, or abnormal motor movements including impaired gait, slurred speech, and incoordination (Bastian et al., 1996; Therrien and

Bastian, 2015; Kemp et al., 2016). As such, the cerebellum has been implicated in many diseases associated with impaired motor control, such as multiple sclerosis, Friedrich’s ataxia, and

Creutzfeldt-Jakob disease (Angel, 1980; Bastian et al., 1996; Schmahmann et al, 2004; Louis et al., 2014).

In accordance with the accepted role of cerebellum in coordinating (but not initiating) movement, variation in cerebellar neuron structure may have adapted to the locomotor needs of

each species (Eccles, 1969). The mechanisms by which organisms maintain equilibrium, coordinate movements, and adapt movements to their respective environments are closely tied to cerebellar development and evolution (Muir, 2000; Hoogland et al., 2015). For example, precocial mammals – those that are able to move around independently, shortly after parturition

– have a well-developed cerebellum at birth. Altricial mammals - those that are still developing locmotor skills after parturition - do not have a well-developed cerebellum at birth (Sanchez-

Villagra et al., 2002). As such, precocial mammals may experience cerebellar plasticity due to synaptic changes in pre-existing circuits (Sanchez-Villagra et al., 2002). Motor aspects of postural adjustment are then generated by areas of the motor cortex (Takukusaki, 2017). Sensory signals flow into the CNS to the brainstem, cerebellum, thalamus, and cerebral cortex; the cerebellum and basal ganglia help integrate posture-gait control through reciprocal connections with the cortex and brainstem (Konnerth et al., 1990; Takakusaki, 2017). Other areas involved in integrating motor programs include the vestibular nuclei and the somatosensory cortex. Further, the cerebellum is also associated with long- and short-term motor plasticity (Anderson, 1993;

Jorntell and Hansel, 2006; Sdrulla and Linden, 2007; Hoxha et al., 2016).

Across vertebrates, the cerebellum varies in gross anatomy from a row of putative cerebellar cells in the lamprey, to the highly folded (foliated) structure seen in birds and mammals (Butz et al., 2014; Jacobs et al., 2014). Among reptiles, the cerebellum is often described as resembling a smooth dome or sheet-like structure (Larsell, 1926; Voogd and

Glickstein, 1998; Wylie et al., 2016). Located on the rostral roof of the fourth ventricle, the cerebellum was derived from the rhombic lip in the hindbrain (Butz et al., 2014). Despite the obvious variation in relative size and shape, the cerebellum of all vertebrates shares the same basic three-layered organization, as well as a comparable arrangement of afferent, efferent, and

intrinsic connections (Banga and Ten Donkelaar, 1982; Voogd and Glickstein, 1998). The layers of the cerebellum include: the cell-rich granular layer, the cell-sparse molecular layer, and, nested between the two, the Purkinje cell layer (Voogd and Glickstein, 1998; Butts et al., 2014).

Each layer is characterized by a distinctive population of neurons. The granular layer has abundant and densely organized glutamatergic neurons known as granular cells, whereas the molecular layer is invested with three different types of interneurons: stellate cells, basket cells, and Golgi cells (Klein et al., 2005; Dine et al., 2013). The most distinctive cells of the cerebellum are those of the Purkinje layer, the appropriately named Purkinje cells (Murchison et al., 2002). Purkinje cells are large, gamma-aminobutyric acid (GABA)-ergic, calbindin-positive neurons, located at the junction between the molecular and granule layers (Zupanc and Zupanc,

2006; Sillitoe et al., 2008). Among reptiles, the thickness of the Purkinje layer varies from one

(crocodiles and chameleons) to five (various lizards) cells thick (Shanklin, 1930; Nieuwenhuys,

1967).

Sensory information enters the cerebellar cortex via two excitatory afferents: mossy fibers, whose axons originate from many different types of neurons, and climbing fibers, which project from the medulla oblongata (Konnerth et al., 1990; Anderson and Korbo, 1993). Mossy fibers innervate granule cells, which in turn give rise to the parallel fibers that innervate Purkinje cells (via their dendrites) and interneurons (Rondi-Reig et al., 2014) (Figure 1.2). In contrast, climbing fibers directly innervate the Purkinje cell soma and proximal portions of the dendrites

(excitatory input), and also influence molecular layer interneurons (such as basket cells), which then inhibit Purkinje cells (Konnerth et al., 1990). Purkinje cells receive convergent inputs of the same multi-sensorial information from both mossy and climbing fiber inputs (Rondi-Reig et al.,

2014; Hoxha et al., 2016). In mammals, Purkinje cells are sites of maximal cerebellar

convergence, with more than 100,000 parallel fiber synapses, and maximal signal divergence, with approximately 100 trillion synaptic connections in humans and 60 billion synaptic connections in the rat (Hoxha et al., 2016). Although a similar pattern of connectivity is likely conserved across species, some variation does exist (e.g., reptiles lack basket cell inhibition)

(Llinas and Nicholson, 1971; Bangma and Ten Donkelaar, 1982). Climbing and parallel fiber synapses are hypothesized to play a role in motor learning, inducing long-term potentiation and/or long-term depression in the strength of Purkinje cell-parallel fiber inputs (Jorntell and

Hansel, 2006; Sdrulla and Linden, 2007).

In addition to coordinating movement, the cerebellum also contributes to somatosensation during active movement (Gao et al, 1996; Hartmann and Bower, 2001; Therrien and Bastian, 2015), receiving input from proprioceptive, tactile, vestibular, and visual receptors

(Reed-Geaghan and Maricich, 2012). As a result, damage to the cerebellum can impair both sensory and motor function (Gao et al., 1996; Schmahmann, 2004; Baumann et al., 2015). For example, cerebellar dysfunction can affect perception in movement control as well as somatosensory and visual domains (Therrien and Bastian, 2015). In an unusual case study, it was shown that an individual with a damaged right lateral cerebellar hemisphere had difficulty evaluating the weights of objects (abarognosis) when asked to lift them with his ataxic arm

(Angel, 1980). Inaccurate perception of load during movement may also relate to the disruption of the predicted sensory consequences, or efference copy of the motor command (Requarth and

Sawtell, 2014). This efference copy is a second, internal copy of movement-generating information that is relayed to the cerebellar cortex (Requarth and Sawtell, 2014). Damage to cerebellum distorts the efference copy, inducing a negative bias in limb dynamics (Therrien and

Bastian, 2015). Purkinje cells have been shown to trigger both single-joint motor learning, as

well as multi-joint behaviors (Black et al., 1990; Hoogland et al., 2015). For example, mutant mice with impaired Purkinje cell output and impaired Purkinje cell potentiation, have severely impaired locomotor performance and adaptation (Veloz et al., 2015). In addition, premature degeneration of Purkinje cells, as seen in patients with the neurodegenerative disease spinocerebellar ataxia type 1, causes impairments in motor coordination (Stucki et al, 2016).

At the level of neuromorphology, two main features characterize Purkinje cells: a relatively large-sized soma, and a complex and highly ramifying dendritic tree (Nieuwenhuys,

1967; Anderson and Korbo, 1993; Voogd and Glickstein, 1998). The distal dendritic branches are covered in spines (membranous protrusions of dendrites), whereas the more proximal branches are smooth (Nieuwenhuys, 1967; O’Brien and Unwin, 2006). Purkinje cell axons represent the sole output of the cerebellar cortex, and terminate on neurons of the deep cerebellar nuclei located in the cerebellar white matter (Konnerth et al., 1990). Neurons of the deep cerebellar nuclei then send projections to the brainstem or cerebral cortex via the thalamus (Butts et al., 2014). The brainstem and cerebral cortex then participate in equilibrium, posture, and vestibular and oculomotor control (Horak, 2010; Baumann et al., 2015).

1.4 Dendritic Morphology

Dendrites are branched extensions of a neuron that receive and transmit electrochemical stimulations to the cell soma (Tavosanis, 2011; Fujishima et al., 2012), and are well understood to be capable of plastic changes during development and throughout adult life (Butz et al., 2009;

Mendell et al., 2017; Louth et al., 2018). Although historically thought to be passive, electrophysiological recording techniques have since shown that neurons express numerous cell-specific forms of localized dendritic signaling (Eilers and Konnerth, 1997; Wong and Wong,

2000; Butz et al., 2009). Further, different neurons have distinctive corresponding dendritic

morphologies, and it has been established that there is a link between dendritic structure and function (Gao, 2007; Fiala et al., 2008). For example, dendrite surface area is correlated with number of contacts that a neuron receives, and therefore the size of the dendritic tree is a limiting factor for how many synaptic inputs the neuron can receive (Chiu et al., 2008; Tavosanis, 2011).

Modifications to dendrite structure are hypothesized to lead to changes in synaptic connections and input processing in the neuron, and represent a plausible mechanism for the nervous system to response to intrinsic and extrinsic changes (Butz et al., 2009). In addition, it is predicted that neuromorphological changes represent the basis for maintaining functional homeostasis in lieu of changed input (Tavosanis, 2011).

Most excitatory input in the CNS is facilitated by synapse formation on spines, located on dendrites of neurons (Konnerth et al., 1990; Radley et al., 2006). Changes in spine density or morphology are associated with changes in learning and memory (Adkins et al., 2002; Lee et al.,

2007). In addition, it is understood that dendritic spines and dendritic branches exist in dynamic balance with one another, with changes in dendritic structure making the neuron more amenable to the formation of new spines (Sweet et al., 2011). How does a change in input activity result in changes in dendrite morphology? Accumulating evidence points to a role for local calcium modulation in regulating dendritic growth cone behavior (Kater et al., 1988; Wong and Wong,

2000; Anwar et al., 2014). Neurotransmitters can either initiate or suppress dendritic growth by altering the postsynaptic membrane potential, which impacts voltage-dependent calcium channels (Zheng et al., 1994; Murchison et al., 2002; Butz et al., 2009). In turn, this results in modified intracellular calcium concentrations, which modulates growth cone behavior at the dendritic tips (Zheng et al., 1994; Butz et al., 2009). Therefore, changes in intracellular calcium concentration, triggered by neurotransmitter release, play an essential role in long-term plasticity

(Kater et al., 1988; Zheng et al., 1994; Anderson, 1993). These neuromorphological changes are also associated with downstream changes in neurophysiology. For example, there is a positive correlation between dendritic diameter and action potential amplitude (Hansson et al., 1994;

Vetter et al., 2001). There are also diameter dependent effects on intracellular calcium concentration, which has a role in the informational processing capabilities of neuronal dendrites

(Vetter et al., 2001), as well as the aforementioned role in maintaining plasticity. Overall, there is strong evidence to show that Purkinje cells play an important role synaptic plasticity. It is still unknown whether the changes occur in the Purkinje cells themselves, or if they induce synaptic plasticity elsewhere (or both). In either case, there are consequent behavioral implications in motor control.

One of the most common methods to evaluate changes in dendritic morphologies involves the use of the Golgi-Cox method. Golgi-Cox staining is a histological technique that allows for the in situ analysis of neuron morphology using light microscopy (Golgi, 1891; Louth et al., 2017). First developed by Golgi (Golgi, 1873) and then modified by Cox (Cox, 1891), this technique visualizes spine density, cell body shape, and dendritic tree morphology. Significantly, only a small percentage (~2%) of neurons are stained (Pasternak and Woolsey, 1975). To date, the reason for this selectivity remains undetermined. Ramon y Cajal used the Golgi-Cox technique to provide the first detailed tracings of cells across the nervous system, as well as the first efforts to map the neuromorphological changes that occur during development (Garcia-

Lopez et al., 2010). The Golgi-Cox technique allows for three-dimensional visualization of neurons, thus permitting details such as length, diameter, number of intersections, and number of terminals to be quantified (Louth et al., 2017). A common method for quantifying these features is the Sholl analysis (Sholl, 1953). Traditional Sholl analyses involve a mapping a series of

nested concentric circles, radiating out from the center of the cell soma, to index a variety of metrics including intersections, diameters, lengths, and areas (Sholl, 1953). Current Sholl analysis involves nested concentric spheres, allowing for analyses in three-dimensions. To date,

Golgi-Cox staining and three-dimensional imaging of Purkinje cells has mainly been performed in mammalian species (Garcia-Lopez et al., 2010; Jacobs et al., 2014, with only a few representative species shown for other classes of vertebrates (Nieuwenhuys, 1967; Uray and

Gona, 1978; Chan and Nicholson, 1986).

The size and complexity of a dendritic arbor can be altered by a number of factors, including electrical activity, drugs and chemicals, hormones, trophic factors, and external environment (Woodward et al., 1975; Floeter and Greenough, 1979; Wong and Wong, 2000;

Lein et al., 2007). Compared to other neuron types, the dendritic arbors of GABAergic neurons, such as Purkinje cells are known to be particularly dynamic (Llinas and Sugimori, 1980;

Tavosanis, 2011). For example, late postnatal mice exposed to regular exercise have markedly larger Purkinje cell dendritic trees and greater numbers of dendritic spines, than their less active counterparts (Pysh and Weiss, 1979). Further, many pathological conditions (e.g., Alzheimer’s disease) result in stress-induced remodeling of Purkinje cell dendrites (Mavroudis et al., 2010).

In humans, substantial alterations of the dendritic arbor are associated with cognitive impairment and motor deficits (Angel, 1980; Mavroudis et al., 2013; Schmahmann, 2004; Louis et al., 2014).

Essential tremor, a common movement disorder in humans characterized by postural and kinetic tremor, is associated with significant reductions in Purkinje cell dendritic complexity (Louis et al., 2014). In contrast, studies on rats have shown an increase in Purkinje cell synaptic plasticity associated with learning complex motor skills (Black et al., 1990; Lee et al., 2007).

1.5 Appendage Loss and the Leopard Gecko

Appendage loss is a dramatic, and often debilitating form of injury. In addition to the obvious trauma at the site of amputation, there are cascades of additional impairments that profoundly influence locomotor performance, behavior, and physiology (Dial and Fitzpatrick,

1981; Martin and Salvador, 1993; Alupay, 2013; Gilbert and Vickaryous, 2013; Gillis and

Higham, 2016). These include sensory deprivation from the amputated region, altered limb coordination, and various neurophysiological changes, such as a loss of GABAergic inhibition, as well as long-term potentiation (Wu and Kaas, 2002; Flor, 2008; Chen et al., 2012; Makin et al., 2013). In combination, these maladaptive alterations often trigger cortical reorganization, a form of neuroplasticity within the primary somatosensory cortex (Flor, 2008; Makin et al.,

2015). Phantom limb pain, the continued sensation of pain from a missing limb, is thought to occur as a byproduct of this maladaptive reorganization (Makin et al., 2013). A classic example of this cortical reorganization comes from the studies investigating proportional innervation density of the primary somatosensory cortex. This so-called sensory homunculus is essentially a proportionally deformed neurological map of somatosensation, dominated by regions of the body receiving the largest cortical representation: the face, hands, and feet (Pubols and Pubols, 1971;

Makin et al., 2015). Reorganization of the sensory homunculus occurs following amputation and other types of sensory deprivation (Pubols and Pubols, 1971; Wu and Kaas, 2002; Makin et al.,

2015). For example, in both humans and owl monkeys, digit amputations result in increasing cortical representation of the remaining fingers in the days (for humans) and months (for monkeys) following appendage loss (Merzenich et al., 1984). In another study, the somatosensory sensitivity of war veterans that were upper limb amputees was tested using monofilaments. This work revealed that amputees had significantly increased tactile sensitivity

in their residual limb compared to unamputated controls (Haber, 1955). In another study, following denervation of the forelimbs, the somatosensory cortex of rats is altered, along with remodeling of dendritic arbors of neurons in the forebrain (Hickmott and Steen, 2005).

Autotomy is a distinctive form of appendage loss, wherein the appendage is voluntarily self-amputated, often in response to the act or threat of predation (Jacyniak et al., 2017). For many species, the ability to autotomize is matched by an equally dramatic ability to spontaneously regenerate the lost appendage (Delorme et al., 2012). Among vertebrates, one of the best-known examples of autotomy and regeneration comes from lizards. Many lizards, including members of Gekkota (geckos), Scincidae (skinks), and Dactyloidae (anoles) are able to autotomize a portion of the tail and then regenerate a replacement appendage (Jacyniak et al.,

2017). As an anti-predation mechanism, autotomy is a rapid phenomenon and hence represents an extreme example of a rapid change in mass (Jagnandan and Higham, 2018). Recent studies investigating Eublepharis macularius, the leopard gecko, have revealed that autotomy can result in a near instantaneous loss of 1/3 of the animal’s total body length and 1/4 total body mass

(Jagnandan et al., 2014). As a result, there is a significant cranial shift in CoM, which impacts body orientation, postural stability, and locomotor performance (Jagnandan et al., 2014).

Following tail loss, there is also a reduction in vertical ground reaction force, and re-distribution of mass across the body (Jagnandan et al., 2014). Over approximately one month, a replacement tail is grown in place of that which was lost (McLean and Vickaryous, 2011). Although the new tail is superficially similar to the original, it differs in various structural details in terms of the composition of the skeleton (bone is replaced by cartilage) and arrangement of skeletal muscles

(McLean and Vickaryous, 2011; Gilbert et al., 2013). Furthermore, the CoM is never fully restored to back to its original (baseline) position (Jagnandan et al., 2014). Remarkably, stability

and gait are maintained in the leopard gecko following autotomy, and therefore running speed is not altered (Jagnandan et al., 2014). This appears to be achieved through joint kinematic alterations at the hindlimb, including changes in femur depression and knee angle, to take on a more sprawled posture (Jagnandan et al., 2014). Along with these postural alterations, there is a shift in hindlimb muscle recruitment, in muscles associated with propulsion (Jagnandan and

Higham, 2018).

Although the mechanism by which tail autotomy is initiated remains poorly understood, evidence points towards a role for cutaneous mechanoreceptors, scale sensilla (Russell, 1986;

Russell et al., 2014). Sensilla mediate extrinsic sensory input related to tactile and positional information (Russell, 1986). Although sensilla are found distributed across the entire body, lizards such as the leopard gecko demonstrate regional variation in receptor density (Russell et al., 2014). More specifically, there is a dorsoventral gradient, with the highest density of sensilla located on the dorsal surface of the tip of the original tail (Russell et al., 2014). While the exact role of sensilla in gecko tactile sensitivity and somatosensation remains to be explored, the observed pattern of distribution points towards two obvious (but not necessarily exclusive) roles:

(1) mechanical detection of tail grasping by predators (Russell et al., 2014); and (2) providing positional information associated with posture and counterbalance (Jagnandan and Higham,

2017).

Lizards compensate for autotomy using a variety of different strategies. Rock lizards

(Lacerta monticola) that have undergone autotomy spend more time heating their bodies than conspecifics with intact tails (Martin and Salvador, 1992). During tail regeneration in a different species of gecko, Hemidactylus flaviviridis, hemodynamic changes have been found to occur as an adaptive systemic response to tail regrowth (Hiradhar et al., 1978). On a cellular level,

immunological adaptations are seen in leukocyte activation featuring antimicrobial beta defensins, following autotomy (Alibardi et al., 2012). In terms of the nervous system, adaptations or physiological compensation following autotomy remain poorly understood.

