Delayed Developmental Loss of Regeneration in Xenopus

laevis tadpoles

A thesis submitted to the Graduate School

of the University of Cincinnati

In partial fulfillment of the requirements for the degree of Master of Science

In the department of Biological Sciences

of the McMicken College of Arts and Sciences

by

Justin Y. He

B.S. Biology, University of the Pacific

Committee:

Dr. Daniel Buchholz- Chair

Dr. Ed Griff

Dr. Josh Benoit

March 2021

i Abstract: The prospect of spinal cord regeneration in humans is an exciting medical advance, but one that remains elusive from the complicated cellular and molecular mechanisms that prevent regeneration from happening. Various model organisms that do possess regenerative ability have been studied in hopes of understanding how spinal cord regeneration can be facilitated in humans. Recent studies in non-regenerative mammalian organisms however have uncovered the role of T3 signaling pathways in inhibiting regenerative capacity. These previous studies have shown inhibition of T3 in-vitro and in-vivo in various model organisms has increased the capacity for regeneration even in organisms that typically do not have such an ability. My dissertation provides a broad examination of previous literature exploring the barriers to regeneration in a wide range of model organisms, as well as potential therapeutic targets for inducing regeneration. Here, I also show how inhibition of T3 in X. laevis tadpoles allows for increased functional recovery from spinal cord transection.

ii

© Copyright by Justin He 2021 All Rights Reserved

iii Acknowledgements

As I conclude my studies at UC in the midst of the COVID-19 pandemic, thank you to all of my friends, colleagues, and family for their love and support in these hectic times. Thank you to my advisor Dr. Daniel Buchholz for his support in my research and writing, and to my lab members for constructive feedback on my presentations during our lab meetings. As I transition into a career as a K-12 teacher, I would also like to thank Dr. Brent Stoffer, the head of the BIOL

1081L & 1082L lab series, for showing us how to effectively teach the material during my time as a TA and for his guidance on transitioning to the world of virtual teaching during this pandemic. Special thanks to him for also providing me with the pedagogical skills necessary to be an effective science educator.

iv Table of Contents Chapter 1

Introduction……………………………………..………………………………...…...…………..2

Evolutionary and Developmental Loss of Regeneration………………………..………………...3

Extrinsic Factors Inhibiting Regeneration: Inflammatory Responses and Myelin…………………………………...……..………………….5

Intrinsic Factors: Neural Signaling Pathways and Crucial Neural Genes………………………...…...……………..8

Summary and Next Steps………………………………...………………………………………11

References……………………………...………………...………………………………………12

Chapter 2

Abstract………………………………...………………...………………………………………17

Introduction…………...…………………………………………………….……………………17

Materials and Methods…………………………………………..………….……………………21

Results…………………………………………..………………….……….……………………25

Discussion………………..………………….……………………………...……………………28

Tables and Figures……………………………...………………………………………………..35

Chapter 3

Where do we target? …………………………………………………………………...………..41

Recent advances in our understanding of neural regeneration……………………...…………………………………………...………..43

Implications of facilitating regeneration?...... ……………………………………44

Concluding Remarks ...... ………...... ………………………………………….45

References Cited…...... ………...... ………………………………………… 47

v

Chapter 1

1 Introduction

The capacity for neural regeneration specifically recovery from spinal cord injury varies across the animal kingdom (Alibardi, 2019; Lee-Liu et al., 2013). Basal vertebrates of the animal kingdom such as anamniotes including fish and amphibians possess the ability to recover from neural injury post embryonically unlike amniotes including mammals which only possess regenerative ability in the embryonic stages (Kundi, 2013). Understanding the mechanisms of regenerative abilities and developmental loss of regeneration across the vertebrate groups can answer questions on how neural injuries including spinal cord injury can be cured in non- regenerative organisms. Many studies in various model organisms have been conducted to understand cellular and molecular barriers to regeneration. These studies have suggested potential therapeutic targets for facilitating regeneration, detected cellular functions impeding regeneration, as well as identified key regenerative genes that have been lost throughout the evolutionary history of vertebrates.

Peripheral nervous system neurons in mammals (PNS) unlike central nervous system neurons have been found to be capable of recovering from injury (Filbin, 2003; Mietto et al.,

2015). Both extrinsic and intrinsic factors have been studied in understanding differential ability to regenerate between CNS and PNS neurons. Extrinsic factors include external neural factors such as glial scar formation or myelin and intrinsic factors include internal neural factors such as cytoskeletal organization after injury or expression of certain transcription factors. It has been previously concluded that the different external environments of CNS and PNS neurons are the main factor in differential regeneration ability (David, S; Aguayo, 1981; Reier et al., 2017). This is based off on previous observations showing how PNS neurons transplanted into a CNS

2 environment are unable to elongate despite their know regenerative capabilities (Aguayo et al.,

1981). It has also been shown that CNS myelin is inhibitory to regeneration but not PNS myelin

(Yiu & He, 2006). The extrinsic environment of CNS neurons is influenced by certain inhibitory molecules that are not expressed around the PNS (Mietto et al., 2015; Toy & Namgung, 2013).

These extrinsic factors of the CNS can be targeted to create a more permissive environment for spinal cord injury recovery (Ferguson & Son, 2011; Kim et al., 2012). Here I discuss key studies across various model organisms that show evolutionary and developmental loss of regeneration, extrinsic and intrinsic factors inhibiting regeneration, as well as potential therapeutic targets to facilitate regeneration.

Evolutionary and Developmental Loss of Regeneration

In many vertebrates, some degree of regenerative ability in the CNS can be found at the earlier stages of development especially in the embryonic stages as it is the stage where there is much potential for neural stem cells to differentiate into their functional form (Alibardi, 2019). In vertebrates that undergo metamorphosis, however, any traces of regenerative ability in CNS neurons cease to exist as development progresses. Why and how metamorphic changes are associated with loss of regeneration is continuing to be studied but various theories have been formulated discussing how developmental loss of regeneration occurs and why mammals and birds experience this phenomenon. From an evolutionary standpoint, it is believed that the expression capabilities of regenerative genes were lost from the transition from water to land due to the need for more specialized organs both internally and externally to perform complex functions on land (Alibardi, 2019). Various inhibitory mechanisms have evolved to suppress

3 neural plasticity as memory and behavioral functions would be lost from constant remodeling of the nervous system.

At what stage of life developmental loss of regeneration occurs varies across vertebrates.

In humans, regenerative ability is lost at the onset of birth. Other mammals such as opossums possess the ability to recover from spinal cord injury within only 9-12 days postnatal (Mladinic et al., 2009; Wheaton et al., 2020). In certain amphibians such as Xenopus laevis frogs on the other hand, the loss of regeneration occurs during the transition from tadpole to froglet (Gaete et al., 2012; Helbing et al., 2003). The developmental transition from tadpole to froglet is analogous to the transition from fetus to birth in humans as well as developmental loss of regeneration in humans making X. laevis a valuable model organism to study developmental loss of regeneration (Buchholz, 2015). The difference in regenerative capabilities between X. laevis tadpoles and froglets is also evident from other studies that have shown the differences in regenerative ability between tadpoles and froglets. Through histological studies, a negative correlation between neural regrowth across the lesion and developmental stage of X. laevis tadpoles was shown (Muñoz et al., 2015). Lower developmental staged tadpoles experienced a greater degree of neural regrowth across the lesion site than the more mature tadpoles. At the end of 20 dpt (days post transection) lower staged tadpoles had formed an ependymal canal, a central canal of the spinal cord, across the lesion site (Muñoz et al., 2015). While some cellular material did cross the lesion site of higher staged tadpoles at 20 dpt, an ependymal canal was not formed indicating incomplete spinal cord regeneration (Muñoz et al., 2015). Understanding the process of developmental loss of regeneration in these model organisms and what signaling pathways are involved is of great interest in understanding the barriers to regeneration.

4 Metamorphosis is induced by thyroid hormone (T3) suggesting a role of T3 in inhibiting regeneration. T3 is also required for brain remodeling and developmental transition during birth in mammals including humans. Extensive studies on T3 function in other model organisms have already shown the inhibitory effects of T3 on regenerative capacity. Studies on organotypic brain slice culture from mice have shown that blocking T3 signaling allows for axon growth across the lesion site (Avci et al., 2012). Rearing zebrafish in T3 inhibitory chemicals accelerated optic nerve repair after optic nerve crush injury (Bhumika et al., 2015). Thyroxin, a precursor form of

T3, inhibits tail regeneration of Rana catesbeiana tadpoles and induces resorption of regenerative tissue at the peak of metamorphosis (Li & Bern, 1976). Studies on X. laevis have also shown the implications of delayed and accelerated metamorphosis on regenerative capacity.