CHAPTER 1 FIGURES

Figure 1.1. A schematic representation of central and peripheral nervous system integration with a focus on the coordination of movement.

Sensory input from the nervous system results in neuromuscular motor output. Information from the internal and external environment are relayed through sensory receptors of the peripheral nervous system (PNS) and are relayed to the central nervous system (CNS) via the spinal cord.

The integration of multiple sensory modalities (i.e., somatosensory, visual, and vestibular input) occurs in the brain stem. The motor cortex of the cerebral cortex integrates with the cerebellum in initiating and regulating motor programs, to initiate movement.

Figure 1.2. Synaptic connections of the cerebellar cortices.

Vertebrate cerebellar schematic with focus on the sole motor output of the cerebellar cortex, the

Purkinje cell. The conserved three-layered arrangement of the cerebellum is denoted on the right.

The gecko Purkinje cell, shown with its characteristic ramifying dendritic arbor, receives synaptic input from parallel and climbing fibers. Climbing fibers synapse on proximal portions of the dendritic arbor (one climbing fiber/Purkinje cell), while parallel fibers synapse the middle-distal part of the arbor (many parallel fibers/Purkinje cell). Note, basket cell and stellate cell input are excluded from this schematic. Excitatory (+) and inhibitory (-) input are denoted in brackets. The direction of action potentials is shown with arrows.

RATIONALE

Mass change is a biological phenomenon that is rarely addressed in the context of the nervous system. It is well understood that animals can respond to intrinsic and extrinsic changes by modulating sensory input and neuromorphology. Localized mass change in the form of appendage loss results in a shift of the center of body mass (CoM), which has important implications for posture, balance, limb coordination, and locomotor performance. Neural compensation, mediated through the peripheral nervous system (PNS) and central nervous system (CNS), is required to restore stability and biomechanical function. Two key elements of the neural compensatory response are the cutaneous somatosensory system (a component of the

PNS) and the cerebellum (of the CNS).

One of the most dramatic examples of localized mass change occurs in organisms capable of autotomy. Autotomy is an evolved voluntary mechanism of self-amputation commonly used by lizards to escape predation. Tail autotomy represents a significant and sudden change in body mass, leading to a cranial shift in the CoM. Over time, a new tail is regenerated, but the position of the CoM is never fully restored. Therefore, lizards represent a unique opportunity to investigate the neural compensatory response to: (a) a rapid change in mass, (b) a gradual restoration of mass, and (c) a permanent shift in CoM. Our work seeks to investigate the short and long-term adaptations of the nervous system to compensate for tail loss in a representative tail-autotomizing and tail-regenerating species, the leopard gecko (Eublepharis macularius). We hypothesized that, following tail loss and during regeneration, there will be somatosensory and neuromorphological changes to compensate for the change in the mass.

The objectives of my study were:

1) To characterize tactile sensitivity of the limbs and tail in geckos with their original tails,

and following tail loss and regeneration

2) To characterize Purkinje cell neuromorphology in geckos with their original tails, and

following tail loss and regeneration

CHAPTER 2: CHARACTERIZATION OF CUTANEOUS TACTILE SENSITIVITY

FOLLOWING TAIL LOSS AND REGENERATION IN THE LEOPARD GECKO

(EUBLEPHARIS MACULARIUS)

2.1 INTRODUCTION Sensory input involves the integration of multiple sensory modalities, including visual, vestibular, and somatosensory (Meyer et al., 2004; Carver et al., 2006; Horak, 2010).

Somatosensation, or tactile sense, refers to the detection and transduction of sensory information from the body surface (including skin and mucous membranes), muscles, limbs, joints, and internal organs to the central nervous system (CNS), and their interaction with the environment

(Shumway-Cook and Horal, 1986; Quai et al., 2004). This involves participation of several different types of receptors including pain (nociception), temperature (thermosensation), and touch (mechanoreception) (Reed-Geaghan and Maricich, 2011; Roudaut et al., 2012; Bent and

Lowrey, 2013). Cutaneous mechanoreceptors (those within the skin), detect mechanical pressure associated with deformation of the skin, enabling the perception of touch (Johnson, 2001; Reed-

Geaghan and Maricich, 2011). In addition to detecting relatively benign pressures, cutaneous mechanoreceptors provide sensory feedback used in regulating stability and movement in accordance with changes in the external environment (Shumway-Cook and Horak, 1986;

Stzalkowski et al., 2015).

Tactile mechanoreceptors of the skin (hereafter ‘mechanoreceptors’) have low activation thresholds (i.e., high sensitivity), and thus even weak stimulations of the skin can initiate action potentials (Purves et al., 2001, Macefield, 2005). Each mechanoreceptor consists of one or more peripheral (sensory) nerve ending, with cell bodies or somas located within the dorsal root

ganglia (Johnson, 2001). Classically, mechanoreceptors are categorized as either slowly (firing for the duration of stimulus) or rapidly adapting (bursts of firing at the beginning and end of stimulation), and unencapsulated (free nerve endings; unencapsulated dendrites of sensory neurons) or encapsulated (dendrites contained within a capsule) (Knibelstol and Vallbo, 1970,

Macefield, 2005). Mechanoreceptors generate impulses that are transmitted along large myelinated axons, which rapidly transmit sensory information to the spinal cord and the CNS

(Reed-Geaghan and Maricich, 2011). The heterogeneous organization and density of different mechanoreceptors provides variation in tactile thresholds and receptive fields (the region of sensory space that affects firing of a neuron) (McPoil and Cornwall, 2006). Mechanoreceptors with small receptive fields (e.g., Merkel and Meissner receptors) allow for more precise detection than mechanoreceptors with large receptive fields (e.g., Ruffini and Pacinian receptors)

(Macefield, 2005). Interestingly, the receptive fields of tactile sensory neurons, to perception, are adaptable and can become larger or smaller in response to injury or experience (Weinberger,

1995).

Whereas the general organization of the somatosensory system is conserved across vertebrates (Roudaut et al., 2012; Schneider et al., 2016), the homology of mechanoreceptors across taxa is poorly understood. For example, both rapid and slow acting mechanoreceptors have been identified in reptiles (Siminoff and Kruger, 1968; Hiller, 1978; Bauer and Russell,

1988), but how these compare with their mammalian counterparts is unknown. Reportedly, crocodylians have mechanoreceptors resembling Merkel cells, and Pacinian corpuscles, and those of snakes are mostly unencapsulated (Siminoff and Kruger, 1968), but for most species the structure and diversity of these tactile organs are yet to be investigated. Highlighting this uncertainty, mechanoreceptors in reptiles (and many invertebrates) have been referred to as

“corpuscles”, “tubercles”, “papillae” and “integumentary sensory organs,” although current usage favors “scale sensilla” or simply “sensilla” (Hiller, 1976, 1978; Ananjeva et al., 1991;

Russell et al., 2014; Crowe-Riddell et al., 2016). Whereas reptilian sensilla are predominantly mechanoreceptive in function (Hiller, 1978), there is evidence to suggest that additional sensory roles may be present in some species. For example, in sea snakes (Elapidae, Hydrophiinae) sensilla are thought to function as both tactile and hydrodynamic receptors, which sense water displacement (Crowe-Riddell et al., 2016), while in some lizards they play roles in hydro- and thermosensitivity (Hiller, 1976; Ananjeva et al., 1991). Like mammals, the cell bodies of mechanoreceptors reside within the dorsal root ganglia of the spinal cord (Johnson, 2001).

Across lizard species, sensillum morphology is widely conserved (often described as rounded papilla nested within a depression on the scale surface) (Hiller, 1978; Russell, 1986;

Bauer and Russell, 1988). However, the spatial organization and distribution of sensilla is taxonomically variable. For example, iguanids have ~8 times more sensilla than agamids

(Ananjeva et al., 1991). These distinctions most likely relate to differences in regional integumentary function as well as ecological factors, such as prey capture and foraging habits, and environment such as nocturnality (Schneider et al., 2016). Related to this, many lizards have a high concentration of sensilla on the head, likely associated with roles in feeding (Jackson,

1977), and along the dorsal surface of the tail, likely associated with tail autotomy (Russell et al.,

2014).

Tail autotomy is voluntary form of self-amputation used by many species of lizard to escape predation (Cromie and Chapple, 2013; Gillis and Higham, 2016; Jacyniak et al., 2017).

The act of grasping the tail initiates a reflexive behavior that results in the loss of self-detachment of a portion of the appendage. Although the exact mechanism initiating tail

autotomy remains unclear, a role for mechanoreceptors, detecting deformation of the skin during a predation event, is strongly indicated (Maclean 1980; Russell et al., 2014). In support of this prediction, the highest density of sensilla in geckos is associated with the dorsal side of the tail

(Russell et al., 2014). Tail autotomy is facilitated by a number of structural adaptations that minimize damage to adjacent tissues of the retained tail stump. These include fracture planes

(zones of weakness within each vertebra that subdivide the centrum during tail detachment), as well as arterial sphincters (Mclean and Vickaryous, 2011; Delorme et al., 2012; Jacyniak et al.,

2017). Arterial sphincters are circular bundles of smooth muscle wrapped around the main artery of the tail, and are located in advance of each fracture plane. When the tail detaches, the sphincters contract and blood loss is minimized (Delorme et al., 2012). Interestingly, the nervous system appears to lack any evidence of structural modification associated with autotomy.

Therefore, during tail autotomy, the tail spinal cord and nerves are effectively (and even uncontrollably) torn (Duffy et al., 1992; McLean and Vickaryous, 2011).

Tail regeneration is spontaneously initiated once the tail is detached and can be broadly characterized into two main events: wound healing and regenerative outgrowth (McLean and

Vickaryous, 2011). Wound healing begins with clotting at the wound surface, retraction of the ruptured spinal cord, blastema formation, and the recruitment of osteoclasts and chondrocytes

(McLean and Vickaryous, 2011). Once the stump has re-epithelialized, regenerative outgrowth begins with the formation of a cartilaginous skeleton around the ependymal tube, tissue differentiation, axonal regrowth, scale formation, and pigmentation (McLean and Vickaryous,

2011). Although the resulting tail grossly resembles the original, it differs in various anatomical details (Gilbert et al., 2013). For example, the skeleton of the regenerated tail is an unsegmented cone of cartilage, whereas the original skeleton is a series of bony vertebrae (McLean and

Vickaryous, 2011; Delorme et al., 2012). In addition, whereas the spinal cord of the original tail closely resembles that of the body, the regenerated organ lacks grey matter (Gilbert and

Vickaryous, 2018).

Although tail autotomy provides an immediate means for evading predation, it also has important consequence on various fundamental traits. For example, tail autotomy is associated with an instantaneous change in overall body mass and length, and therefore a significant cranial shift in center of mass (CoM) (Jagnandan et al., 2014). This shift thus imparts a re-distribution of body mass among the limbs. Recent work has determined that while there are transient changes in hindlimb ground reaction forces and hindlimb joint angles following tail autotomy (largely resolved following tail regeneration), there are no significant changes in forelimb kinematics

(Jagnandan et al., 2014). In humans, there is an established link between sensory input from plantar surfaces in the feet, and gait kinematics (Magnusson et al., 1990; Nurse and Nigg, 1999).

This link is emphasized in circumstances of human limb loss (Templeton et al., 2018). Whether the transient changes observed in geckos are also associated with changes in tactile sensitivity (as has been reported for in humans) remains unknown.

Here, we conducted a spatiotemporal investigation (before and after tail loss, and once regeneration is complete) of tactile sensitivity at multiple locations across the ventral body surface of a tail-autotomizing lizard, the leopard gecko (Eublepharis macularius). More specifically, we performed monofilament testing to quantify sensitivity changes in response to immediate mass loss (tail loss) and gradual mass gain (tail regeneration). The leopard gecko is a well-established model of tail autotomy and regeneration (Mclean and Vickaryous, 2011;

Delorme et al., 2012; Peacock et al., 2015; Gillis and Higham, 2016; Jacyniak et al., 2017;

Gilbert and Vickaryous, 2018). Tail regeneration in this species is a relatively rapid (~30 days),

epimorphic (blastema-mediated) process that generates a structurally similar but non-identical replacement appendage (McLean and Vickaryous, 2011; Delorme et al., 2012; Gilbert et al.,

2013). Previous work has determined that tail loss in leopard geckos results in a cranial shift in

CoM that is never fully restored - even once the tail is completely regenerated (Jagnandan et al.,

2014). We predicted that tactile sensitivity would be permanently altered following tail loss.

Unexpectedly, we found that while tactile sensitivity does change following tail loss, this effect was only transient.

2.2 MATERIALS AND METHODS

2.2.1 Experimental Animals and Animal Care

Captive bred leopard geckos (E. macularius; hereafter ‘geckos’) were obtained from a commercial supplier (Global Exotic Pets, Kitchener, Ontario, Canada). All geckos were subadults, less than one year in age and had an average body mass of 20.2g (17.1-22.4g). For the main experiments (tactile sensitivity [Chapter 2] and neuromorphology [Chapter 3]), we used a cohort of 15 geckos; for the serial histology characterization, we used a separate group of three subadult geckos. Growth of all geckos was tracked weekly throughout the experimental period by measuring snout-vent length, tail length, and body mass. Animal Utilization Protocols (AUPs) were approved by the University of Guelph Animal Care Committee (Protocols #1954, 3772) and followed the guidelines of the Canadian Council on Animal Care. Animal husbandry follows the work of McLean and Vickaryous (2011). Gecko colonies were housed in a secure vivarium

(Hagen Aqualab facility) at the University of Guelph, inside a temperature controlled environmental chamber (average ambient temperature, 27.5 ̊C). Geckos were housed individually in 5 galleon polycarbonate tanks, with a heat cable (Hagen Inc., Baie d’Urfé,

Québec, Canada) set at 32ºC and placed underneath one side of the enclosure to establish a

thermal gradient. Each enclosure included two hide boxes and a water dish. Geckos had constant access to fresh drinking water, and were fed 3 larval mealworms (Tenebrio spp.) daily, dusted with powdered calcium and vitamin D3 (cholecalciferol; Zoo Med Laboratories Inc., San Luis

Obispo, California, USA). The gecko chamber was kept on a year-round 12h light:12h dark photoperiod, and all experiments were conducted in a 6-week span from November-December.

2.2.2. Experimental Design

Both the tactile sensitivity and neuromorphological experiments (Chapters 2 and 3) used the same cohort of 15 geckos. These geckos were randomly assigned into one of three groups

(n=5 per group): control (tail intact); tail loss (8 days post-autotomy); and tail regenerated (32 days post autotomy). The experimental timeline was either eight days (for the tail loss group) or

32 days (for the control and tail regenerated groups) (Figure 2.1). Autotomy for the tail loss and tail regenerated groups occurred on day 1 of the experiment; control geckos retained their original tails for the duration of the experiment. Each gecko was used for both tactile sensitivity testing at multiple time points and neuromorphological evaluation (following euthanasia).

2.2.3 Tail Autotomy

Self-detachment of the tail, tail autotomy, is a naturally evolved anti-predation strategy common to many species of lizard. Tail autotomy was performed by manually restraining the gecko, and then pinching the base of the tail between the forefinger and thumb. The tail was then self-detached, and the gecko was released back into its enclosure. All geckos survived the autotomy procedure and all autotomies occurred at roughly the same point along the tail

(adjacent to the tail base), corresponding with a maximal tail loss. Control geckos were manually restrained to simulate autotomy conditions, but did not have their tails removed.

2.2.4 Monofilament Testing

Cutaneous sensory responses were tested by applying Semmes-Weinstein monofilaments

(pressure aesthesiometers; hereafter ‘monofilaments’) to six sites of interest following the method of Field et al. (1997). Sites of interest included ventral surfaces of: the left and right hindlimbs (plantar surface of the pes), left and right forelimbs (palmer surface of the manus), and the (proximal) base and (distal) tip of the tail (Figure 2.2).

Geckos undergoing monofilament testing were placed in a clear plexiglass chamber

(31.5cm wide, 29.5cm high, 31.5cm deep) mounted on a raised, perforated (~10mm diameter) platform. A mirror suspended at an oblique angle below the perforated platform was used to direct the monofilaments through the perforations (Figure 2.3A-C). A video camera (Panasonic

SDR-H85) was mounted outside the chamber to record the trials, and the front of the chamber was covered with a dark partition, to blind the gecko to the presence of the experimenter. Geckos were acclimated to the testing chamber for one hour prior to the start of each monofilament testing session. Monofilaments were applied perpendicular to the skin in an area roughly corresponding to the middle of each site of interest. Pilot studies established that geckos responded to the most lightly-weighted monofilaments (of the set). Thus, to evaluate each gecko’s withdrawal response, we used an ascending series of calibrated monofilaments

(Table 2.1) measuring force in grams, starting with 0.091g (monofilament #3). Each monofilament was applied perpendicular to the skin until buckling. Once buckled, the monofilament was maintained in position for approximately one second, before being withdrawn. A positive withdrawal response from the gecko was scored when application of the monofilament resulted in lifting of the foot (when testing the forelimbs and hindlimbs) or body/tail (when testing the tail) within the 1 second of application. When no withdrawal

response was observed, the application was deemed negative. Following a negative response, the next heaviest weighted monofilament was selected, and the site of interest was re-tested (Figure

2.4). This process was repeated until a positive withdrawal response was observed. The smallest weighted monofilament (in grams) that evoked a positive response was recorded as the tactile threshold (hereafter, ‘threshold’). Re-application of the series was done for each site (a test and re-test), and the average of the two thresholds was taken to be the lowest sensory response. The order of site testing was chosen pseudorandomly, and somewhat opportunistically, as sites of interest could only be tested when they were placed over the perforations in the raised platform.

Monofilament testing was only administered when the gecko was motionless, with all four feet contacting the platform.

Prior to autotomy (day 0 of the experiment), baseline thresholds were established for all

15 geckos. Geckos were then tested on day 2 (one day post-autotomy for the experimental groups), and then every seven days thereafter, for 30 days. All geckos in the same group (tail intact, tail loss, or tail regenerated) were tested on the same day. Daily feeding for geckos being tested took place only after all monofilament testing for the day was finished. One gecko from the control group (gecko #5) did not react to monofilament testing and was removed from the experiment.

2.2.5 Euthanasia, Tissue Collection, and Preparation

Geckos were euthanized with intra-muscular injection of 150μL Alfaxan (Alfaxalone) into the epaxial muscles of the neck. Geckos collected for histochemistry and immunofluorescent staining were then fixed by transcardial perfusion of phosphate buffered saline (PBS), followed by 10% neutral buffered formalin (NBF; Fisher Scientific, Waltham, Massachusetts, USA).

Whole brains were then dissected out of the skull, and further fixed in 10% NBF for 22 hours.

Tissues were then transferred to 70% ethanol until processing. Geckos collected for Golgi-Cox staining were immediately dissected, the brains removed (without fixation), and placed into

Golgi-Cox solution.

2.2.6 Statistical Analysis Statistical analyses for the monofilament data were performed using Statistical Analysis

Software (SAS Institute Inc, Cary, NC), with descriptive statistics reported as the normalized means ± SEM. Outliers were calculated as values greater than ± 2 Standard Deviations (SD) from the mean for the entire data set. Data was then pooled by site and outliers removed outside of the mean ± 2 SD for each site. When an outlier was removed, the second (re-test) value for that test replaced it, except if the test and re-test values were adjacent, up to 0.369g

(monofilament #7). For each group, data were normalized to baseline values (i.e., day 0) and expressed as a ratio (Table 2.2). Data was log-transformed to meet assumptions of parametric testing. Normality was tested using the Shapiro-Wilk test (p<0.05), and Brown & Forsythe’s test determined homogeneity of variance. A three-way repeated measures analysis of variance

(ANOVA) (Group (3) x Site (6) x Time (6)) was performed to ensure there was not a three-way interaction between measures. After ensuring there was no significant difference between groups prior to autotomy, an additional one-way repeated measures ANOVA assessed the raw monofilament data (in grams) at baseline, pooling the three experimental groups to identify any regional differences (Figure 2.5). Lastly, the data was separated into the four regional site subsets

(hindlimbs, forelimbs, tail base, and tail tip) to examine the effect of tail loss and regeneration over time using a two-way repeated measures ANOVA (group (3) x time (6)) (Figures 2.6-2.9).

All post hoc analyses examined pair wise comparisons using Fisher’s least significant difference

(LSD) test. Significance level for all analysis was determined as p< 0.05. Based on the research aims, apriori hypotheses were developed to examine specific post hoc comparisons for each

ANOVA, outlined within the results section. Only significant post hoc comparisons were reported.

2.3 RESULTS

2.3.1 Experimental Geckos

At the start of the experiment, all geckos were subadults, with no signs of secondary sexual characteristics, and had an average mass of 20.2g (17.1-22.4g), an average snout-vent body length of 98mm (90-100mm), and an average tail length of 80.6 (70-90mm) (Appendix I,

Table A1). All geckos continued to grow throughout the experimental timeframe, including those undergoing tail autotomy. Following autotomy, geckos in the tail loss and tail regeneration groups lost an average of 19.28% of their body mass, and an average of 39.64% of their body length (Appendix I, Tables A3, A4). The three experimental gecko groups all followed the same timeline, but whereas the tail loss group was euthanized on day 8 of the experiment, the tail regeneration and control groups were euthanized on day 31 (Figure 2.1). As such, there is no data for the tail loss group at the last three timepoints of the experiment.

By the end of the experiment, the average masses for each of the groups were: 24.8g

(control), 19.6g (tail loss), and 21.8g (tail regeneration). Snout-vent body lengths for all groups closely paralleled one another throughout the experiment, with an average length of 105mm (95-

110mm) for the control and tail regeneration groups on day 31 (Appendix I, Table A2). At day 8 of the experiment, geckos in the tail loss group remained in the wound healing phase of regeneration – i.e., they either retained the clot covering the site of autotomy (representing stage

II of regeneration; Mclean and Vickaryous, 2011) or the clot was lost but no new outgrowth had occurred (i.e., stage III). At day 31 of the experiment, all geckos in the tail regeneration group

had fully regrown new tails (i.e., stage VII), and hence were well within the regenerative outgrowth phase.

Six sites of interest were investigated using monofilaments (one on the palmer/plantar surface of each limb [manus and pes] and two locations on the ventral side of the tail). Tail sites included the tail base, which was tested in all geckos, and the tail tip. The original tail tip was investigated for all groups at day 0 (baseline), and the control group throughout the experiment.

The regenerate tail tip was only investigated in the tail regeneration group, but only once regenerative outgrowth had occurred (starting at day 14). Beyond day 0, tail tip data was not collected from the tail loss group.

2.3.2 Monofilament Testing

Monofilaments used in the experiment were calibrated prior to testing (range: 0.091g to

67.1g) (Table 2.1). No gecko was tested with a monofilament rated higher than 6.2g during the experiment (that was not considered an outlier). On average, thresholds for experimental groups ranged between 0.09-0.2g for all sites (Appendix III, A5-A7). Test and re-test values were either the same for each site, or within +/- two logarithmic intervals (i.e., two monofilaments).

Comparing all groups over time (3 groups x 4 sites x 6 timepoints), we found there was a main effect of group (three-way repeated measures ANOVA, F2,7=5.36, p=0.04), site (three-way repeated measures ANOVA, F3,11=7.1, p=0.006). and time (three-way repeated measures

ANOVA, F5,32=9.26, p<0.001) (Appendix V, Table A9). Additionally, an interaction effect existed between group and site (three-way repeated measures ANOVA, F5,13=2.98, p=0.05).

Based on a priori hypotheses, post hoc comparisons (Fisher’s LSD) for the group and time interactions were examined, to identify how tail loss and regeneration influenced regional differences in cutaneous sensitivity. Post hoc analyses revealed that there were no significant

differences between left and right forelimbs and hindlimbs (p>0.05). As a result, data for left and right sides were pooled for forelimbs and hindlimbs. Post hoc analyses also showed that at Day

0, there were no significant differences between experimental groups (p>0.05). Therefore, groups were pooled when looking at regional differences at prior to autotomy.

2.3.3 Regional Differences Prior to Autotomy

To determine if there were differences in sensitivity between the forelimbs, hindlimbs, tail base, and tail tip prior to autotomy, we investigated each group at day 0 (=baseline), when all geckos still had original tails. We found there was a significant main effect of site (one-way repeated measures ANOVA, F3,11=6.43, p=0.0089) (Figure 2.5). The most sensitive site of interest (i.e., the site of interest with the lowest threshold) was the base of the tail (0.11g±0.009), followed by the forelimbs (0.13g±0.01), original tail tip (0.14g±0.001), and the hindlimbs

(0.19g±0.007). The threshold of the hindlimbs was significantly higher than the forelimbs

(Fisher’s LSD post hoc, p=0.005) and tail base (Fisher’s LSD post hoc, p=0.003). There was no baseline measurement of the regenerate tail, as this was not measurable until day 14 of the experiment. Based on these regional differences in sensitivity, we used targeted ANOVAs to evaluate each site separately for the remainder of the analysis.

2.3.4 Sensitization of Control Group

Over the time course of the experiment, all groups experienced repeated exposure to monofilament testing. In the control group, the only group that did not undergo autotomy, there was a significant decrease in threshold (i.e., sensitivity increased) at all sites (Figures 2.6-2.9), compared to baseline. This began at day 2 for the forelimbs (Fisher’s LSD post hoc, p=0.003) and tail base (Fisher’s LSD post hoc, p=0.009), day 7 for the hindlimbs (post hoc, p=0.05), and

day 14 for the tail tip (Fisher’s LSD post hoc, p=0.02). At each site, sensitization was maintained for the duration of the experiment, with the exception of the tail tip at day 21 (Fisher’s LSD post hoc, p=0.16). There were no significant differences between consecutive timepoints, apart from baseline and day 2 (17-30% reduction). The sites most affected by sensitization over time were the hindlimbs and forelimbs (Figure 2.6, 2.7), which dropped to 67% (day 14) or 68% (day 31) of baseline threshold, respectively (Table 2.2).

2.3.5 Hindlimb Sensitivity Following Tail Loss and Regeneration

Focusing on the hindlimb data across all groups over time, we found there was a significant main effect of time (two-way repeated measures ANOVA, F5,123=5.29, p<0.001), but no effect of group (two-way repeated measures ANOVA, F2,123=1.78, p=0.174), and no interaction between group and time (two-way repeated measures ANOVA, F7,123=0.52, p=0.817). Overall, hindlimb threshold decreased in all three groups (Figure 2.6). At day 2 (one day after autotomy), the hindlimbs in the tail loss group (Fisher’s LSD post hoc, p=0.04) and tail regeneration (Fisher’s LSD post hoc, p<0.001) groups were significantly different from baseline threshold, while the control (Fisher’s LSD post hoc, p=0.2) was not. Additionally, on day 2, threshold of the tail regeneration group was significantly lower than the control group (Fisher’s

LSD post hoc, p=0.023). By day 7, and throughout the remainder of the experiment, all three groups were significantly lower than baseline threshold. Hindlimb sensitivity in the control group and the tail regeneration group were not significantly different from one another on days 7,

14, 21, and 31.

2.3.6 Forelimb Sensitivity Following Tail Loss and Regeneration

Focusing on the forelimb data across all groups over time, we found there was a significant main effect of group (repeated measures two-way ANOVA, F2,123 =7.27, p=0.001), and a significant main effect of time (repeated measures two-way ANOVA, F5,123=3.29, p=0.0008), with no interaction effect (repeated measures two-way ANOVA, F7,123=1.13, p=0.349) (Figure 2.7). At day 2, forelimb threshold of the control group (Fisher’s LSD post hoc, p=0.003), but not the tail loss (Fisher’s LSD post hoc, p=0.88) or tail regeneration (Fisher’s LSD post hoc, p=0.44) groups, was significantly different from thresholds at day 0 (baseline). This is essentially the opposite trend as was observed for the hindlimbs at the same time point. When compared as a percentage of change from the day 0 (baseline) threshold, the control group at day

2 decreased 30.4%, while the tail loss and tail regeneration groups decreased only 2.5% and

8.8%, respectively. Further, both the tail loss (Fisher’s LSD post hoc, p=0.003) and tail regeneration (Fisher’s LSD post hoc, p=0.03) groups had significantly higher thresholds than controls at day 2. At day 7, the tail loss group had significantly higher threshold than controls

(Fisher’s LSD post hoc: p=0.006), and also (unexpectedly) the tail regeneration group (Fisher’s

LSD post hoc: p=0.02). Forelimb threshold of the tail regeneration group did not decrease significantly from day 0 (baseline) until day 31 (Fisher’s LSD post hoc, p=0.02).

2.3.7 Tail Base Sensitivity Following Tail Loss and Regeneration

Focusing on the tail base data across groups over time, there was a significant main effect of group (two-way repeated measures ANOVA, F2,54 =6.46, p=0.003), and a significant main effect of time (two-way repeated measures ANOVA, F5,54=4.58, p=0.002), with no interaction between group and time (two-way repeated measures ANOVA, F7,54=1.11, p=0.378) (Figure

2.8). At day 2, threshold of the control group (Fisher’s LSD post hoc, p=0.009) and the tail

regeneration group (Fisher’s LSD post hoc, p=0.008) were significantly lower than baseline, while the tail loss group was not (Fisher’s LSD post hoc, p=0.965). Comparing experimental groups at day 2, the tail loss group had significantly higher threshold than the control group

(Fisher’s LSD post hoc, p=0.005) and (unexpectedly) the tail regeneration group (Fisher’s LSD post hoc, p=0.009). The tail loss group did not significantly differ from its baseline at day 2 or day 7. At day 7, threshold of the tail loss group was significantly higher than the control group

(Fisher’s LSD post hoc, p=0.05). The tail regeneration group and the control group were significantly lower than their respective baselines for all timepoints, except for the tail regeneration group at day 21 (Fisher’s LSD post hoc: p=0.15). At day 21, threshold of the tail regeneration group was significantly higher than the control group (Fisher’s LSD post hoc, p=0.05). Within the tail regeneration group, all geckos responded to the lowest-weighted monofilament (=0.091g). Therefore, the threshold values for both the test and re-test did not vary within the group (Figure 2.8). This most likely represents a ceiling effect of the monofilament testing.

2.3.8 Tail Tip Sensitivity Following Tail Loss and Regeneration

At day 0, tail tip data was collected from the original tail for all groups. For the tail regeneration group, tail sensitivity at subsequent timepoints was measured on the replacement

(i.e., regenerated) tail. Testing of the regenerated tail tip began on day 14, once the replacement tail was long enough to be tested. The tail loss group was euthanized at day 8, and therefore was not included in this comparison. Focusing on tail tip data in the control and tail regeneration groups over time, there was a significant main effect of group (two-way repeated measures

ANOVA, F1,25 =4.20, p=0.05), and a significant main effect of time (two-way repeated measures

ANOVA, F3,25=6.44, p=0.002), but no interaction between group and time (two-way repeated

measures ANOVA, F3,25=0.62, p=0.606). Overall, thresholds for both groups decreased from baseline during the experimental timeframe (Figure 2.9). At no point were thresholds of the control and tail regeneration groups significantly different from one another, although the tail regeneration group was consistently more sensitive (i.e., threshold of the tail regeneration group was 67% of baseline, whereas the threshold of the control group was 78% of baseline). Within the tail regeneration group, all geckos responded to the lowest-weighted monofilament

(=0.091g). Therefore, threshold values, for both the test and re-test at day 14 and day 31 did not vary within the group (Figure 2.9). This most likely represents a ceiling effect of the monofilament testing.

2.4 DISCUSSION

Autotomy is an evolved form of self-mutilation employed to escape predation (Jacyniak et al., 2017). Previous research has demonstrated that lizards undergoing tail autotomy experience a rapid and dramatic change in total body length and body mass, and a cranial shift in

CoM (Jagnandan et al., 2014). Using monofilaments, we determined that there are regional differences in tactile sensitivity prior to and following autotomy. With repeated testing, the forelimbs, hindlimbs, tail base, and tail tip of control (tail-intact, un-autotomized) geckos, and the hindlimbs, tail base, and tail tip of tail-autotomized geckos became sensitized. Curiously, the forelimbs of tail autotomized geckos were resistant to sensitization, at least until the tail was fully regenerated. These findings indicate that autotomy, and the resulting associated cranial shift in CoM, affect peripheral tactile sensitivity in geckos.

To the best of our knowledge, this study is the first to use monofilaments on a species of lizard, and the first to perform a multi-site comparison during regeneration. In humans, monofilaments are most commonly used as clinical tools to evaluate tactile sensitivity under

pathological or amputation conditions (Haber, 1955; Thornbury and Mistretta, 1981; Thornbury and Mistretta, 1982; Templeton et al., 2018). Monofilaments have also been used as tools to evaluate instinctive (i.e., non-anticipated responses) in various rodents (Field et al., 1999; Ren,

1999; Tanaka et al., 2013; Bradman et al., 2015), as well as horses (Redua et al., 2002), octopuses (Alupay, 2013), and turtles (Gornik et al., 2015). While the hands and feet are the most common targets for testing, monofilaments have also been used to test sensitivity of the eyeball (Gornik et al., 2015), autotomized arm (Alupay, 2013), and thigh (Redua et al., 2002).

2.4.1 Regional differences in tactile sensitivity exist prior to autotomy

This study determined that there are regional (i.e., site-specific) differences in tactile sensitivity across the ventral surface of geckos prior to autotomy. More specifically, we found that the ventral surface of the tail base had the lowest threshold (0.11g±0.009), followed by the forelimbs (0.13g±0.01), original tail tip (0.14g±0.001), and hindlimbs (0.19g±0.007). Variation in threshold across sites likely relates to the pattern of sensilla distribution across the body.

Previous research has demonstrated that gecko sensilla are more densely distributed on the ventrolateral surface of the original tail (mean: 17.4/mm2) as compared to the ventral surface of the hindlimb (mean: 15.5/mm2) (Russell et al., 2014). This pattern of distribution is consistent with the prediction of skin deformation playing a role as an initiating factor in autotomy

(Maclean, 1980; Russell et al., 2014). Interestingly, while the same study determined that there is a greater density of mechanoreceptors (sensilla) at the tail tip (mean: 25/mm2), compared to the tail base (mean: 17.4/mm2), we found that the tail base (and not the tip) was more sensitive. Our findings suggest that receptor density is not necessarily an accurate predictor of overall tactile sensitivity. Sensory acuity and central processing are most likely playing a role in the heightened sensitivity of the tail tip. By way of explanation, we note that from an evolutionary perspective,

the tail base is more functionally relevant than the tail tip. The tail base supports the load of the length of the tail, as well as facilitating pelvic rotation and stabilizing lateral undulations

(Jagnandan and Higham, 2017). The tail base is also valuable in terms of being closer to the body, where there is higher risk of life-threatening damage. A similar trend towards increasing sensitivity in proximal locations is seen in the octopus, another species capable of autotomy of its arms. As in geckos, the tactile sensitivity of the octopus is significantly higher at the arm base compared to the arm tip (Alupay, 2013).

Differences in threshold may also relate to differences in limb and joint kinematics, appendage loading, and predator detection (Foster and Higham, 2012; Jagnandan et al., 2014;

Russell et al., 2014). In the context of locomotion, the asymmetrical sensitivity of the forelimbs and the hindlimbs is matched by differences in functional roles: hindlimbs are primarily involved in propulsion, whereas forelimbs participate as brakes, energy absorption, and stability (Lee,

2010; Foster and Higham, 2012; Autumn et al., 2016). When faced with environmental challenges in arboreal and terrestrial environments, lizard species show differential responses between the forelimbs and hindlimbs, as a result of differences in function, anatomy, kinematics, and behavior (Losos, 1990; Foster and Higham, 2012; Jagnandan et al., 2014). Hence, the greater sensitivity of the forelimbs, as compared to the hindlimbs, may be due to the transferred load to the forelimbs following autotomy in the leopard gecko during regular stance and locomotion. In humans and quadrupeds, body weight is a key determinant in ground force and a crucial parameter for plantar pressure distribution, therefore affecting distribution in vertical forces in load-bearing areas such as the foot pad (Magnusson et al., 1990; Nurse and Nigg, 1999).

Previous studies have shown that some quadrupeds (such as canines and horses) have vertical

forces in the forelimb are consistently higher than those in the hindlimbs (Besancon et al., 2004;

Munoz-Nates et al., 2017).

2.4.2 Monofilament testing results in sensitization

Unexpectedly, we determined that geckos with intact original tails (control group geckos) became, and remained, sensitized to repeated tactile testing. More specifically, following first exposure to the monofilaments, we found there was a significant decline in threshold at every site (at day 2), compared to day 0. This significant decrease (from baseline) was maintained throughout the experiment (with the exception of day 21 in the tail tip), but did not significantly change following day 2. A significant increase in sensitivity at day 2 was also seen in the hindlimbs, tail base, and tail tip of the tail regeneration group, and the hindlimbs of the tail loss group. Sensitization, the tendency towards increased responsiveness to stimuli, is often associated with avoidance or fear (Gotz and Janik, 2011). In contrast, habituation – the tendency towards decreased responsiveness to stimuli – is associated with non-stressful stimuli, and is a common outcome of many behavioral tests (Bolivar, 2009). Both sensitization and habituation are forms of non-associative learning, allowing organisms to ignore or prioritize stimuli

(Mcsweeney et al., 1996; Bolivar, 2009). As we observed that sensitization was almost global

(i.e., across multiple sites), it seems reasonable to predict that repeated testing affected the entire peripheral system and not just regional targets. Sensitization is often seen in fear-induced acoustic startle responses, resulting in increased response magnitude (Gotz and Janik, 2011), and hence is often correlated with fear or stress (Rau et al., 2005; Tanaka et al., 2013). Therefore, it is possible the geckos experienced stress during testing associated with handling and testing in the monofilament chamber. However, it is worth noting that all geckos continued to grow and eat throughout the experimental timeframe, and none demonstrated any other abnormal behaviors

indicative of stress, such as aggression, anorexia, or hyperactivity (Warwick et al., 2013).