Development was inhibited in X. laevis by treating tadpoles with the T3 inhibiting drug methimazole, while premature metamorphosis was induced through T3 treatment (Gibbs et al.,

2011). Methimazole-treated tadpoles that have undergone spinal cord transection also experienced axon regrowth through the lesion site (Gibbs et al., 2011).

Extrinsic Factors Inhibiting Regeneration: Inflammatory Responses and Myelin

Inflammatory reactions that occur after neural injury have been extensively studied as it is hypothesized that the immune system has evolved to prevent neural plasticity of neurons

(Alibardi, 2017). An inflammatory reaction occurs through reactions of astrocytes and recruitment of immune cells to the injury site forming a glial scar, a physical barrier to axon growth (Lee-Liu et al., 2013; Michael V. Sofroniew, 2010). After neural injury, a blastema, which is characteristic of development, forms along the injury site (Alibardi, 2019). In pre- metamorphic Rania dalmatina tadpoles however, the blastema from amputated has been found to

5 be infiltrated with immune cells suggesting an impediment to development from scar tissue that has formed (Alibardi, 2017). In organisms whose immune systems continue to be more active as development progresses, lack of inflammatory responses after neural tissue damage was shown at the early stages of development but is continuously enhanced as development progresses

(Eming et al., 2009). The dramatic changes in immune response to neural injury is reflected in the development of the immune system in X. laevis. While X. laevis do have similar immune systems to mammals, the loss of regenerative capacity is correlated with progressive maturation of the immune system (Robert & Ohta, 2009; Simon et al., 2015). It has been found that dendritic cells MHC class II molecules begin to appear in X. laevis skin during the midlarval stages but become more abundant at the pro-metamorphosis stages (Mescher et al., 2007). This has led to some speculation that continuing development of the immune system is the cause for the gradual loss of regenerative ability in X. laevis. (Eming et al., 2009; Lee-Liu et al., 2013;

Murray Blackmore Paul C. Letourneau, 2006).

Despite the likely hypothesis that development of the immune system is associated with changes in regenerative capacity of tadpoles, other organisms that maintain regenerative ability throughout life such as planaria or zebrafish still have immune systems. Interestingly, such organisms do not exhibit glial scar formation after neural injury even with fully developed immune systems (Cigliola et al., 2020; Eming et al., 2009; Bloom, 2014). In zebrafish, however, it has been found that the immune response that is initiated after injury differs from the immune response of other organisms with lack of regenerative ability. The immune response of zebrafish after spinal cord injury facilitates regeneration unlike the immune systems of non-regenerative organisms which impedes regeneration. After spinal cord injury in zebrafish, glial cells form a bridge across the lesion site which serves as a scaffold for axons to regrow on. The formation of

6 glial bridges is induced by the expression of ctgfa around glial cells (Mokalled et al., 2016).

Overexpression of ctgfa accelerated the regeneration process, while ctgfa knockout zebrafish showed defects in glial bridge formation. As of now, ctfga has not been found in other model organisms so it is unknown if lack of ctgfa expression in mammals would explain the formation of glial scars. Aside from the expression of ctfga however, the formation of glial bridges in zebrafish is also dependent on a family of fibroblast growth factor (fgf) genes (Goldshmit et al.,

2018). fgf3 is a particular fibroblast growth factor gene that induces neurogenesis certain subset of motor neurons (Goldshmit et al., 2018). Analysis of longitudinal sections of fgf3 injected zebrafish spinal cord injury has shown enhanced recovery 10 days after injury (Goldshmit et al.,

2018). The fgf3 gene however has not been found to be expressed in rat cells explaining the possibility of lack of regenerative capacity without expression of the gene (Goldshmit et al.,

2018). Studies are continuously being conducted in understanding how the immune systems of regenerative organisms do not inhibit regeneration, but the differences in immune signaling pathways between zebrafish and rats provide clues into how regeneration is facilitated in organisms that maintain regenerative abilities throughout life.

Myelination from oligodendrocytes is another extrinsic factor of interest inhibiting regeneration (Ferguson & Son, 2011). Certain oligodendrocyte receptors have been identified as inhibitory receptors for regeneration. While both PNS and CNS neurons are myelinated, it has been found that certain receptors found CNS oligodendrocytes but are not expressed in PNS oligodendrocytes. The differential expression of certain receptors between PNS and CNS neurons has allowed for the identification of crucial receptors involved in the inhibition of axon regrowth after injury. It is hypothesized that oligodendrocyte receptors serve as inflammatory mediators after injury. The receptors on myelin debris as a result of neural injury activates

7 complement systems initiating phagocytosis of myelin debris. Myelin debris can even diffuse into other areas of the CNS and can affect neurons not in direct contact with myelin debris

(McKerracher & Rosen, 2015). This was shown in studies on a known inhibitory molecule, myelin-associated glycoprotein (MAG). MAG knockout mice displayed improved recovery after

CNS injury (Tang et al., 2001). In wild-type mice, however, the same effects were observed through the introduction of monoclonal antibodies targeting MAG debris released from myelin

(Tang et al., 2001). These studies show that while MAG is an inhibitory myelin protein, the effects are much more potent when released from damaged myelin. (McKerracher & Rosen,

2015) Other myelin receptors including NogoA have also been of interest in understanding the inhibitory effects of CNS myelin (Schwab, 2004). Inhibiting NogoA in the same manner how

MAG was inhibited in previous studies showed noticeable increase in axon growth after injury

(Simonen et al., 2003). Nogo-A knockout mice like MAG knockouts showed a moderate but noticeable increase in axon elongation (Schwab, 2004). The inhibitory effects of NogoA has also been observed when transgenic mice expressing NogoA from PNS myelin were unable to recover from PNS injury (Pot et al., 2002). Which myelin receptor is more potent in inhibiting neural regeneration remains unknown but the identification of these myelin receptors as inhibitory factors have identified additional potential therapeutic targets to induce regeneration.

Intrinsic Factors: Neural Signaling Pathways and Crucial Neural Genes

While targeting extrinsic factors inhibiting regenerative capabilities have shown promising paths in facilitating regeneration, they only show moderate regeneration (Fang & He,

2011; Yiu & He, 2003). This has led to additional studies focusing on intrinsic factors to regeneration in hopes of improving neural regeneration capabilities to a much greater degree

8 than focusing on extrinsic factors (Apara & Goldberg, 2014). The organization of the cytoskeleton in neurons after injury is a major intrinsic factor. It has been shown that CNS neurons form retractions bulbs after injury, while PNS neurons form growth cones (Blanquie &

Bradke, 2018; Ertürk et al., 2007). Growth cones are cytoskeletal extensions from an injured neuron attempting to reach a synaptic target and retraction bulbs are swellings from disorganized cytoskeleton inhibiting growth capabilities (He & Jin, 2016). The formation of retraction bulbs appears to be induced by inhibitory molecules secreted by glial cells after injury (Blanquie &

Bradke, 2018). Inhibitory signaling molecules from glial cells induces the trafficking of fibroblasts (Kawano et al., 2012). The cytoskeleton in neurons can also be targeted to stop the migration of fibroblasts which provides the structural frame work for the formation of glial scars

(Blanquie & Bradke, 2018). The cytoskeleton of neurons serves to traffic vesicles containing fibroblasts which form the structural framework of glial scars. Altering the organization of cytoskeleton to convert retraction bulbs into growth cones reduces scaring and induces axon growth after spinal cord injury in rats (Ruschel et al., 2015). These studies show the capability of overcoming extrinsic barriers to regeneration by altering the organization of the cytoskeleton in neurons after injury.

Certain molecular pathways that are active in CNS neurons are not present in PNS neurons leading to interest in identifying proteins that may play a significant role in inhibiting

CNS regeneration. Proteomic and RNA-seq analysis on X. laevis tadpoles and froglets have revealed differential expression of certain genes expressed specifically in regenerative or non- regenerative stages providing candidate genes to conduct further developmental and regenerative studies on (Lee-Liu et al., 2018). With much interest on T3 as a hormonal factor inhibiting generation, many studies have also been conducted in understanding T3 signaling pathways as

9 well as genes that are upregulated or downregulated specifically in regenerative and non- regenerative stages. A transcription factor that is known to maintain pluripotency of stem cells includes SOX2 (Gaete et al., 2012; Muñoz et al., 2015). The necessity for sox2 expression in order for tail regeneration to occur is shown from transgenics that express a dominant negative form of sox2 which do not have the capability to recover from tail amputation even in the tadpole stage (Gaete et al., 2012). The roles of T3 as a hormonal factor inducing developmental changes in neurons is shown from how hypothyroid mice have increased neural levels of SOX2 (López-

Juárez et al., 2012). Increased levels of T3 on the other hand suppresses sox2 expression (López-

Juárez et al., 2012).