Although some geckos did assume an upright (elevated) body posture while in the testing chamber (a recognized form of gecko anti-predation behavior; Landova et al., 2016), similar postures have also been observed during routine handling (not associated with monofilament testing).

Sensitization (and habituation) can occur within or between testing sessions (Bailey and

Chen, 1983). Therefore, it is possible that the observed sensitization to repeated testing of geckos was the result of hypersensitivity: from being tested at frequent intervals (McSweeney et al.,

1996). Another factor may have been the novelty of the testing environment/protocol. We did not expose geckos to the testing chamber prior to the start of the experiment, and exploratory behavior (associated with novel environments) has been correlated with higher degrees of sensitivity, or hypersensitivity (Honey et al., 2007). Alternatively, sensitization may be associated with the monofilament testing protocol mimicking a predatory encounter. In the wild, a sympatric predator of leopard geckos is the red sand boa, Eryx johnii. Sand boas are subterranean predators that employ a sit-and-wait strategy (Landova et al., 2015): they stay buried in the ground (e.g., sand), until prey comes nearby before striking. A recent behavioral study demonstrated that even captive-bred and predator-naïve geckos have an innate anti- predation reaction to the presence of sand boas (Landova et al., 2015). We postulate that monofilament testing may imitate the initiation of the sand boa encounter, which may explain why our geckos – including the control group – became sensitized. Overall, our data indicates that for most sites, sensitization is most likely a result of the monofilament assay itself, and not tail autotomy. Thus, a general level of sensitization exists in all groups.

2.4.3 Sensitization is attenuated in the forelimbs and exaggerated in the hindlimbs, following tail loss

Following tail autotomy, the forelimbs and hindlimbs demonstrated different trends in tactile sensitivity, especially between day 0 and day 2, corresponding to the wound healing phase of tail regeneration. Whereas the sensitivity of the forelimbs of autotomized geckos was largely unchanged following tail loss, the sensitivity of the hindlimbs was affected. Interestingly, while the hindlimbs were initially the least sensitive site tested, following autotomy it became significantly more sensitive even compared to controls. By day 7 post-autotomy, the differences between the control and tail autotomy groups were gone - both significantly more sensitive than baseline. In contrast, threshold of the forelimbs in the autotomized groups was not significantly altered from baseline until the tail was fully regenerated, at day 31. Overall, decreased sensitivity in the forelimbs and increased sensitivity in the hindlimbs are seen during the wound healing phase of tail regeneration. It is unclear whether this change is happening long enough to gain adaptive control (Cheadle et al., 2015), or whether this effect is a transient alteration to compensate in the interim (until the replacement tail is regrown). As the hindlimbs are less sensitive compared to other regions prior to autotomy, it is most likely easier to see sensitization effects compared to their baseline. It should be noted that deafferentation of limbs in humans leads to increased sensitivity and sensory acuity in neighboring areas (Bjorkman et al., 2009;

Lundborg et al., 2010). Based on proximity to the tail, the hindlimbs may be experiencing this effect in response to autotomy.

Discrepancies between the autotomized groups at day 7 (Figure 2.7) may be due to behavioral differences observed between the tail loss group and the tail regeneration group.

Though not the focus of this study, it was observed that geckos in the tail loss group were more

active following tail autotomy (days 2 and 7), than geckos from the tail regeneration group.

Individual variability seen in this study may have been heightened by the small sample size.

Future investigations plan to elucidate the role of behavior and its implications for tactile sensitivity.

Differential changes in tactile sensitivity between forelimbs and hindlimbs may correlate with biomechanical changes in posture and locomotion that occur following tail autotomy. Tail autotomy results in a cranial shift of the CoM, a decrease in peak vertical ground reaction force, and an increase in propulsive hindlimb ground reaction force (Jagnandan et al., 2014). As a result, there is a body-wide re-distribution of mass. In addition, there is a transient change in the knee angle of the hindlimbs (resulting in sprawled posture), but not at the elbow of the forelimbs

(Jagnandan et al., 2014). Concomitant changes in propulsive muscle recruitment and activation

(e.g., the caudofemoralis and gastrocnemius muscles) also occurs in the hindlimbs following autotomy (Jagnandan and Higham, 2018). However, these studies showed that there were no changes in the recruitment and activation of muscles of the forelimbs (e.g., the biceps and triceps brachii), or postural muscles of the hindlimb (e.g., the pubioschiotibialis) (Jagnandan and

Higham, 2018). Further, there was no change to running speed post-autotomy (Jagnandan et al.,

2014). Non-lizard studies have also shown that changes in mass lead to compensatory redistribution of load (Besancon et al., 2004; Munoz-Nates et al., 2017). For example, during human pregnancy, the substantial increase in anterior mass is compensated for by a proportional increase of opposing muscle force, such that overall gait patterns do not change (Mitternacht et al., 2013). In horses with forelimb lameness, increase mass and pressure are taken up by the opposite hindlimb (Weishaupt et al., 2006), while in quadrupeds carrying objects in their mouth

(rostral weight gain), the peak vertical force and foot pressure contact increases at the forelimbs

and decreases at the hindlimbs (Bockstahler et al., 2016). Importantly, studies investigating tactile sensitivity of the human foot have addressed the relationship between somatosensation and biomechanics. Reduced cutaneous input from the plantar surface of the human foot

(reflecting a decrease in discharge from plantar mechanoreceptors) is associated with changes in load and pressure distribution during locomotion, and modified patterns in walking (Eils et al.,

2002). Taken together with previously shown changes in biomechanics (loading and force) in the leopard gecko following autotomy (Jagnandan et al., 2014), it is likely that differential changes in tactile sensitivity of the hindlimbs and forelimbs confer biomechanical advantage so that parameters such as running speed may be maintained despite a drastic reduction in mass.

2.4.4 Tactile sensitivity is variable in the tail base, and unchanged in the tail tip, following tail loss

Some of the most unexpected findings of our investigation were observed at the tail base.

We determined that the tail base is the most sensitive site of interest prior to tail autotomy. Post autotomy, we found that the sensitivity of the tail base was strikingly variable between the tail loss and tail regeneration groups even though both cohorts underwent the same autotomy protocol. By way of explanation, we suggest that these differences are associated with the imperfect nature of spinal cord and peripheral nerve rupture. Unlike the skeleton and vascular system, there are no structural adaptations to the nervous system of the gecko tail that would minimize adjacent tissue damage following autotomy (McLean and Vickaryous, 2011; Delorme et al., 2012). Instead, the spinal cord and nerves are torn as the tail skeleton and associated structures give way (Duffy et al., 1992). Moreover, tail autotomies were induced by bilateral compression of the tail base (using pinching of the fingers), which may have resulted in unintended crush-like injuries to the nervous tissue. Although these injuries did not interfere with

spinal cord regeneration and axonogenesis in the long-term, they may have generated short-term symptoms such as muscle weakness and changes in sensation, creating inconsistencies between the tail loss and tail regeneration group. In order to test this hypothesis in the future, we propose removing the tails using a surgical amputation (done previously, Delorme et al., 2012), a more reproducible form of tail loss.

The sensitivity of the regenerated tail tip was never significantly different from original- tailed controls at any time point, indicating that tactile sensitivity is restored during tail regrowth.

Sensilla density of the regenerate gecko tail is documented to be much lower than corresponding segments on original tails (Russell et al., 2014), which is expected since tail loss is an energetically costly process, reducing mating and reproductive success (Dial and Fitzpatrick,

1981; Martin and Salvador, 1992). Curiously, tactile sensitivity between the original and regenerate tail as determined by monofilament testing, was comparable. This may be explained by functional roles of the regenerate tail, as it is still an ecologically and evolutionarily valuable appendage. Like the original tail, it is used for counterbalance (Jagnandan et al., 2014), but it also has comparably higher fat content. Leopard geckos specifically are a species that use their tails for fat storage, as a source of energy when food is scarce (Bustard, 1967). Lipid stores from the tail are also used for vitellogenesis (yolk deposition), in female geckos (Dial and Fitzpatrick,

1981; Doughty and Shine, 1998).

2.4.5 Differential changes in tactile sensitivity may be attributed to selective weighting of cutaneous feedback in the CNS, or altered skin capillary permeability

Broadly stated, our findings of differential changes in tactile sensitivity at the forelimbs and hindlimbs following the loss of the tail appendage, may be considered in the context of human amputation studies. The loss of an appendage results in a loss of afferent feedback from

that area, impacting functional recovery of movement. Differential changes in tactile sensitivity are often modulated through central processing changes, or selective weighting of cutaneous feedback in the CNS (Mugge et al., 2009; Strzalkowski et al., 2015). Following amputation of upper of lower limbs in humans, tactile sensitivity and acuity of both upper and lower residual limbs have been found to increase, compared to intact (control) limbs (Haber, 1955; Wu and

Kaas, 2002; Templeton et al. 2018). In general, amputation-mediated changes in sensitivity are associated with cortical reorganization, which may happen on a timescale of seconds (Haber,

1955; Merzenich et al, 1984; Makin et al., 2015). This is also seen in cases of hyperalgesia, where reversible changes in signal processing in the CNS render light tactile stimuli quite painful

(Torebjork et al., 1992).

An additional contributing factor involved in tactile sensitivity is skin capillary permeability and vasoconstriction (Edelberg, 1961). Tactile sensory nerves mediate, and in turn are mediated by, blood flow in the skin (Petrofsky, 2012). Vascular changes associated with stretching or relaxation of the skin, are known to alter tactile thresholds (Edelberg, 1961). This phenomenon is often seen in diabetic patients experiencing tissue ischemia, with reduced circulation to the skin resulting in decreased sensitivity of the hands and feet (Fromy et al.,

2002). Although the relationship between tactile sensitivity and vascular permeability in lizards remains unclear, body-wide hemodynamic changes have been documented during tail regrowth in the gecko, Hemidactylus flaviviridis (Hiradhar et al., 1978). In the future it would be worthwhile to establish whether changes in cutaneous blood flow (such as during tail autotomy and subsequent tail regeneration) influence or alter mechanoreceptor function.

2.4.6 Future Directions

This study established that there are regional differences in tactile sensitivity prior to tail loss, likely correlated to different functional roles of body areas in the gecko. Sensitization following autotomy was widely seen across experimental groups, which may be attributed to the monofilament assay itself. Tactile sensitivity of the regenerate tail tip following autotomy and regeneration was comparable to the original tail tip, whereas sensitivity of the tail base, was variable (likely due to nervous tissue injury following manual autotomy). The hindlimbs and forelimbs showed differential changes in tactile sensitivity, presumably conferring biomechanical advantage so that locomotor function may be maintained following a drastic loss in mass. Future work should investigate gecko behavior simultaneous with monofilament testing, to provide additional information related to sensitization, and functional changes related to altered tactile sensitivity.

CHAPTER 2 TABLES

Table 2.1 Monofilament buckling force values.

Calibrated in 2018 with Denver Instrument SI-603 scale. Grayed values were not used in this study.

Monofilament First Second Average Number Measurement (g) Measurement (g) (g) 3 0.094 0.088 0.091 4 0.133 0.124 0.1285 5 0.175 0.154 0.1645 6 0.258 0.256 0.257 7 0.383 0.355 0.369 8 0.575 0.534 0.5545 9 1.15 1.088 1.119 10 2.381 2.221 2.301 11 3.16 3.3 3.23 12 6.395 6.03 6.2126 13 11.5 11.6 11.55 14 18 18.4 18.2 15 40.3 39.9 40.1 16 50.2 48.9 49.55 17 66.1 68.1 67.1

Table 2.2 Tactile sensitivities across the gecko body: raw monofilament threshold and proportion of threshold.

DAY Control Tail Loss Tail Regeneration HINDLIMBS Threshold (g) % of baseline Threshold (g) % of Baseline Threshold (g) % of Baseline D0 0.19503125 100 0.157575 100 0.19565 100 D2 0.1593625 82.8 0.117025 74.5 0.1161 57.1 D7 0.13859375 74.8 0.118675 75.4 0.14645 71.5 D14 0.1293125 67.3 n/a n/a 0.1168 57.3 D21 0.13390625 70.7 n/a n/a 0.13275 64.9 D31 0.1409375 74.9 n/a n/a 0.1503 77.5 FORELIMBS Threshold (g) % of baseline Threshold (g) % of Baseline Threshold (g) % of Baseline D0 0.14525 100 0.11055 100 0.1336 100 D2 0.100375 69.6 0.107725 97.5 0.119775 91.2 D7 0.1095625 76 0.113275 102.5 0.10585 79.7 D14 0.10028125 69.6 n/a n/a 0.106 80.1 D21 0.10496875 72.6 n/a n/a 0.107875 81.6 D31 0.09803125 67.6 n/a n/a 0.1003 75.4 TAIL BASE Threshold (g) % of baseline Threshold (g) % of Baseline Threshold (g) % of Baseline D0 0.128125 100 0.0985 100 0.11335 100 D2 0.1001875 78.2 0.0985 100 0.091 80.3 D7 0.1001875 78.2 0.091 92.4 0.091 80.3 D14 0.0956875 74.7 n/a n/a 0.091 80.3 D21 0.0956875 74.7 n/a n/a 0.1021 90.1 D31 0.0956875 74.7 n/a n/a 0.091 80.3 TAIL TIP Threshold (g) % of baseline Threshold (g) % of Baseline Threshold (g) % of Baseline D0 0.1395 100 n/a n/a 0.1375 100 D14 0.1050625 75.3 n/a n/a 0.091 66.2 D21 0.11875 85.1 n/a n/a 0.09475 68.9 D31 0.10975 78.7 n/a n/a 0.091 66.2

Threshold values measured in grams (g).

CHAPTER 2 FIGURES

Figure 2.1. Schematic illustration of experimental design for monofilament assay.

Black vertical ticks correspond to monofilament testing days. Gray vertical ticks correspond to euthanasia timepoints. Each group followed the same timeline (with the tail loss group being euthanized on day 8), with all geckos in the same group tested (or euthanized) on the same day.

All testing was performed within 1.5 months.

Figure 2.2. Locations of the sites of interest for monofilament testing.

White circles denote regions across the ventral surface of the body of the leopard gecko that were tested for tactile sensitivity. Sites included: the left and right hindlimbs (plantar surface of the pes), left and right forelimbs (palmer surface of the manus), and the (proximal) base and (distal) tip of the tail. The distal tip of the tail was first tested for original tails (for baseline measurements of all groups; all timepoints for control geckos), and then tested for regenerate tails 14 days following autotomy (for the tail regeneration group). Monofilaments were applied perpendicular to the skin in an area roughly corresponding to the middle of each site of interest.

Figure 2.3. Monofilament testing chamber and a representative Semmes-Weinstein monofilament.

(A) The monofilament chamber: a clear plexiglass chamber (31.5cm wide, 29.5cm high, 31.5cm deep) mounted on a raised, perforated (~10mm diameter) platform. A mirror suspended at an oblique angle below the perforated platform was used to direct the monofilaments through the perforations. A black partition (not shown) covered the front of the chamber, to blind the gecko to the presence of the experimenter. A video camera (Panasonic SDR-H85) was mounted outside the chamber to record the trials. Application of a monofilament through a perforated hole is shown. (B) A leopard gecko within the chamber, standing on the perforated platform. The ventral surface of the body is reflected in the mirror below. Geckos were acclimated to the testing chamber for one hour prior to the start of each monofilament testing session. (C) A

Semmes-Weinstein monofilament, with tip diameter corresponding to a specific force in grams

(Table 2.1). Each monofilament was applied perpendicular to the skin until buckling.

Figure 2.4. Ascending stepwise monofilament method.

The monofilament assay was performed following the protocol of Field et al. (1997), using an ascending stepwise approach. Testing began with the lightest weight monofilament, applied to the site of interest until buckling. A positive response from the geckos consisted of a withdrawal, lifting of either the foot (when testing the forelimbs and hindlimbs), or body/tail (when testing the tail). If no withdrawal response was observed, the application was deemed negative, and the next largest monofilament was applied. If positive, the response was recorded and a re-test was done. The average of the two thresholds was taken to be the lowest sensory response. Flow chart adapted from Bradman et al. (2015).

Figure 2.5. Regional sensitivity differences at baseline.

Raw monofilament data (measured in grams) at each site at day 0 of monofilament testing, reported as means and standard errors. There was a main effect of site. The hindlimbs were significantly less sensitive than the forelimbs (p=0.005) and tail base (p=0.003). Experimental groups were pooled for analysis, presented as means of 14 geckos with standard errors. Data was analyzed using a one-way repeated measures ANOVA, and post hoc comparisons using Fisher’s

LSD test. Asterisks (*) denote significance.

Figure 2.6. Changes in hindlimb sensitivity with respect to baseline following tail loss and tail regeneration.

Tactile threshold of the hindlimbs of experimental groups over time with respect to baseline, reported as means (n=4, control; n=5, tail loss; n=5, tail regeneration) and standard errors. There was a main effect of time. Hindlimb threshold significantly decreased in all three groups compared to baseline (except for the control group at day 2). At day 2, threshold of the tail regeneration group was significantly lower than the control group (p=0.023). Statistical analysis was done using a two-way repeated measures ANOVA, and post hoc comparisons using Fisher’s

LSD test. Asterisks (*) denote significant differences between groups. Letters (‘a’ control, ‘b’

tail loss group, ‘c’ tail regeneration group) denote significant differences from respective baselines. Phases of tail regeneration are shown below the x axis.

Figure 2.7. Changes in forelimb sensitivity with respect to baseline following tail loss and tail regeneration.

Tactile threshold of the forelimbs of experimental groups over time with respect to baseline, reported as means (n=4, control; n=5, tail loss; n=5, tail regeneration) and standard errors. There was a main effect of group and time. At day 2, forelimb threshold of the control group was significantly lower than baseline (p=0.003), but the tail loss and tail regeneration group were not.

At day 7, forelimb threshold was significantly different between the control and tail loss group

(p=0.006) and significantly different between the tail loss and tail regeneration group (p=0.02).

While controls were significantly lower than baseline for the duration of the experiment, tail regeneration was not until day 31. Statistical analysis was done using a two-way repeated measures ANOVA, and post hoc comparisons using Fisher’s LSD test. Asterisks (*) denote significant differences between groups. Letters (‘a’ control, ‘b’ tail loss group, ‘c’ tail

regeneration group) denote significant differences from respective baselines. Phases of tail regeneration are shown below the x axis.

Figure 2.8. Changes in tail base sensitivity with respect to baseline following tail loss and tail regeneration.