Another prominent transcription factor that is expressed in response to T3 is KLF9 which plays a role in significantly decreasing the capacity for axon growth during development. KLF9 was identified as a crucial developmental protein as high expression levels of klf9 were detected from developing mice RGCs. KLF9 also seems to serve as a mediator protein for T3 functions in inducing developmental loss of regeneration as KLF9 is expressed from a positive feedback mechanism in response to T3. The inhibitory effects of KLF9 on neural growth is evident from studies on klf9 function in mice. In one study, mice injected with a virus containing anti-KLF9 shRNA experienced greater recovery from optic nerve injury (Trakhtenberg et al., 2019). In the same studies that assessed growth axon on brain slices from mice of varying T3 conditions, neural klf9 knockout mice even with T3 treatment still experiences axon growth along the lesion site showing KLF9 as a mediator protein for T3 function (Avci et al., 2012). The long growth and large area of axon growth from klf9 knockouts shows a crucial intrinsic factor inhibiting growth across the lesion site.

10 Summary and Next Steps

Facilitating functional recovery from CNS injury is complex from the various cell types involved and the wide range of neuronal factors involved. Overtime an intricate network of barriers and signaling pathways have continued to evolve in vertebrates to prevent neuroplasticity of neurons. These evolving traits have resulted in the inability of CNS neurons in mammals to recover from injury. Advancing research has allowed us to understand external and internal neuronal factors for developmental loss of regeneration and how to overcome these barriers. Varying factors including but not limited to cellular barriers formed from inflammatory response, intrinsic signaling pathways, and cytoskeletal organization have been found to be involved in the loss of regenerative ability. Targeting these factors have allowed for some degree of axon growth across the lesion site. More recent studies however have now been focused on targeting intrinsic factors in hopes of increasing the capacity for neural regeneration to a greater degree than targeting extrinsic factors. The long regrowth of axons as a result of knocking out certain developmental genes have shown crucial intrinsic factors to target in hopes of facilitating neural growth. As it is already known that neurons do possess some intrinsic capability to regenerate, additional studies could be conducted into understanding how changes in neural signaling pathways in-vivo could affect the capacity for regeneration. Understand neural specific factors responsible for developmental loss of regeneration and how to overcome cellular barriers in other model organisms can provide promising avenues in determining therapeutic approaches for treating spinal cord injury in humans.

11 References Cited

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14 Responses after Spinal Cord Injury. 38, 201–211. Tang, S., Qiu, J., Nikulina, E., & Filbin, M. T. (2001). Soluble myelin-associated glycoprotein released from damaged white matter inhibits axonal regeneration. Molecular and Cellular Neuroscience, 18(3), 259–269. https://doi.org/10.1006/mcne.2001.1020 Toy, D., & Namgung, U. (2013). Role of Glial Cells in Axonal Regeneration. Experimental Neurobiology, 22(2), 68–76. https://doi.org/10.5607/en.2013.22.2.68 Trakhtenberg, E. F., Li, Y., Feng, Q., Tso, J., Paul, A., Goldberg, J. L., & Benowitz, L. I. (2019). Regeneration After Optic Nerve Injury. 22–29. https://doi.org/10.1016/j.expneurol.2017.10.025.Zinc Wheaton, B. J., Sena, J., Sundararajan, A., Umale, P., Schilkey, F., & Miller, R. D. (2020). Identification of regenerative processes in neonatal spinal cord injury in the opossum (Monodelphis domestica): A transcriptomic study. Journal of Comparative Neurology, July, 1–18. https://doi.org/10.1002/cne.24994 Yiu, G., & He, Z. (2003). Signaling mechanisms of the myelin inhibitors of axon regeneration. Current Opinion in Neurobiology, 13(5), 545–551. https://doi.org/10.1016/j.conb.2003.09.006 Yiu, G., & He, Z. (2006). Glial inhibition of CNS axon regeneration. Nature Reviews Neuroscience, 7(8), 617–627. https://doi.org/10.1038/nrn1956

15

Chapter 2 - Blocking neuron-specific thyroid hormone signaling prevents developmental inhibition of spinal cord regeneration

Justin Y. He, Kurt M. Gibbs, Daniel R. Buchholz

16 Abstract:

Unlike in mammals, free-living tadpoles of the frog Xenopus laevis have the ability to regenerate spinal cords. This ability however is lost when tadpoles metamorphose into juveniles.