Tactile threshold of the tail base of experimental groups over time with respect to baseline, reported as means (n=4, control; n=5, tail loss; n=5, tail regeneration), and standard errors. There was a main effect of time and group. Tail base threshold significantly decreased compared to baseline for the controls and tail regeneration group for all timepoints (except day 21 for the tail regeneration group). At day 2, threshold of the tail loss group was significantly higher than the control group (p=0.005), and also significantly higher than the tail regeneration group (p=0.009).

At day 7, threshold of the tail loss group was significantly higher than the control group

(p=0.05). Tail regeneration groups and control groups were significantly different at day 21

(p=0.05). Statistical analysis was done using a two-way repeated measures ANOVA, and post

hoc comparisons using Fisher’s LSD test. Asterisks (*) denote significant differences between groups. Letters (‘a’ control, ‘b’ tail loss group, ‘c’ tail regeneration group) denote significant differences from respective baselines. Phases of tail regeneration are shown below the x axis.

Figure 2.9. Changes in tail tip sensitivity with respect to baseline following tail loss and tail regeneration.

Tactile threshold of the tail tip of experimental groups over time with respect to baseline, reported as means (n=4, control; n=5, tail regeneration) and standard errors. There was a main effect of time and group. Tail tip threshold significantly decreased compared to baseline for the controls and tail regeneration group for all timepoints. Testing of the regenerated tail tip began on day 14 when the replacement tail was long enough to be tested. Statistical analysis was done using a two-way repeated measures ANOVA, and post hoc comparisons using Fisher’s LSD test.

Asterisks (*) denote significant differences between groups. Letters (‘a’ control, ‘b’ tail loss group, ‘c’ tail regeneration group) denote significant differences from respective baselines.

Phases of tail regeneration are shown below the x axis.

CHAPTER 3: CHARACTERIZATION OF PURKINJE CELL MORPHOLOGY

FOLLOWING TAIL LOSS AND REGENERATION IN THE LEOPARD GECKO

(EUBLEPHARIS MACULARIUS)

3.1 INTRODUCTION The cerebellum is a morphologically variable region of the hindbrain best known for its roles in fine-tuning motor actions and error control (Anderson, 1993; Requarth and Sawtell,

2014; Baumann et al., 2015). Across vertebrates, the cerebellum varies in shape, from a small mound with a row of putative cerebellar cells in the lamprey, to the highly folded (foliated) structure seen in birds and mammals (Voogd and Glickstein, 1998; Butz et al., 2014; Jacobs et al., 2014). Amongst non-avian reptiles, the cerebellum is often described as resembling a smooth dome or sheet-like structure (Larsell, 1926; Voogd and Glickstein, 1998; Wylie et al., 2016).

Notwithstanding its gross morphological diversity, the cellular organization of the cerebellum are remarkably conserved (Bangma and Ten Donkelaar, 1982; Voogd and Glickstein,

1998). At the level of histology, the cerebellum is organized into three main layers: the molecular layer, the Purkinje cell layer, and the granular layer (Butts et al., 2014). The molecular layer is comparatively cell-sparse, but includes populations of stellate cells and basket cells, while the granular layer is cell-dense, and includes abundant glutamatergic granule neurons and interneurons (including Golgi, Lugaro, unipolar brush cells) (Dine et al., 2013; Badura and De

Zeeuw, 2017). Between the two is the Purkinje cell layer, dominated by large, gamma-aminobutyric acid (GABA)-ergic Purkinje cells. Although typically arranged in a monolayer in mammals (Anderson and Korbo, 1993; Jacobs et al., 2014), the organization of

Purkinje cells is often less structured among non-avian reptiles, amphibians, and teleost fish,

including scattered cells (in some snakes, and fish) or multiple layers (in some lizards) (Larsell,

1926; Ten Donkelaar, 1998).

The cerebellum integrates with visual, vestibular, and somatosensory systems, to regulate motor actions, and may also play a role in cognition and motor learning (Black et al., 1990;

Konnerth et al., 1990; Hoogland et al., 2015; Takakusaki, 2017). Sensory input to the cerebellar cortex comes from mossy (originating from many neuronal sources) and climbing fibers

(originating from the medulla oblongata), which innervate granule and Purkinje cells, respectively (Konnerth et al., 1990; Dusart and Flamant, 2012). Each Purkinje cell receives multiple synaptic contacts from a single climbing fiber (Dusart and Flamant, 2012). Each climbing fiber wraps around the Purkinje cell soma, as well as the smooth (i.e., lacking dendritic spines) proximal dendrites, to provide strong excitatory input (Mauk et al., 1997; Voogd and

Glickstein, 1998). The second primary afferent input, the mossy fibers, synapse on granule cells

(Dusart and Flamant, 2012). Granule cell axons then pass into the molecular layer and split into

‘T’- shaped horizontal branches called parallel fibers (Voogd and Glickstein, 1998). These parallel fibers weave within the dendritic arbors of Purkinje cells, making numerous (up to 200

000 per Purkinje cell in mammals) weakly excitatory synapses (Hoxha et al., 2016). Purkinje cells serve as the sole output of the cerebellar cortex, and pass inhibitory projections to the deep cerebellar nuclei, located in the white matter of the cerebellum (Rondi-Reig et al., 2014).

Neurons of the deep cerebellar nuclei then send projections to the brainstem or cerebral cortex via the thalamus (Konnerth et al., 1990).

Purkinje cells are amongst the largest and most distinctive cell types of the nervous system. First described by Purkinje in 1837 (and later by Golgi, 1886), Purkinje cells are characterized as having large cell bodies (or somas), and an elaborate, highly complex or

arborized arrangement of dendrites (Anderson and Korbo, 1993; Voogd and Glickstein, 1998).

Among reptiles, Purkinje cells are typically described as having a long primary dendrite, with shorter secondary and (sometimes) tertiary branches, terminating in spiny branchlets

(Nieuwenhuys, 1967; Bangma and Ten Donkelaar, 1982; Chan and Nicholson, 1986). Dendrites closest to the soma typically lack dendritic spines, and are referred to as being smooth; in more distal dendrites, spines are often densely organized (Voogd and Glickstein, 1998). Collectively,

Purkinje cells receive more synaptic input than any other type of cell in the brain, and are known to be capable of changing the morphology and functional attributes of their dendrites - features that are thought to facilitate cerebellar plasticity (Floeter and Greenough, 1979; Komandatov and

Ascoli, 2009; Tavosanis, 2011).

Although dendrites were once viewed as playing a passive role in synaptic signaling, it is now understood they are capable of conducting input signals, back-propagating action potentials, integrating synaptic input, and reshaping network connectivity (Eilers and Konnerth, 1997;

Ascoli et al., 2001). As a result, changes in dendritic morphology have been correlated with changes in neurophysiology, which in turn is modulated (in part) by changes in intracellular calcium concentrations (Kater et al., 1988; Eilers and Konnerth, 1997; Anwar et al., 2014).

Neurotransmitters alter postsynaptic membrane potentials, impacting voltage-dependent calcium channels, which in turn regulate dendritic growth cone behavior (Zheng et al., 1994; Murchison et al., 2002; Butz et al., 2009). Among Purkinje cells, changes in dendritic morphology have been associated with a wide range of stressors, including alterations in social environment, exercise, age, neurodevelopmental disorders, and chemical treatments (Woodward et al., 1975;

Floeter and Greenough, 1979; Lein et al., 2007). This plasticity is predicted to permit adaptations in response to changes in their internal and external environment, which includes motor changes

and motor learning (Anderson, 1993; Jorntell and Hansel, 2006; Sdrulla and Linden, 2007).

Whether this adaptive response extends to other forms of biological perturbation remains unexplored.

Changes in body mass are a common form of biological perturbation experienced by many organisms. These include alterations in internal (e.g., pregnancy) or external physical loading (carrying offspring or external loads, space flight), body-wide ontogenetic growth, or the discrete development (e.g., antlers) or loss (e.g., amputation) of specific structures of specific structures (Makin et al., 2013; Lowrey et al., 2014; Jagnandan and Higham, 2018). Mass loss following amputations is a particularly traumatic example of perturbation that has profound implications at the level of physiology, behavior, and locomotion (Haber, 1955; Makin et al.,

2013; Templeton et al., 2018). These include sensory deprivation from the amputated region, altered limb coordination, and a variety of neuromorphological and neurophysiological changes

(Wu and Kaas, 2002; Chen et al., 2012; Makin et al., 2013). Interestingly, some species employ a specialized form of amputation, known as autotomy, as a means to avoid predation. Among vertebrates, the best-known example of autotomy comes from lizards (Jacyniak et al., 2017).

Many species of lizard are able to self-detach, or voluntarily amputate, a portion of the tail to escape capture (Cromie and Chapple, 2013; Gilbert et al., 2013; Gillis and Higham, 2016;

Jacyniak et al., 2017). Perhaps not surprisingly, tail autotomy impacts how lizards move. For example, investigations using the lizard Eublepharis macularius, (the leopard gecko; hereafter,

‘gecko’), determined that tail autotomy results in a near instantaneous loss of 1/3 of the lizard’s total length, and ¼ of its total body mass (Jagnandan et al., 2014). As a consequence, there is a cranial shift in center of mass (CoM), which in turn impacts body orientation, postural stability, and locomotor performance (Jagnandan et al., 2014). Curiously, hindlimb and forelimb

kinematics do not appear to change follow tail autotomy (Jagnandan et al., 2014), indicating the participation of one or more as-of-yet undetermined short-term compensatory mechanisms.

For most lizards, tail autotomy is matched by the spontaneous ability to regenerate a replacement. In geckos, tail regeneration takes approximately 30 days (Mclean and Vickaryous,

2011). Although the newly regenerated tail is superficially similar to the original organ it differs in various structural details, including the organization of skeletal muscle and the tissue composition of the skeleton (e.g., bony vertebrae are replaced by a cartilaginous cone) (McLean and Vickaryous, 2011; Delorme et al., 2012). Further, the CoM is never fully restored to its original (baseline) position (Jagnandan et al., 2014). Adjustments to long-and short-term motor changes is known to be elicited by the cerebellum in the form of multi-limb coordination (Veloz et al., 2015). Although a role for the cerebellum in maintaining fine motor coordination before and after autotomy, and throughout tail regeneration is strongly indicated, to date little is known.

Here, we used a modified Golgi-Cox method to provide the first ever neuromorphological characterization of gecko Purkinje cells. To confirm our results, we performed a parallel investigation on CD-1 mice, a multipurpose murine model whose Purkinje cell morphology is well documented (Soha and Herrup, 1995; Billeci et al., 2010; Dusart and

Flamante, 2012). Next, we sought to investigate if the neuromorphology of gecko Purkinje cells changed in response to tail autotomy. We reasoned that since tail autotomy results in significant changes in gecko CoM, it may also induce neuromorphological alterations in the overall size, shape, and complexity of the Purkinje cell dendritic tree. We determined that there was no overall change in the morphology of Purkinje cells following tail autotomy. However, we did find that there were significant changes in the medial-distal segments of the dendritic arbor – regions corresponding with the distribution of parallel fibers. We predict that plasticity following

tail autotomy may be restricted to the arrangement and distribution of synaptic connections in the cerebellum, facilitated through isolated changes in neuromorphology of the dendritic tree of

Purkinje cells.

3.2 MATERIALS AND METHODS

3.2.1 Experimental Animals and Animal Care

Captive bred leopard geckos (E. macularius) were obtained from a commercial supplier

(Global Exotic Pets, Kitchener, Ontario, Canada). All geckos were subadults, less than one year in age and had an average body mass of 20.2g (17.1-22.4g). For the main experiments (tactile sensitivity [Chapter 2] and neuromorphology [Chapter 3], we used a cohort of 15 geckos; for the serial histology characterization, we used a separate group of three subadult geckos. Growth of all geckos was tracked weekly throughout the experimental period by measuring snout-vent length, tail length, and body mass. Animal Utilization Protocols (AUPs) were approved by the

University of Guelph Animal Care Committee (Protocols #1954, 3772) and followed the guidelines of the Canadian Council on Animal Care. Animal husbandry follows the work of

McLean and Vickaryous (2011). Gecko colonies were housed in a secure vivarium (Hagen

Aqualab facility) at the University of Guelph, inside a temperature controlled environmental chamber (average ambient temperature, 27.5 ̊C). Geckos were housed individually in 5 galleon polycarbonate tanks, with a heat cable (Hagen Inc., Baie d’Urfé, Québec, Canada) set at 32ºC and placed underneath one side of the enclosure to establish a thermal gradient. Each enclosure included two hide boxes and a water dish. Geckos had constant access to fresh drinking water, and were fed 3 larval mealworms (Tenebrio spp.) daily, dusted with powdered calcium and vitamin D3 (cholecalciferol; Zoo Med Laboratories Inc., San Luis Obispo, California, USA). The

gecko chamber was kept on a year-round 12h light:12h dark photoperiod, and all experiments were conducted in a 6-week span from November-December.

CD1 strain mice were obtained as part of a collaboration with Dr. C. Bailey, University of Guelph. Mice were purchased from Charles River Canada (Saint-Constant, Quebec, Canada), and maintained within a secure vivarium (Central Animal Facility, Guelph) at the University of

Guelph (average ambient temperature of 22.5ºC). AUP #3622 was approved by the University of

Guelph Animal Care Committee, and closely follows the guidelines of the Canadian Council on

Animal Care. Animal husbandry and breeding follows the work of Chung (2018). Pups were collected at postnatal day 14 (the day of birth for the litter representing postnatal day 0). Mice had consistent ad libitum access to food and water, and were kept on a 12-hour reverse light cycle.

3.2.2 Experimental Design

Both the tactile sensitivity and neuromorphological experiments (Chapters 2 and 3) used the same cohort of 15 geckos. These geckos were randomly assigned into one of three groups

(n=5 per group): control (tail intact); tail loss (8 days post-autotomy); and tail regeneration (31 days post autotomy). The experimental timeline was either eight days (for the tail loss group) or

31 days (for the control and tail regenerated groups) (Figure 3.1). Autotomy for both the tail loss and tail regenerated groups was induced on day 1 of the experiment; control geckos retained their original tails for the duration of the experiment. Each gecko was used for both tactile sensitivity (Chapter 2) and neuromorphological evaluation (Chapter 3) following euthanasia.

Mice were used an internal control for the neuromorphological experiment.

3.2.3 Tail Autotomy

Tail autotomy is a naturally evolved anti-predation strategy common to many species of lizard. Tail autotomy was performed by manually restraining the gecko, and then pinching the base of the tail between the forefinger and thumb. The act of pinching induces the tail to self-detach, after which the newly tailless gecko was released back into its enclosure. All geckos survived the autotomy procedure and all autotomies occurred at roughly the same point along the tail (the second proximal fracture plane from base), corresponding with a maximal tail loss.

Control geckos were manually restrained but did not have their tails removed.

3.2.4 Euthanasia, Tissue Collection, and Preparation

Whole brains were then dissected out of the skull, and further fixed in 10% NBF for 22 hours.

Tissues were then transferred to 70% ethanol until processing. Geckos collected for Golgi-Cox staining were immediately dissected, the brains removed (without fixation), and placed into

Golgi-Cox solution. Mice were anesthetized using 5% isoflurane and subsequently decapitated, with brains immediately removed and placed in Golgi-Cox solution.

Fixed tissues collected for histochemistry and immunofluorescent procedures were prepared using an automated processor (Fisher Scientific, Waltham, Massachusetts, USA).

Briefly, tissues stored in 70% ethanol were dehydrated to 100% isopropanol, cleared in xylene, and then embedded in paraffin wax. Sectioning was done using a rotary microtome (Shandon

Finesse ME+, Thermo Fisher Scientific), producing sections 5μm thick which were then

mounted on charged slides (Surgipath X-Tra; Leica Microsystems, Ontario, Canada). Prior to histology or immunofluorescence procedures, slides were incubated overnight at 37ºC or 60ºC, respectively.

3.2.5 Golgi-Cox Impregnation

To visualize neurons, we used the Golgi-Cox method, modified from the work of Louth et al. (2017). Briefly, whole brains (gecko and mouse) were dissected out of the skull immediately following euthanasia, and placed in a scintillation vial (covered with aluminum foil) filled with Golgi-Cox solution (1% (w/v) potassium dichromate, 0.8% (w/v) potassium chromate and 1% (w/v) mercuric chloride). The brains were incubated in the Golgi-Cox solution, in the dark, for 29 days at room temperature. At the end of the incubation period, brains were transferred to a scintillation vial containing sucrose cryoprotectant (30% (w/v) sucrose in 0.1 M phosphate buffer, pH 7.4) for 72 hours at 4°C in the dark. The recipe and protocol for Golgi-Cox staining are found within Appendix II.

3.2.6 Golgi-Cox Sectioning

Prior to sectioning, brains were removed from the sucrose cryoprotectant solution and trimmed using a razor blade. Brains were then embedded in longitudinal orientation - lateral side down in molten agar (3% (w/v) in water heated for 50 seconds using a microwave) in a small weigh dish. Once the agar had cooled, the resulting block was trimmed to leave ~2 mm solid agar surrounding the brain, and then glued onto the stage of the vibratome (Leica VT1000S) using ethyl cyanoacrylate glue. The vibratome stage area was then filled with enough sucrose cryoprotectant to cover the brain. The brain/agar block was then sliced at a thickness of 400μm

(blade advancement speed 0.125mm/s, vibration frequency 86Hz). Sagittal brain sections were picked up with a small paint brush and placed into a well of a 6-well tissue culture plate, with

mesh-bottom inserts (3 brain slices per well). Wells were pre-filled with 10mL of sucrose (6%

(w/v)) in 0.1M phosphate buffer, pH 7.4. Sections were then incubated in the dark at 4ºC overnight.

3.2.7 Golgi-Cox Brain Section Processing

Brain sections were transferred into new wells containing 5mL of 2% (w/v) paraformaldehyde in 0.1 M phosphate buffer, using the removable mesh-bottom inserts, and incubated on a rocker (moderate speed [20rpm]) in the dark for 15 minutes. Sections were then washed twice by transferring them to new wells containing 5mL of water (slow speed [15rpm]; 5 mins each), then new wells containing 5mL of 2.7% (v/v) ammonium hydroxide, incubated in the dark (moderate speed [20rpm]; 15 minutes), followed by two water washes (slow speed

[15rpm]; 5 mins each) in new wells. Next, the brain sections were transferred into new wells with 5mL of Kodak Fixative A (recipes in Appendix II), diluted 10x in water, and incubated in the dark (moderate speed [20rpm]; 25 minutes), before two final washes in water (slow speed

[15rpm]; 5 mins each) in new wells.

3.2.8 Golgi-Cox Mounting and Dehydrating Brain Sections

Brain sections were mounted onto Superfrost Plus microscope slides using a small paintbrush. Excess agar was removed using tweezers, and excess water wicked away using

Kimwipes. Sections were air dried at room temperature, for either 45 minutes (geckos) or 60 minutes (mice), then dehydrated by placing them into Coplin staining jars containing a graded ethanol series (2 minutes in each of 50%, 75%, and 95%, followed by two 5 minute washes in

100% ethanol) followed by 5 minute of clearing in the organic solvent Citrisolv (Fisher

Scientific, 22-143-975). Slides were then coverslipped using Permount (Fisher Scientific, SP15-

100) an anhydrous mounting medium, and dried for 5 days in the dark.

3.2.9 Microscopy and Imaging Stained Neurons

Histological and immunofluorescent sections were visualized using the 20x and 40x objectives of an Axio Imager D1 Microscope (Carl Zeiss Canada Ltd., Toronto, Ontario,

Canada). To visualize Purkinje cells, we used brightfield microscopy, using a BX53 Olympus microscope (12-bit color camera, MBF Biosciences, Williston, Vermont, USA), and Neurolucida software (version 10, MBF Bioscience). High-resolution z-stack images of neurons of interest were acquired using the 30x 1.05 N.A. silicone oil-immersion objective. The step distance between images in the stack was set to 1μm, and overlapping stacks in the x and y-axes were stitched together in three-dimensions. For each gecko cerebellum, all Purkinje cells within a given brain section were traced, provided they met the following selection criteria: (1) the soma was located in Purkinje cell layer, 20μm above or below the junction of the molecular and granular cell layers; (2) dendrites and somata were fully contained within the section (i.e., not passing out of the section and therefore truncated); (3) individual Purkinje cells were reasonably isolated from adjacent neurons (i.e., overlap was minimal); and (4) the dendrites projected into the molecular layer, with a characteristic Purkinje cell arborization. This resulted in a gecko control group (n=3), tail loss group (n=5), and tail regeneration group (n=4). Neuron averages per individual are available in Appendix IV, Table A8. For the mice cerebella, two Purkinje cells were randomly sampled from each brain (from two individuals, n=2), using the same selection criteria outlined above. Statistical analyses were not performed on mouse Purkinje cells – they were qualitatively described in this study. Species comparisons were based on numbers present and not statistically sound.

3.2.10 Histology: Hematoxylin and Eosin

To visualize the tissue architecture of the cerebellum, representative sections of the hindbrain were stained with hematoxylin and eosin. Slides were deparaffinized with three xylene washes (2 minutes each), then rehydrated to water through three washes of absolute isopropanol

(2 minutes each), one wash of 70% isopropanol (2 minutes), and one wash in deionized water

(dH2O) for 2 minutes. Slides were then immersed in Harris Hematoxylin (Fisher Scientific,

Waltham, Massachusetts, USA) for 10 minutes. Alternating rinses with dH2O accompanied 5 dips in acid alcohol (1% hydrochloric acid in 70% isopropanol), dips in ammonia water (until blue), and 6 dips in 70% isopropanol. Slides were then immersed in eosin for 1 minute, before being subjected to four washes of absolute isopropanol (2 minutes each), three washes of xylene

(2 minutes each), and finally coverslipped using Cytoseal (Fischer Scientific, Waltham,

Massachusetts, USA). Slides were then allowed to dry overnight. The recipe and protocol for hematoxylin and eosin staining are found within Appendix II.

3.2.11 Immunofluorescence

A two-day immunofluorescence protocol was used to visualize calbindin, a calcium-binding characteristic of Purkinje cells. On day one, slides were deparaffinized with three xylene washes (2 minutes each), then rehydrated to water through three washes of absolute isopropanol (2 minutes each), one wash of 70% isopropanol (2 minutes), and one wash in dH2O

(2 minutes). They were then immersed in 1x phosphate buffered saline (PBS) for 15 minutes.

Following this, they underwent heat-induced antigen retrieval in citrate buffer at 95˚C for 12 minutes, with a 20-minute cooling period. Slides were then washed with PBS (2 minutes), before undergoing further antigen retrieval by being treated with 0.1% trypsin (Sigma-Aldrich, St.

Louis, Missouri, USA), diluted with sterile PBS at a temperature of 37ºC for 20 minutes.

Following another wash with PBS, slides were blocked for one hour with 5% normal goat serum

(NGS), mixed with diluent (1% BSA, 0.5% Tween 20 [Sigma-Aldrich, Oakville, Ontario,

Canada], 0.1% sodium azide [Fisher Scientific, Waltham, Massachusetts, USA] in PBS) at a temperature of 37ºC for 30 minutes. Next, slides were incubated in primary antibody mixed with diluent, at 4ºC overnight (mouse anti-calbindin [1:50] Sigma-Aldrich, St. Louis, Missouri, USA).

One section on each slide served as a negative control and was incubated in only diluent without primary antibody.

On day two of the protocol, sections were washed in PBS three times (two minutes each), before being incubated with secondary antibody (Alexa Fluor-488 labeled goat anti-mouse for calbindin [1:100]) diluted in sterile PBS, at room temperature for one hour. The slides were then again washed in PBS three times (two minutes each), before applying DAPI nuclear stain (Life

Technologies, Eugene, Oregon, USA) diluted in sterile PBS [1:5000], for two minutes. Slides were then washed in PBS three final times (2 minutes each), and then coverslipped with fluorescent mounting medium (DAKO, Glostrup, Denmark). The recipe and protocol for immunofluorescence is found within Appendix II.

3.2.12 Sholl and Branch Analyses Data were generated using Neurolucida Explorer software (MBF biosciences). For each gecko, one to six Purkinje cells within each cerebellum were traced, and the neuromorphological measurements were averaged for each individual (Appendix IV, Figure A1, Table A8). In 3 geckos, no Purkinje cells meeting the selection criteria (outlined above) were observed. Data are presented as mean ± 1 SEM, with three to five geckos in each group. To quantitatively analyze the neuromorphology of the Purkinje cells for both mice and control (i.e., tail intact) geckos, we generated three-dimensional representations of select neurons. Modified three-dimensional Sholl analyses (15μm intervals between nested spheres radiating from the cell soma) were performed

to measure number of dendritic intersections and average dendritic diameter. To quantitatively analyze the neuromorphology of the Purkinje cells for both species, we used Neurolucida software to evaluate several key metrics of dendritic complexity: total length of all dendrite matter associated with the neuron; dendrite volume (the total dendrite diameter and surface area); the number of dendritic terminals, and the length of the furthest terminal (indicative of neuron size).

3.2.13 Statistical Analysis Statistical analyses for the Purkinje cell neuromorphological data were performed using

GraphPad Prism 7 (GraphPad Software, La Jolla, CA). Sholl data were analyzed using a two-way repeated measures ANOVA to assess main effects of distance from the soma and tail loss and/or regeneration groups, and potential interaction between these two variables. Branch structure analyses were analyzed using a one-way ANOVA. Significance level for all analysis was determined as p< 0.05. All post hoc analyses examined pair wise comparisons using the

Bonferroni method. Statistical output for Sholl analyses is shown in Appendix V, Figure A10.

3.3 RESULTS 3.3.1 Experimental Geckos

shared a similar experimental timeline but with two different euthanasia endpoints (Figure 3.1).

The tail loss group was euthanized on day 8 of the experiment, whereas the tail regeneration and control groups were euthanized on day 31. By the end of the experiment, the average masses for each of the groups were: 24.8g (control), 19.6g (tail loss), and 21.8g (tail regeneration).

Snout-vent body lengths for all groups closely paralleled on another throughout the experiment, with an average length of 105mm (95-110mm) for the control and tail regeneration groups on day 31 (Appendix I, Table A2). At day 8 of the experiment, geckos in the tail loss group had yet to begin regenerating their tails, and either retained the clot covering the site of autotomy

(representing stage II of regeneration [Mclean and Vickaryous, 2011] or the clot was lost but no new outgrowth had occurred (stage III). At day 31 of the experiment, all geckos in the tail regeneration group had fully formed new tails (stage VII).

3.3.2 Organization of the Cerebellar Cortex

The cerebellum is located dorsal to the fourth ventricle, caudal to the optic tectum, and rostral to the medulla of the brainstem (Figure 3.2A,B). Similar to birds and mammals, the reptilian cerebellar flocculus (or auricles) extends laterally, along with the cerebellar peduncle

(Voogd and Glickstein, 1998). As with other reptiles, the gecko cerebellum lacks foliation, and instead resembles a simple, unfolded sheet (Figure 3.2A,B). Similar to most lizard species, the gecko cerebellum is everted, or tilted towards the optic tectum. As a result, the granular layer is positioned dorsally. The grey matter of the cerebellar cortices has a highly conserved three-layered arrangement (Figure 3.3A,B). Purkinje cells have large soma that are conspicuously immunopositive for calcium-binding protein, calbindin, and loosely arranged in a layer 1-3 cells deep, at the junction between the molecular and granular cell layers (Figure

3.4A,B). In sagittal sections, the granular layer is 150-400μm thick, while the Purkinje cell layer is 25-75μm thick, and the molecular layer spans 300-600μm thick.

3.3.3 Purkinje Cell Visualization

To characterize the neuromorphology of Purkinje cells, we used Golgi-Cox staining, on

400um thick sections. Cerebella from geckos were collected following monofilament testing

(Chapter 2), with each brain yielding six tissue slices, all of which were analyzed (Figure 3.5A).

Cerebella from mice were collected without monofilament testing (Figure 3.5B). In addition to

Purkinje cells, Golgi-Cox staining visualized: astrocytes and stellate cells (within the molecular layer); basket cells (adjacent to the Purkinje cell layer) and climbing fibers and Bergmann glia

(projecting into the molecular layer).

Using Neurolucida software, we created three-dimensional Z-stacks of Purkinje cells, 40-

60μm deep (Figure 3.6A,B). Purkinje cells of both mice and geckos are characterized by a large soma (~400μm2). Gecko Purkinje cells have a single primary dendrite, whereas those of mice may have one or two. All dendrites of all Purkinje cells pass into the molecular layer before ramifying to the characteristically elaborate pattern of dendritic branching. Among geckos, the primary dendrite of the Purkinje cell gives rise to comparatively shorter secondary and tertiary branches, with branching occurring deeper in the molecular layer than compared to the pattern observed in mice. In both geckos and mice, proximal areas of the dendritic arbor are smooth

(lacking spines) whereas more distal segments seem are covered in spines. The spines of gecko

Purkinje cells were notably smaller than that of the mouse.

3.3.4 Species Differences in Purkinje Cell Neuromorphology

Overall, the mouse Purkinje cell data collected in this study were consistent with previously published findings of CD-1 mice (Soha and Herrup, 1995; Billeci et al., 2010).

Species-specific Purkinje cell measurements are presented in Table 3.1. Overall, we determined that the dendritic arbors of gecko Purkinje cells were less complex than those of mice. The total dendrite length of gecko Purkinje cells is less than half of that of mice (means: 1445μm [gecko],

3740μm [mouse]); while the volume of dendrite matter is less than one-quarter (means: 4887μm

[gecko]). 22238μm [mouse]). In addition, geckos had approximately 100 fewer dendritic terminals compared to mice (means: 47.67 [gecko]);146.25 [mouse]). Interestingly, the length of the furthest dendritic terminal (means: 210.6μm [gecko]; 233.75μm [mouse]); and the average length of the dendritic terminals (means: 137.3μm [gecko]; 143.0μm [mouse]); of Purkinje cells of geckos and mice were similar.

3.3.5 Purkinje Cell Neuromorphology With Tail Loss and Regeneration

To determine if tail autotomy (i.e., tail loss), and subsequent tail regeneration resulted in changes to Purkinje cell neuromorphology, we generated three-dimensional representations of

Purkinje cells and quantified using Sholl and branch structure analyses (Figures 3.7,3.8). A representative two-dimensional projection with Sholl circles is shown in figure 3.7A, and data from Sholl and branch structures are shown in Figures 3.7 and 3.8.

The number of dendritic intersections passing through each concentric Sholl sphere was significantly affected by increasing distance from the soma (two-way repeated measures

ANOVA, (F18,162=17.32, p < 0.001), but not by tail loss or tail regeneration (two-way repeated measures ANOVA, F2,9=1.62, p=0.25). There was no interaction between distance and treatment

(two-way repeated measures ANOVA, F36,162=1.24, p=0.1849). The number of dendritic intersections significantly differed between the tail regeneration group and each of the tail loss and control groups at specific distances (=intervals) from the soma (Figure 3.7B). In particular, the tail regeneration group had a significantly higher number of dendritic intersections at 135μm

(Bonferroni post hoc, compared to the tail loss group only, p=0.001), and 150μm (Bonferroni post hoc, compared to the tail loss group, p=0.001, and control group, p=0.03) away from the soma. At these distances the tail regeneration group had more than double the number of intersections compared to the other experimental groups. Purkinje cells from control and tail loss groups also demonstrated an increase in the mean number of intersections up to the distance interval of 90μm from the soma (~10 intersections/shell), after which the number of dendritic intersections decreased. In contrast, neurons from the tail regeneration group increased in mean number of dendritic intersections up to a distance interval of 150μm (~8 intersections/shell) from the soma, after which the number of dendritic intersections decreased. At a distance interval of

200μm from the soma, all three groups had a similar number of dendritic intersections (~2 intersections/shell).

The average diameter of dendrites within each concentric Sholl sphere (Figure 3.7C) was also significantly affected by distance away from the soma (two-way repeated measures

ANOVA, F18,162=53.78, p < 0.001), but not by tail loss or tail regeneration (two-way repeated measures ANOVA, F2,9=0.68, p=0.53). There was no significant interaction between distance and treatment (two-way repeated measures ANOVA, F36,162=1.175, p=1.175). In all groups, dendritic diameter was inversely proportional to increasing distance from the soma. However, the tail regeneration group had significantly greater dendritic diameter than the tail loss group

(Bonferroni post hoc, p=0.03) and the control group (Bonferroni post hoc, p=0.027) at the

distance interval of 165μm from the soma. The tail regeneration group was also significantly greater than the control group (Bonferroni post hoc, p=0.048) at the distance interval of 180μm from the soma. At these distances, the tail regeneration group was more than double the diameter of both the tail loss and tail regeneration groups.

Branch analyses were then used to assess three measures of dendritic complexity: total dendrite volume, farthest dendritic terminal, and total number of dendrite terminals (Fig.3.8A-C).

Unlike the Sholl analyses, which describe neuron shape at set distances from the soma, branch analyses are summation measurements of total neuron characteristics. Total dendrite volume ranged from 4800-8300μm3 depending on the group, but was not significantly affected by tail loss or tail regeneration (one-way ANOVA, F2,9=0.339, p=0.723) (Figure 3.8A). The farthest terminal (longest dendrite from the soma) for all groups was approximately 220μm from the soma, and was also not significantly affected by tail loss or tail regeneration (one-way ANOVA,

F2,9=0.82, p=0.471) (Figure 3.8B). Lastly, the number of terminals (dendrite endings) was approximately 50 for all groups, and not significantly affected by tail loss or tail regeneration

(one-way ANOVA, F2,9=0.05, p=0.95) (Figure 3.8C).

3.4 DISCUSSION This study is the first to characterize and quantify the neuromorphology of lizard Purkinje cells using the Golgi-Cox method. We found that tail loss and tail regeneration have no significant overall effect on Purkinje cell neuromorphology. However, we did determine that there are significant differences in dendritic intersections and diameter, within medial-distal segments of the dendrites once regeneration is complete. These findings are the first to document the consequences of tail autotomy on neuronal structure in the brain, and point towards an

adaptive response possibly underpinning biomechanical changes brought about by a shift in

CoM.

At the level of gross morphology, the gecko cerebellum closely resembles that of other lizards, including varanids (Nieuwenhuys, 1967), agamids (Wylie et al., 2016), and lacertids

(Giacometti et al., 2009) in that it is unfoliated and inverted, curving cranially towards the optic tectum. At the level of histology, the Purkinje cell layer of geckos is 1-3 cells thick, similar to that of agamids (Wylie et al., 2016), but unlike those of other lizards (reportedly 1-5 cells thick;

Nieuwenhuys, 1967). As for other vertebrates (e.g., Wylie et al., 2016; Ariel et al., 2009; Zupanc and Zupanc, 2006), gecko Purkinje cells are immunopositive for the calcium-binding protein calbindin (Zupanc and Zupanc, 2006; Ariel et al., 2009).

3.4.1 A phylogenetic trend is apparent in the dendritic complexity of Purkinje cells

Although Purkinje cells are one of the most distinctive neurons of the brain, our findings confirm that many obvious species-species differences exist. Purkinje cells are classically characterized as having a comparatively large soma (~400μm2), and relatively complex, ramifying dendritic arbors (Llinas and Nicholson, 1971; Voogd and Glickstein, 1998; Anderson and Korbo, 1993; Mavroudis et al., 2010). Like other reptiles, amphibians, and some species of fish, each gecko Purkinje cell has a single primary dendrite that passes deep into the molecular layer, before branching (Nieuwenhuys, 1967; Uray and Gona, 1978; Ariel and Tolbert, 2009). In contrast, among rodents, Purkinje cells may sometimes have two primary dendrites (Nedelscu and Abdelhack, 2018), and subsequent branching into secondary and tertiary branches occurs in more proximal portions of the molecular layer (Anderson and Korbo, 1993; Soha and Herrup,

1995). Although details of neuromorphology remain poorly documented for most species, available evidence reveals a trend towards increasing complexity when comparing reptiles to

mammals (Nieuwenhuys, 1967; Eccles, 1969; Takeda et al., 1992) (Figure 3.9). For example, we found that the dendritic arbors of gecko Purkinje cells had less than half of the total dendrite length, less than a quarter of the total dendritic volume of mice, and ~100 fewer dendritic terminals. A similar trend towards increasing Purkinje cell complexity has been reported elsewhere (Takeda et al., 1992), with mammals and birds revealing the most densely organized dendritic arbors (Jacobs et al., 2012). By way of explanation, Purkinje cell neuromorphology

(and, to a similar extent, the gross morphology of the entire cerebellum) appears to correlate with complexity and sophistication of locomotor mechanics and requirements (Eccles, 1969; Bell,

2002). More specifically, the evolution of precise motor control, and the associated requirement for larger scaffolds for synaptic connections, is predicted to necessitate an increase in Purkinje cell dendritic complexity (Eccles, 1969; Takeda et al., 1992). Similarly, it stands to reason that the recently recognized role of the cerebellum in motor cognition, including tool use, foraging, and the planning and execution of motor planning (Barton, 2012), also impacts Purkinje cell morphology.

3.4.2 Tail loss and regeneration does not significantly affect Purkinje cell neuromorphology

This study is the first to characterize and quantify the neuromorphology of lizard Purkinje cells using the Golgi-Cox method. We have found that tail loss and regeneration following autotomy does not significantly alter the overall neuromorphology of gecko Purkinje cells.

Among mammals, numerous studies have shown that the dendritic arbors of Purkinje cells are highly responsive to a wide variety of extrinsic and intrinsic stimuli, including social interactions, stress, exercise, age, neurodevelopmental disorders, and chemical injuries

(Woodward et al., 197; Floeter and Greenough, 1979; Wong and Wong, 2000; Lein et al., 2007).