Previous studies have shown that thyroid hormone (T3) inhibits axon growth across spinal cord lesion sites during metamorphosis, but the cellular targets of T3 signaling responsible for blocking regeneration are not known. To examine the role of T3 signaling specifically in neurons in regeneration, we used tadpoles transgenic for neuron-specific inducible expression of a dominant negative T3 receptor (TRDN). We verified the neuron-specific action of TRDN to inhibit T3-induced changes in gene expression and morphology. Differential recovery from spinal cord injury in transgenic and wild type (WT) tadpoles was then assessed through behavioral studies of swimming ability. In behavioral studies, WT tadpoles treated with T3 experienced a significant delay in recovery. This delay was not seen in TRDN transgenic tadpoles treated with T3. These studies suggest that T3 action on neurons contributes to the loss of spinal cord regeneration during development.

Introduction:

Mammals unlike fish or salamanders do not have the capacity for spinal cord regeneration (Alibardi, 2019; Lee-Liu et al., 2013). However, even in non-regenerative organisms, vestiges of regenerative ability can still be found in embryonic stages (Kundi, 2013).

Some mammals such as opossums are even capable of recovery from spinal cord injury at 1-3 weeks after birth because of the immature level of development at birth (Wheaton et al., 2020).

The mechanisms of regenerative ability and developmental loss of this ability remain elusive, but studies on rat retinal ganglion neurons (RGCs) have served as a model for studying how central

17 nervous system (CNS) neurons lose their regenerative ability (Goldberg et al., 2002). Hormonal factors induce a wide range of intrinsic signaling pathways in RGCs that decrease their capacity for regrowth as development progresses (He & Jin, 2016). These studies have continued to drive research into understanding what hormonal or genetic factors contribute to neural remodeling during nervous system maturation and how to reverse or inhibit the effects of these factors to facilitate recovery from spinal cord injury.

Thyroid hormone (T3) is of interest in current regenerative studies as it is required for inducing physical changes associated with development in vertebrates as well as developmental loss of regeneration (Denver et al., 1997; Gibbs et al., 2011; Wen et al., 2019). T3 regulates the developmental transitions in vertebrates, e.g., birth, hatching, metamorphosis, which includes many brain remodeling events in mammals including humans (Choi et al., 2015; Furlow & Neff,

2006; Schlosser et al., 2002). It is hypothesized that when T3 induces cell differentiation into a mature form as yet incompletely understood, the neurons no longer have the ability to regenerate

(Easterling et al., 2019). Hypothyroidism has been shown to prevent differentiation of neural stem cells possibly maintaining their pluripotency (Bhumika & Darras, 2014). Whether or not inhibiting proliferation of stem cells can maintain regenerative ability is unknown but previous studies have shown that T3 does inhibit axon regrowth after injury in various model organisms.

One such study includes treating zebrafish with iopanoic acid, a chemical that inhibits production of thyroid hormone into its functional form, which allowed for increased recovery from optic nerve crush (Bhumika et al., 2015).

Much focus in understanding developmental loss of regeneration has centered on how T3 affects neurons themselves as opposed to other T3-responsive cells (Bloom, 2014; He & Jin,

18 2016). Specifically, molecular pathways that T3 induces within neurons have been of much interest as it has been found that certain T3 induced transcription factors are correlated with neuronal changes associated with decreased capacity for axon growth (Denver & Williamson,

2009; Fawcett & Verhaagen, 2018; Goldberg et al., 2002; Moore et al., 2011). One of these transcription factors is KLF9 which is heavily involved in developmental neural signaling pathways inhibiting regeneration (Apara & Goldberg, 2014; Hu et al., 2016; Zhang et al., 2017).

In neurons, T3 induces expression of klf9 which allows for increased sensitivity to T3 (Cvoro et al., 2013; Bagamasbad et al., 2012). Neural maturation is heavily dependent on this positive feedback mechanism as klf9 expression induces neurite growth during development. The maturation of neurons due to KLF9 during birth is correlated with decreased regenerative capacity that occurs at this stage of development suggesting KLF9 may be a key player in developmental loss of regeneration (Moore et al. 2009). In RGCs, klf9 expression has been shown to be increased by over 250 fold at developmental stages where the capacity for neurite growth has been significantly decreased (Moore et al. 2009). The inhibitory effects of klf9 expression in neuron regeneration has been shown from in-vitro studies on organotypic culture of mouse brain slice culture. While brain slices from hyperthyroid mice showed decreased axon growth across the lesion site, the inhibition of axon growth as a result of increased levels of T3 is dependent on klf9 expression. Brain slices from klf9 knockout specifically in neurons even with

T3 treatment showed the same axon growth conditions as hypothyroid mice (Avci et al., 2012).

Overall the roles of KLF9 in these neural signaling pathways studies suggest klf9 expression is a major neuronal intrinsic factor inhibiting the ability for neurons to regenerate after injury (Apara et al., 2017). These studies however were not conducted in-vivo nor were axon target finding and

19 behavioral recovery assessed. It is also unknown if the effects of T3 or klf9 expression would apply to spinal cord injury.

Xenopus laevis is an ideal model organism for studying spinal cord regeneration since they have the capacity for spinal cord regeneration as tadpoles but lose this ability as they undergo T3-dependent metamorphosis (Edwards-Faret et al., 2018; Lee-Liu et al., 2014, Beattie., et al 1990). Regeneration studies using X. laevis can provide insight into molecular mechanisms involved in both inducing and inhibiting tissue repair in the tadpole and froglet stages (Gaete et al., 2012). X. laevis tadpoles can also be used to study hormonal effects of spinal cord regeneration. The rearing water of tadpoles can be exogenously supplied with hormones to induce metamorphic changes in virtually every tissue of the body (Buchholz et al., 2003). The effects of altered T3 signaling on spinal cord regeneration has been studied in-vivo using X. laevis tadpoles (Gibbs et al., 2011). Tadpoles treated with methimazole (blocks synthesis of T3) experienced blocked metamorphic development maintaining their larval stages which also allowed for axon growth across the lesion site following spinal cord transection (Gibbs et al.,

2011). At the behavioral level, methimazole-treated tadpoles were able to recover swimming functions close to that of non-chemically treated control tadpoles (Gibbs et al., 2011). These recovery events were not observed in tadpoles that underwent premature metamorphosis from T3 treatment (Gibbs et al., 2011). These studies have led to the conclusion that maintenance of larval features in X. laevis tadpoles is necessary to retain the ability for spinal cord regeneration

(Gibbs et al., 2011).

While these studies show a profound role for T3 in spinal cord regeneration, the critical targets for T3 in blocking regeneration, e.g., neurons, glia, immune cells, vasculature, are still

20 unknown (Bhumika & Darras, 2014; Prezioso et al., 2018; Trentin, 2006). Here, we examine in- vivo effects of lack of T3 signaling specifically in neurons using doxycycline- (Dox-) inducible dominant-negative thyroid hormone receptor (TRDN) transgenic tadpoles. In the presence of

Dox, transgenic tadpoles exhibit neuron-specific GFP and TRDN expression, thereby precluding

T3 responses in neurons. Previous studies have already shown the developmental effects of tadpoles with constitutive, neuron-specific expression of the NbT/TRDN transgene, namely limb paralysis and death at the climax of metamorphosis (Marsh-Armstrong et al., 2004). Here, we study the effect of impaired T3 signaling in neurons in-vivo on spinal cord regeneration using neuron-specific TRDN transgenic tadpoles. We verify neuron specificity of TRDN action through study of morphological changes in tadpoles after T3 treatment and expression of two genes known to be upregulated in response to T3: Krüppel-like f\actor 9 (klf9) and thyroid hormone receptor-ß (thrb), a T3 receptor isoform. After verifying functionality of transgenics, we study the role of lack of neural T3 signaling in recovery from spinal cord injury through behavioral studies.