For geckos, tail autotomy is not only a traumatic (albeit life-saving) injury, it represents a sudden

and significant change in body length (up to a 40% loss) and body mass (up to 20% loss), and a drastic relocation of CoM (Jagnandan et al., 2014). Whereas autotomized tails do regenerate, the

CoM is not fully restored even once regrowth is completed (Jagnandan et al., 2014). Hence, the absence of obvious and significant changes to gecko Purkinje cell neuromorphology suggests that, at least in this species, these neurons are unexpectedly tolerant of certain forms of biological perturbation. Although this neuromorphological stability may represent evolved cellular adaptations to autotomy, it may also reflect the prioritization of cerebellar control of coordinated multi-limb movements (for escape). Related to this, in mice and humans, it has been shown that synchronous activation of Purkinje cells initiates and coordinates movements of the trunk, limbs, and tail (mice), during locomotion experiments (Bastian et al., 1996; Hoogland et al., 2015).

3.4.3 Purkinje cells show dendritic remodeling at specific intervals following tail regeneration

While our dendrite branch analyses (including Purkinje cell total dendrite volume, farthest dendritic terminal, and total number of dendrite terminals) did not reveal any significant differences between control and experimental (tail loss and tail regeneration) groups, we did find evidence for dendritic remodeling using Sholl analyses (dendritic intersections and diameters).

More specifically, comparing control with tail regeneration geckos we found significant differences in the number of dendritic intersections and dendritic diameters within middle-distal intervals away from the soma (intersections: 135-150μm from soma; diameters:165-180μm from soma). Intriguingly, at these distances from the soma the dendrites become covered in dendritic spines; at more proximal intervals, the dendrites are smooth (i.e., lacking spines) (Nieuwenhuys,

1967; O’Brien and Unwin, 2006). Furthermore, the middle-distal intervals away from the soma also correspond to the position of parallel fiber synapses (Ichikawa et al., 2002; Hoxha et al.,

2016). In contrast, the smooth proximal portions of dendrites, along with the Purkinje cell somas, receive excitatory input from climbing fibers (Crepel et al., 1980; Ichikawa et al., 2002). Parallel fibers – axons from granule cells – synapse on dendritic spines, and are thought to modulate synaptic plasticity underlying motor learning, through long-term depression and/or long-term potentiation (Jorntell and Hansel, 2006; Hoxha et al., 2016). In mice, motor learning has been shown to cause synaptogenesis in the cerebellum, while regular exercise does not (Black et al.,

1990; Lee et al., 2007). Motor learning is a procedural process relating to practice and memory that results in the formation of a new motor skill (Black et al., 1990; Anderson, 1993). Therefore, it is possible that in the changes observed in the gecko dendritic tree at these intervals is related to compensatory motor learning, as the geckos adapt to the permanently changed weight distribution and CoM.

Previous studies have shown a positive correlation between increased spine density and increased dendritic branching (Robinson and Kolb, 1999; Pysh and Weiss, 1979; Adkins et al.,

2002; Ferrante et al., 2013). Furthermore, dendritic morphology of cells within the cerebellum is known to regulate sensory-motor circuit function (Nedelscu and Abdelhack, 2013; Therrien and

Bastien, 2015). In mice, exercise during development and motor skill learning following injury show increased spine density and altered dendritic morphology (Pysh and Weiss, 1979; Bury and

Jones, 2002). It is possible that dendritic plasticity at the medial-distal dendritic interval may also represent an inherent property of terminal branches seen during dendritogenesis and synaptogenesis (Wong and Wong, 2000). For example, during dendritic development, terminal dendritic processes are capable of extensive structural plasticity, whereas primary and secondary branches are relatively more stable (Wong and Wong, 2000; Fujishima et al., 2012).

Additionally, as structural remodeling occurs, dendritic material is added and removed, but there

is no overall change in dendritic length and complexity (Wong and Wong, 2000), similar to this study.

As for other studies, the number of dendritic intersections in gecko Purkinje cells has a unimodal distribution; the number of intersections is greatest mid-distance from the soma, and relatively lower in proximal and distal positions (Billeci et al., 2010). The area with the greatest number of intersections in a dendritic tree is said to be representative of peak dendritic complexity. In mouse CA1 hippocampal pyramidal neurons, the greatest number of intersections is seen ~70μm from the soma (Lein et al., 2007; Elston and Rosa, 2000). CD-1 mouse Purkinje cells in culture have been shown to have a maximum of 14 intersections, at approximately

150μm from the soma (Billeci et al, 2010). In geckos, the control and tail loss groups have their peak number of intersections (~10) at 80-100μm from the soma, while the largest number of intersections from the tail regeneration group (~8) was observed 135-150μm from the soma. This topological shift in dendritic organization likely reflects a change in the number of synaptic inputs Purkinje cells can receive, and thus has downstream implications for neuron signaling and connectivity (Black et al., 1990; Chiu et al., 2008; Hoxha et al., 2016).

Similar to the topology of dendritic intersections, dendritic diameter was also affected by tail regeneration. More specifically, at 165-180μm from the soma, the tail regeneration group had significantly thicker dendritic diameters than other groups (three-dimensional Sholl analysis).

Although our branch analysis showed that total dendritic volume is not significantly different between groups, within the context of a Sholl analysis, diameter is significantly different depending on distance from the soma. Consistent with previous studies (Sdrulla and Linden,

2007; Anwar et al., 2014; Louth et al., 2018), all groups show decreasing diameter with distance, with the proximal primary dendrite being the thickest. Previous studies in mice have shown that

layer VI pyramidal neurons of the medial prefrontal cortex have proximal dendritic diameters as thick as 2.5μm, (Louth et al., 2018), whereas gecko Purkinje cells began with diameters as thick as 3.5-4.0μm thick. This is expected as Purkinje cells are quite large compared to other neurons, especially at the level of the cell soma. The most distal Purkinje cell dendritic branches also have the thinnest diameters, less than 0.5μm wide, which is on par with other mouse studies (Chiu et al., 2008). Dendritic diameter is linked to many neurophysiological effects, including action potential amplitude and intracellular calcium channel changes (Anwar et al., 2014). Calcium dynamics are in turn connected to the regulation of dendritic cone behavior, which prompts dendritic growth or reduction (Zheng et al., 1994; Murchison et al., 2002; Butz et al., 2009).

Increasing dendritic diameter likely decreases axial resistance, which has an effect on the kinetics of input going from dendrites to soma (Mainen et al., 1995). Generally, these dendritic diameter alterations are not known to affect total volume of the neuron (Mainen et al., 1995), which is also consistent with this study.

3.4.4 Purkinje cell morphology does not change shortly after tail loss

We found no significant neuromorphological differences between Purkinje cells from control geckos and the 8 days post-tail loss geckos. In contrast, during development, dendrite remodeling (including the addition or subtraction of dendrite matter) can occur on a timescale of seconds (Wong and Wong, 2000). In culture, a complete Purkinje cell dendritic arbor can be established in 14 days for CD-1 mice (Billeci et al., 2010). By way of explanation, we note that

Purkinje cells must continue to participate in fine motor coordination and regulation in order to survive following autotomy, so that they can competently perform basic survival activities such as roaming and hunting. Since these activities do resume following autotomy, it is possible that immediate compensation may be seen elsewhere in the CNS or PNS other than the cerebellum

(Chapter 2). How the gecko CNS addresses the immediate impacts of tail autotomy remains unknown.

At this time, it remains unclear whether neuromorphological changes observed in gecko

Purkinje cells are permanent or reversible. The experimental timeframe for this study permits regrowth of the tail to occur (i.e., 30 day days; Mclean and Vickaryous, 2011; Delorme et al.,

2012), but did not address whether dendritic plasticity would continue or ablate in the long run.

Previous research using a different species of gecko (Chondrodactylus turnery) revealed that microgravity conditions were sufficient to induce pathomorphological changes in Purkinje cells, but that these alterations were fully reversed when gravity was restored (Proschina et al., 2017).

If, as we predict, the observed changes in dendritic morphology are associated with changes in parallel fiber-Purkinje cell synapses, this remodeling may be indicative of permanent form of motor learning. The parallel fiber-Purkinje cell is known to be capable of bidirectional plasticity, meaning that long-term potentiation or long-term depression can occur with rapid reversibility between the two, meaning that reversal of synaptic and/or dendritic remodeling may be reversible as well (Jorntell and Hansel, 2006; Hoxha et al., 2016).

3.4.5 Future directions

One obvious target for future investigations is to quantitively evaluate spine density and morphology. Similar to dendrites, dendritic spines are dynamic structures that play an important role in nervous system plasticity (Adkins et al., 2002; O’Brien and Unwin, 2006; Lee et al.,

2007). Changes in spine density and morphology in Purkinje cells are associated with motor adaptations (Sdrulla and Linden, 2007; De Bartolo et al., 2015), and has implications for compensatory mechanisms at the level of the synapse, following changes in body mass. Another logical future investigation would involve the use of in vivo 2-photon microscopy, which, along

with genetic labelling techniques, would allow for dendrites to be visualized repetitively in real time (Tavosanis, 2011; Nedelscu and Abdelhack, 2013). This would be helpful in elucidating precise timepoints when dendritic changes are initiated. Overall, tail loss represents an integrative process in the nervous system. As the gecko is a model of rapid mass loss and gradual mass gain, short- and long-term compensation (or motor learning) may be facilitated by neurostructural changes seen at discrete portions in Purkinje cells. The novelty of this study emphasizes that relatively unexplored concepts such as the shift in CoM, limb dynamics, and gait following autotomy may have emerging implications on neuron structure, and by extension, neural signaling and behavior.

CHAPTER 3 TABLES

Table 3.1 Quantification of mouse and gecko Purkinje cell neuromorphological characteristics.

Comparison of control geckos and CD-1 mice used in this study. Means are not statistically analyzed.

Property Gecko Purkinje Cell (Means) Mouse Purkinje Cell (Means) Dendrite Length 1445μm 3740.8μm Dendrite Volume 4887μm3 22238.5μm3 Furthest Terminal 210.6μm 233.8μm Average Terminal Distance 137.3μm 143μm Number of Terminals 47.7 146.3

CHAPTER 3 FIGURES

Figure 3.1. Schematic illustration of experimental design for the Golgi-Cox procedure.

D1 of the Golgi-Cox timeline corresponds to the euthanasia dates on Figure 2.1 (Chapter 2).

During days 1-26, whole brains were incubated in Golgi-Cox solution.

Figure 3.2 Gross anatomy of the gecko brain.

(A) Dorsal and (B) lateral views. The gecko cerebellum is relatively smooth and lacks foliation.

Meninges are present covering on the outer layer of the brain, and olfactory bulbs are absent.

Imaged with Keyence VHX-1000 digital microscope.

Scale bars = 1000μm. bs = brainstem, cb = cerebellum, ot = optic tectum, tel = telencephalon

Figure 3.3. Histology the gecko cerebellum.

(A) Transverse section through the gecko hindbrain, stained with hematoxylin and eosin. The cerebellum sits dorsal to the fourth ventricle, with the granular layer positioned dorsally, which is inverted compared to mammals. (B) Sagittal section through the gecko cerebellum, stained with hematoxylin and eosin. The gecko cerebellum is unfoliated, and resembles a leaf-like sheet.

Scale bars = 100μm. bs = brainstem, gl = granular layer, ml = molecular layer, pl = Purkinje cell layer, v = fourth ventricle

Figure 3.4 Cortical layers of the gecko cerebellum.

(A) Transverse section of the cerebellum stained with hematoxylin and eosin. The highly conserved three-layer arrangement is visible: the granular layer is cell dense, while the molecular layer is cell-sparse. Between the two are the distinctively large cell bodies of the Purkinje cells

(white triangles) within the Purkinje cell layer. (B) Purkinje cells (white triangles) robustly express the calcium-binding protein calbindin (green). Nuclear marker DAPI labeled neuronal and glial nuclei (blue).

Scale bar = 20μm. gl = granular layer, ml = molecular layer, pl = Purkinje cell layer

Figure 3.5. Golgi-Cox stained cerebella of the gecko and mouse.

In both geckos (A) and mice (B), somata of Purkinje cells are clearly visible at the junction between the granular and molecular layers. Dendritic branching is also clearly visible, projecting into the molecular layer towards the pia mater.

Scale bar = 100μm. gl = granular layer, ml = molecular layer, pl = Purkinje cell layer

Figure 3.6. Two-dimensional tracings of gecko and mouse Purkinje cells.

Representative two-dimensional Z-projections of traced Purkinje cells were compiled using

Neurolucida software. (A) Gecko Purkinje cell tracing from control (tail intact) group. Note the presence of a single long primary dendrite and smaller secondary and tertiary branches.

(B) CD-1 mouse Purkinje cell tracing. Mice had comparatively shorter primary dendrites, and longer tertiary and secondary branches.

Scale bar = 20μm

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Figure 3.7. Morphological analysis of gecko Purkinje cell dendrites.

(A) Representative z-projection of traced gecko Purkinje cell, with concentric Sholl spheres radiating from the cell soma at 15μm increments. Scale bar = 25μm. (B) Three-dimensional

Sholl analysis measuring the average number of dendritic intersections with respect to distance from the cell soma, with standard errors (gecko groups: control, n=3; tail loss, n=5; tail regeneration, n=4). Dendritic intersections and diameters were not significantly affected by tail loss or tail regeneration. Post hoc comparisons (Bonferroni method) showed that the tail regeneration group was affected at discrete intervals along the dendritic arbor. More specifically, the tail regeneration group had significantly more dendritic intersections than the tail loss group

(p=0.001) at 135μm from the soma, and 150μm from the soma (p=0.001), and more than the control group (p=0.03) 150μm from the soma (analyzed with repeated measures two-way

ANOVA). (C) Three-dimensional Sholl analysis measuring the average dendritic diameter, with respect to the cell soma, with standard errors (gecko groups: control, n=3; tail loss, n=5; tail regeneration, n=4). Although there were no overall significant differences between groups in terms of dendritic diameters, post hoc comparisons (Bonferroni method) showed that the tail regeneration group had significantly larger dendritic diameters than the tail loss (p=0.03) and control (p=0.027) groups at 165μm from the soma, and significantly larger dendritic diameters than controls (p=0.048) at 180μm from the soma (analyzed with repeated measures two-way

ANOVA). Significance is denoted with an asterisk (*).

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Figure 3.8 Branch analyses of gecko Purkinje cell dendrites.

There were no significant effects of tail loss or regeneration on total dendritic volume (A), terminal size (i.e., farthest dendritic terminal) (B), or number of terminals (C). Means were compared between groups (gecko groups: control, n=3; tail loss, n=5; tail regeneration, n=4), with standard errors, analyzed with one-way ANOVA.

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Figure 3.9. Schematic illustration mapping Purkinje cell neuromorphology onto a phylogeny of vertebrates.

Vertebrate phylogeny showing Purkinje neuron development in evolutionary order. One representative organism is shown from each vertebrate class: mammal – CD1 mouse (present study); bird – pigeon (Nieuwenhuys, 1967); reptile – leopard gecko (present study); amphibian – frog (Nieuwenhuys, 1967); lamprey (Nieuwenhuys, 1967). Varying degrees of dendritic complexity is visible, with a phylogenetic trend.

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CHAPTER 4: CONCLUDING STATEMENTS

This study sought to investigate whether elements of the central nervous system (CNS) and peripheral nervous system (PNS) were altered in response to tail autotomy and tail regeneration in the leopard gecko (Eublepharis macularius). Tail autotomy involves a near-instantaneous loss of mass at the distal end of the body (Jacyniak et al., 2018). As a result, there is a cranial shift in the CoM, changes in ground reaction force, as well as kinematic and muscular changes (Jagnandan et al., 2014, Jagnandan and Higham, 2018). Here, we conducted a spatiotemporal examination of tactile sensitivity using monofilaments, and quantified the neuromorphology of Purkinje cells, using Golgi-Cox staining, to assess the nervous system following tail loss and growth. We determined that there are transient changes in tactile sensitivity immediately following tail loss, but that there are no overall changes in dendritic morphology. However, we did find that there were localized changes in dendritic arbor morphology of Purkinje cells following tail regrowth. Taken together, our findings demonstrate the first evidence for compensatory roles of both the CNS and PNS, following tail loss and regeneration in geckos.

4.1 Is post-autotomy compensation occurring in the PNS of the leopard gecko? Using Semmes-Weinstein monofilaments, we identified regional differences in tactile sensitivity across the ventral surfaces of the hindlimbs, forelimbs, and tail of the geckos.

Repeated monofilament testing, even in the absence of tail autotomy, had a significant sensitization effect (i.e., decrease in threshold) compared to baseline, at all locations. This sensitization may have occurred as a result of stress or novelty, of the experimental chamber/protocol, and/or may reflect a defensive response. In future studies, it is recommended that geckos are given time prior to the experimental timeline (they were only given acclimation

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time during the experiment), to familiarize themselves to the chamber. Our application of monofilaments (from beneath the gecko) arguably mimics the predatory strategy of the subterranean snake Eryx johnii. E. johnii co-exists in the wild with geckos, its presence is known to elicit a strong antipredatory behavioral response, even from captive-bred individuals (Landova et al., 2016). Therefore it is possible that the testing procedure itself confused the geckos into misinterpreting the presence of a predator.

Immediately following autotomy, we found that most sites of interest paralleled the sensitization response observed in the control (tail-intact) group with two notable exceptions.

Monofilament testing of the forelimbs revealed no significant effect of sensitization (from baseline, day 0) until the tail was fully regenerated (day 31). In contrast, monofilament testing of the hindlimbs revealed they were even more sensitized than the control group, immediately following tail loss. This likely confers biomechanical advantage following a rapid change in mass, since altered tactile sensitivity has implications on balance and locomotion (Eils et al.,

2002). Interestingly, the tactile sensitivity of regenerated tail tips was not significantly different from original (intact) control group tails. This indicates that tail regeneration effectively restores somatosensory function to the new appendage. Unexpectedly, we found that the tactile sensitivity of the tail base was highly variable, even between the two tail autotomy groups.

Although the cause of this variation remains unknown, one possibility is that our method of inducing tail autotomy (pinching the base of the tail) generates inconsistent damage to the remaining PNS and CNS of the tail stump. In the future, we recommend performing surgical amputations of the tail, so that the spinal cord and peripheral nerves are severed in a consistent manner, thereby minimizing variation in nervous system wounding between geckos.

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Our findings suggest that tactile sensitivity across the ventral body surface is not simply correlated to sensilla density. Whereas previous work on geckos found that the tail tip had a higher density of sensilla than the tail base in original tails (Russell et al., 2014), we determined the tail base was the more sensitive of the two locations. Tactile sensitivity is a product of the density, distribution, and acuity of mechanoreceptors (Purves et al., 2001; Macefield, 2005).

Tactile sensitivity can also be modulated through plastic changes in central processing

(central-reorganization) by the CNS (Makin et al., 2013; Strzalkowski et al., 2015). In addition, mechanoreceptive afferent physiology may also be affected by skin capillary permeability and vasoconstriction, both of which may be altered in response to tail loss. We suggest that the immediate changes in tactile sensitivity observed following tail loss are indicative of a short-term compensatory mechanism, and most likely participate in ensuring locomotor stability following a rapid (and drastic) change in body mass.

4.2 Is post-autotomy compensation occurring in the CNS of the leopard gecko?

Using Golgi-Cox staining, we characterized and quantified the neuromorphology of

Purkinje cells within the gecko cerebellum. Similar to other reptiles, amphibians, and some fishes, gecko Purkinje cells have a large soma, a lengthy primary dendrite passing into the molecular layer, and expansive branching into secondary and tertiary branches. Overall, we found there were no significant changes in Purkinje cell neuromorphology following tail loss and tail regeneration. However, our Sholl analyses revealed significant increases in dendritic diameter, and the number of dendritic intersections, in middle-distal segments of the dendritic arbor 30 days following tail loss. Changes in neuromorphology have important implications on function and connectivity; dendrite structure is implicated in many neurophysiological parameters: action potential amplitudes, axial resistance, and intracellular calcium dynamics, to

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name a few (Zheng et al., 1994; Mainen et al., 1995; Vetter et al., 2001). The middle-distal segment of the dendritic arbor corresponds to the area where parallel fibers synapse on the

Purkinje cell (Ichikawa et al., 2002; Hoxha et al., 2016). Parallel-fiber Purkinje cell synapses have been implicated in motor learning, through long-term potentiation and/or long-term depression (i.e., long-term changes in synaptic efficacy and modifications in synaptic connections) (Jorntell and Hansel, 2006; Sdrulla and Linden, 2007). By way of explanation, we propose that the permanent shift in CoM following tail regeneration may necessitate permanent changes in motor control, and/or motor learning. Our findings also suggest that the need to maintain cerebellar function following tail loss (imperative to survival) may act to attenuate any immediate changes in dendritic morphology resulting from the sudden alteration in body mass and weight distribution. Indeed, running speed of the gecko is unaffected by tail loss (Jagnandan et al., 2014). We propose that the morphological changes observed by the end of regeneration reflect a long-term compensatory mechanism in response to the permanent alteration in biomechanics. A summary of these biomechanical shifts is presented in Table 4.1.

4.3 Integration of the CNS and PNS in the leopard gecko

Taken as a whole, our findings reveal that the effects of tail autotomy are minimized in the forelimbs, but actively accommodated via biomechanical changes at the hindlimbs. This includes changes in tactile sensitivity (our present study), joint angles (resulting in altered posture; Jagnandan et al., 2014), and muscle activation in the hindlimbs as seen in previous work

(Jagnandan and Higham, 2018). The observed disparity in tactile sensitivity may reflect the difference in functional roles of the forelimbs and hindlimbs. Whereas forelimbs are involved in braking, energy absorption and stability, hindlimbs are primarily involved in propulsion (Lee,

2010; Foster and Higham, 2012; Autumn et al., 2016). How these are changed in response to tail

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autotomy is only now being explored. In one recent study, the effect on tail autotomy on locomotion was investigated using a lizard species capable of quadrupedal and bipedal

(hindlimb) running, Acanthodactylus schreiberi. Whereas tail autotomy had no effect on quadrupedal running, it significantly impacted bipedal locomotion, effectively abolishing this behavior in adults (Savvides et al., 2017). Our findings with tactile sensitivity highlight how locomotion and gait parameters can be maintained following autotomy, in spite of substantially altered body size and mass. In support of our findings, tactile afferents in the human hands and feet are known to modulate gait (Meyer et al., 2004; McPoil and Cornwall, 2006), while tactile afferents in human fingers have been shown to facilitate adaptive motor responses (Johansson and Westling, 1987).

Purkinje cells are the sole output of the cerebellar cortex and project to upper motor neurons (through deep cerebellar nuclei via the thalamus), to coordinate movement (Konnerth et al., 1990; Takakusaki, 2017). Our observation that the dendritic arbors of Purkinje cells are altered in response to tail regeneration (specifically in the area corresponding to the parallel fiber synapses) suggest that neuroplasticity may play a role in long-term compensation following regrowth of the tail. In support of this interpretation, Jagnandan et al. (2014) found that the CoM of geckos having undergone tail autotomy and regeneration never returns to baseline. Whether there is an associated change in cortical-reorganization (or cortical plasticity) remains unclear; however in cases of amputated appendages, cortical-reorganization is known to occur (Flor et al.,

2008; Makin et al., 2013). Reptiles do not have a motor cortex or somatosensory cortex homologous to that of mammals (Naumann et al., 2015), although there are functionally comparable regions, such as the lateral portion of the dorsal cortex (Medina and Reiner, 2000). A neuromorphological analysis of neurons within the dorsal cortex would be an interesting target

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for future studies, as there are multiple locations in the CNS implicated in movement (in addition to the cerebellum).

An important avenue for future studies would be to investigate the link between the somatosensory system and the cerebellum. Mammalian studies have revealed an extensive network of cutaneous somatosensory projections to the granule cell layer (via mossy fibers) of the cerebellar hemispheres and the caudal vermis (Welker and Shambes, 1985; Bengtsson and

Jorntell, 2008). This provides a link between the somatosensory input and Purkinje cells

(mediated in part by granule cells, which synapse on Purkinje cells via parallel fibers). At this time, the relationship between tactile sensitivity and changes in neuromorphology of Purkinje cells remains unclear. Further, Purkinje cells were only assessed at two timepoints in this experiment (8 and 30 days post-autotomy). As these cells are notoriously dynamic (with neuromorphological changes potentially happening on a timescale of seconds; Wong and Wong,

2000) it would be worthwhile investigating Purkinje cell dendritic morphology as early as one day following autotomy (similar to monofilament testing).

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Table 4.1. Properties of tail loss and tail regeneration in the leopard gecko.

Biomechanical and nervous system changes following tail loss and regeneration. Parameters undergoing significant or obvious changes are bolded.

Property Tail Loss Tail Regeneration Study

Length Lost 40% 70% restored Present Study Mass Lost 20% resolved Present Study Tactile Sensitivity - Forelimbs unchanged unchanged Present Study Tactile Sensitivity - Hindlimbs decreased resolved Present Study Tactile Sensitivity - Tail Base variable unchanged Present Study Tactile Sensitivity - Tail Tip unchanged unchanged Present Study Neuromorphology Purkinje Cells unchanged overall unchanged - discrete portions Present Study Center of Mass (CoM) Shift 13% anterior shift 6% anterior shift Jagnandan et al., 2014 Running Speed unchanged unchanged Jagnandan et al., 2014 Peak Vertical GRF reduced resolved Jagnandan et al., 2014 Peak Propulsive GRF increased resolved Jagnandan et al., 2014 Posture - Forelimbs unchanged unchanged Jagnandan et al., 2014 Posture - Hindlimbs sprawled resolved Jagnandan et al., 2014 Joint Angles - Forelimbs unchanged unchanged Jagnandan et al., 2014 Joint Angles - Hindlimbs femur depression, knee angle reduction resolved Jagnandan et al., 2014 Muscle Activity - Forelimbs unchanged unchanged Jagnandan and Higham, 2018 Muscle Activity - Hindlimbs propulsive muscles reduced resolved Jagnandan and Higham, 2018

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APPENDICES (I-V) APPENDIX I – EXPERIMENTAL GROUPS

Start of experiments: Nov 3/2017 Monofilament testing occurred between 8:30am-5:30pm each day. Schedule: 1 hour acclimate/30-45 minutes monofilament testing. 5 geckos/per day *Feeding followed monofilament testing at the end of the day. Cage changes always a day after the monofilament testing for the group. Tail loss group experimental dates: Nov 4 – Nov 13 Control group experimental dates: Nov 12 – Dec 13 Tail regeneration group experimental dates: Nov 3 – Dec 4 Euthanasia timepoints: all 5 geckos collected, Golgi-Cox timeline began Monofilament Timeline: D0 – baseline monofilament testing D1 – autotomy D2 – monofilament testing D7 - monofilament testing D14 - monofilament testing D21 - monofilament testing D31 - monofilament testing D32 – euthanasia. This served as D1 for the Golgi-Cox timeline. Golgi-Cox Timeline: D1 – Whole brains into Golgi-Cox solution D26 – Whole brains into sucrose solution D29 - Brain sectioning and embedding D30 - Brain histology and mounting

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Table A1: Experimental group data at start of experiment

Gecko Experimental Starting Mass Starting S-V Tail Length Euthanasia Group (g) length, (mm) (mm) Timepoint #1 - May-01-01 Control 20.5 100 80 31 days #2 - May-01-06 Control 21.2 90 80 31 days #3 - May-01-08 Control 19.3 90 80 31 days #4 - May-01-25 Control 20.5 100 80 31 days #5 - May-01-29 Control 22.4 100 80 31 days #6 - May-01-19 Tail Loss 21.7 100 80 8 days #7 - May-01-27 Tail Loss 22.4 100 80 8 days #8 - May-01-17 Tail Loss 22.3 100 80 8 days #9 - May-01-03 Tail Loss 20.8 100 70 8 days #10 - May-01-04 Tail Loss 22.1 90 80 8 days #11 - May-01-02 Tail Regeneration 19 100 80 31 days #12 - May-01-09 Tail Regeneration 17.5 100 80 31 days #13 - May-01-10 Tail Regeneration 17.1 100 90 31 days #14 - May-01-11 Tail Regeneration 18.5 100 80 31 days #15 - May-01-31 Tail Regeneration 17.9 100 90 31 days Averages 20.21333333 98 80.66666667

Table A2: Experimental group data at euthanasia

Gecko Experimental Mass at S-V Length at Tail Length at Euthanasia Group Euthanasia(g) Euthanasia, (mm) Euthanasia(mm) Timepoint #1 - May-01-01 Control 26.5 100 80 31 days #2 - May-01-06 Control 26.6 95 80 31 days #3 - May-01-08 Control 26 110 80 31 days #4 - May-01-25 Control 27.7 100 80 31 days #5 - May-01-29 Control 29.4 110 90 31 days #6 - May-01-19 Tail Loss 20 100 12* 8 days #7 - May-01-27 Tail Loss 19.6 100 15* 8 days #8 - May-01-17 Tail Loss 19.5 95 11* 8 days #9 - May-01-03 Tail Loss 19 100 11* 8 days #10 - May-01-04 Tail Loss 20.2 95 11* 8 days #11 - May-01-02 Tail Regeneration 23.2 110 20* 31 days #12 - May-01-09 Tail Regeneration 20.6 110 10* 31 days #13 - May-01-10 Tail Regeneration 20.7 100 20* 31 days #14 - May-01-11 Tail Regeneration 22.7 100 20* 31 days #15 - May-01-31 Tail Regeneration 22.2 110 30* 31 days Averages * = regenerate Control 27.24 103 82 Tail Loss 19.66 98 12 Tail Regen 21.88 106 20

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Table A3: Autotomy data, tail loss group

Gecko Total Mass Tail Mass % Tail - Total Tail Autotomized Body Length % Autotomy- (g) (g) Total Mass Length (mm) Tail Length (mm) (S-V, mm) Body Length #11: May01-19 21.7 3.8 17.50% 75 68 95 40% #12: May01-27 22.6 4.5 19.90% 78 68 100 38.20% #13: May01-17 22.7 4.6 20.30% 75 74 100 42.30% #14: May01-03 22.3 4.7 21.10% 90 72 100 37.90% #15: May01-04 24.2 4.6 19% 75 65 96 38% AVERAGE 22.7 4.44 19.56% 78.6 69.4 98.2 39% Table A4: Autotomy data, tail regeneration group

Gecko Total Mass Tail Mass % Tail - Total Tail Autotomized Body Length % Autotomy- (g) (g) Total Mass Length (mm) Tail Length (mm) (S-V, mm) Body Length #6: May01-02 22.8 4.1 18% 80 72 105 38.90% #7: May01-09 20.7 3.9 18.80% 80 70 90 41.20% #8: May01-10 20.7 3.8 18.40% 80 76 107 40.60% #9: May01-11 22.8 4.7 20.60% 77 71 95 41.30% #10: May01-31 21.1 4 19% 80 71 100 39.40% AVERAGE 21.62 4.1 19% 79.4 72 99.4 40.28%

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Appendix II: Detailed Histochemical and Immunohistochemical Protocols and Recipes

Golgi-Cox Staining Protocol (from Bailey lab at University of Guelph)

1. Subject is euthanized and the brain dissected out.

2. Whole brain is placed into a 20 mL scintillation vial containing 20 mL of Golgi-Cox Solution (i.e., topped-up with solution until the container is full). Cover the vial with aluminum foil and incubate for 29 days at room temperature in the dark.

3. Place brain into a 20 mL scintillation vial containing 20 mL of Sucrose Cryoprotectant. Cover the vial with aluminum foil and incubate for 48 hours at 4 °C in the dark.

4. Remove brain from sucrose and block it for slicing by cutting the cerebellum off and leaving a flat edge at the back (caudal) end of the brain. Place the brain standing upright in a small weigh boat/dish. Heat the agar prior to blocking the brain and when it has cooled to near-solid phase, pour it into the weigh boat to encase the brain.

5. Once the agar has solidified, trim excess agar and glue the brain to the stage of the vibratome. Slice in Sucrose Cryoprotectant at 200 to 500 μm thickness, depending on the brain region to be examined. Place slices into a well of a 6-well plate (35 mm diameter) filled with 10 mL of 6% Sucrose and incubate overnight at 4 °C in the dark.

6. Place slices into a well with 5 mL of 2% paraformaldehyde. Moderate speed rocking (20rpm) for 15 min.

7. Wash slices twice in wells containing 5 mL of H2O. Slow speed rocking (15rpm) for 5 mins each.

8. Place slices into a well filled with 5 mL of 2.7% NH4OH. Moderate speed rocking (20rpm) for 15 min.

9. Wash slices twice in wells containing 5 mL of H2O. Slow speed rocking (15rpm) for 5 mins each.

10. Place slices into a well filled with 5 mL of Kodak Fixative A (diluted 10x from purchased concentration). Moderate speed rocking (20rpm) for 25 minutes.

11. Wash slices twice in wells containing 5 mL of H2O. Slow speed rocking (15rpm) for 5 mins each.

12. Mount slices onto gelatinized slides, or Superfrost Plus slides. Remove excess agar and water with a kimwipe.

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13. Allow slides to air dry for 45 mins (400 μm) or 90 mins (500 μm). This timing is critical, as too short a time leads to slices falling off of the slides and too long leads to cracked slices. Should look shiny, with a bit of a halo where water has dried.

14. Dehydrate slices: a. 50% EtOH 2 min b. 75% EtOH 2 min c. 95% EtOH 2 min d. 100% EtOH 5 min e. 100% EtOH 5 min f. Citrisolv for 5 min g. Citrisolv for 5 min.

15. Coverslip each section or slide using Permount and allow slides to dry horizontally for five days in the dark.

Golgi-Cox Solution 1% (w/v) potassium dichromate 0.8% (w/v) potassium chromate 1% (w/v) mercuric chloride

0.4 M Phosphate Buffer Stock 0.9% (w/v) sodium phosphate monobasic (anhydrous) 8.7% (w/v) sodium phosphate dibasic (heptahydrate)

Sucrose Cryoprotectant 30% (w/v) sucrose in 0.1 M phosphate buffer

2% Paraformaldehyde 2% (w/v) paraformaldehyde in 0.1 M phosphate buffer

3% Agar 3% (w/v) agar in water

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Hematoxylin and Eosin Protocol

1. Absolute xylene (3 washes; 2 minutes each) 2. Absolute isopropanol (3 washes; 2 minutes each) 3. 70% isopropanol (2 minutes) 4. dH2O (2 minutes) 5. Modified Harris Hematoxylin (10 minutes) 6. Rinse in running dH2O to remove excess Hematoxylin 7. 1% acid alcohol (4 dips) 8. Rinse in dH2O 9. Ammonia water (until blue, ~6 dips) 10. Rinse in dH2O 11. 70% isopropanol (6 dips) 12. Eosin (1 minute) 13. Absolute isopropanol (3 washes; 2 minutes each) 14. Absolute xylene (3 washes; 2 minutes each) 15. Coverslip

Acid Alcohol 1% HCl in 70% isopropanol

Ammonia Water 5 drops ammonium hydroxide 250mL dH2O

Eosin Stock Solution 10g Eosin Y 1g Phloxine B dissolve in 1000mL 80% ethanol

Eosin Working Solution 200mL Eosin stock solution 200mL dH2O 600mL absolute ethanol 5mL glacial acetic acid

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Immunofluorescence Protocol

1. Absolute xylene (3 washes; 2 minutes each) 2. Absolute isopropanol (3 washes; 2 minutes each) 3. 70% isopropanol (2 minutes) 4. dH2O (2 minutes) 5. Citrate buffer retrieval (12 minutes at 95oC, 20 minutes at room temperature) 6. 1XPBS (3 washes; 2 minutes each) OR 1XPBST (3 washes; 2 minutes each) 7. 5% normal goat serum (NGS) block diluted in 1XPBS (1 hour at room temperature) 8. Tip off blocking solution 9. Incubate in primary antibody diluted in 1XPBS or 1XPBST (overnight at 4oC) 10. 1XPBS (3 washes; 2 minutes each) 11. Incubate in secondary antibody diluted in 1XPBS (1 hour at room temperature) 12. 1XPBS (3 washes; 2 minutes each) 13. Counterstain with DAPI (1:5,000; 2 minutes) 14. 1XPBS (3 washes; 2 minutes each) 15. Coverslip with fluorescent mounting media

Citrate Buffer Stock Solution A (0.1M Citric Acid) 1.92g citric acid powder 100mL dH2O

Citrate Buffer Stock Solution B (0.1M Sodium Citrate dehydrate) 14.7g sodium citrate dehydrate powder 500mL dH2O adjust pH to 6.0

Working Citrate Buffer 1mL Solution A 5mL Solution B 44mL dH2O

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APPENDIX III: RAW MONOFILAMENT DATA Table A5: Raw monofilament data for control group

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Table A6: Raw monofilament data for tail regeneration group

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Table A7: Raw monofilament data for tail loss group

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APPENDIX IV: 2-DIMENSIONAL PURKINJE CELL TRACINGS Table A8: Number of neurons (‘n’) averaged for each gecko and mouse

Control Geckos, n=3 Tail Regeneration Geckos, n=4 Tail Loss Geckos, n=5 Mice, n=2 Animal # G#1 G#2 G#3 G#4 G#5 G #6 G #7 G #8 G #9 G #10 G #11 G #12 G #13 G #14 G #15 M#1 M#2 # of Neurons 3 - 3 - 2 2 1 - 3 2 3 1 2 6 1 2 2

Figure A1: Two-dimensional Purkinje cell tracings. For control, tail loss, and tail regeneration groups. Scale bar = 20μm

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APPENDIX V: STATISTICAL OUPUT Table A9: Three-Way ANOVA output for all monofilament data (normalized to baselines)

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Figure A10: Two-Way ANOVA output for Sholl Analyses of Purkinje cell intersections and diameter

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