Materials and Methods

Animals and Husbandry: The doxycycline (Dox)-inducible NbT/TRDN-GFP (NbT:DNTR

(inducible)) transgenic line was a gift from Donald Brown of the Department of Embryology at the Carnegie Institute of Washington [Fig. 1] (Das & Brown, 2004). The pDRTREG transgenic animals were from the lab colony [Fig. 1] (Kerney et al., 2008). To produce offspring for experiments, adult Xenopus laevis were injected with 0.5 ug/uL ovine luteinizing hormone

(oLH), 4 uL for females, 1 uL for males. Eggs were collected approximately 15-18 hours after hormone injections. Offspring were reared to Nieukoop and Faber (NF) stage 45 (beginning of

21 feeding) and then treated with 50ug/mL Dox in rearing water for 24 hours and then sorted into 6

L tanks at an occupancy of 30 tadpoles each with transgenic tadpoles (green fluorescence) or wild-type (non-fluorescent). Fluorescence was detected under a Leica fluorescence stereo dissecting microscope using a GFP filter. Tadpoles were reared in 22C and fed Sera Micron food everyday ad-lib., with water changes every two days. For each experiment, all tadpoles were exogenously treated with 50 ug/mL Dox and 1-5 nM T3 depending on experiment. No food was given during experiments.

Gene Expression Studies: Quantitative PCR (qPCR) analysis was conducted on brain and tail samples. Tadpoles were obtained by crossing NbT/TRDN (inducible) x wild type. At NF stage

54, transgenic and wild-type tadpoles were treated with 50ug/mL Dox for 48 hours. After 48 hours, water was changed and 50ug/mL Dox and 0 or 2 nM T3 were added. Transgenic tadpoles were treated with 2 nM T3 (n = 8 tail, n = 9 brain), and wild type tadpoles used were separated into 2 nM T3 (n = 8 tail, n = 9 brain) and non-T3 treated (n = 10 tail, n = 9 brain) groups. Tissue harvesting was performed on tadpoles after 24 hours of T3 treatment. Brain and tail tissue were harvested as described (Patmann et al., 2017), snap frozen on dry ice, and stored at -80°C until

RNA extraction with TriZol (Invitrogen). cDNA was synthesized using ThermoFisher High

Capacity Reverse Transcription Kit then subjected to qPCR analysis for klf9 and thrb normalized using rpl8 as a reference gene (Dhorne-Pollet et al., 2013). The 2- ΔΔCT method was applied to quantify gene expression (Livak & Schmittgen, 2001).

Morphological studies showing the effect of TRDN on brain remodeling: To visualize neural tissue and compare T3-induced brain remodeling in tadpoles with and without neuron-specific

TRDN expression, offspring of NbT:TRDN (inducible) x pDRTREG were used. To achieve

22 strong neuron-specific GFP and TRDN expression (+TRDN group), NF stage 47 tadpoles were pretreated with Dox for two days (n = 5). To achieve strong GFP expression without TRDN (-

TRDN group), NF stage 47 tadpoles were pre-treated with 50 ug/mL Dox for three days followed by Dox removal for 3 days to induce long-lasting GFP protein and give time for

TRDN-GFP mRNA to degrade (n = 5) (Kerney et al, 2012). To compare T3-induced brain remodeling between these two groups, tadpoles were then treated with T3 for 6 days, where the

+TRDN group had continued Dox treatment, and the -TRDN group did not (Fig. 2). Dox at

50ug/mL has no known effect on metamorphic progression (Kerney et al, 2018). One exemplar tadpole from each treatment group was used for fluorescent photography. Pictures were taken every day under a Leica fluorescence stereomicroscope during hormone treatment. Water and chemicals were changed every day.

To quantify morphological changes in the head, NF stage 54 tadpoles from NbT:TRDN

(inducible) x wild-type were treated with 50ug/mL Dox for 2d followed by 6d of 2nM T3 and

Dox. Water, Dox, and T3 were changed every 48 hours. Each day during treatment with T3, transgenic and wild-type (n = 10) tadpoles were anesthetized in 0.02g/200 mL tricaine (MS-222,

Fisher Scientific) followed by measurements of olfactory nerve, brain length, brain width, and head width (defined as dorsal width across the two eyes). Measurements were taken with a Leica stereo dissecting microscope fitted with a reticle at the following magnifications (olfactory nerve

3X magnification, brain width and brain length 1.25X magnification, head width 2X magnification). Measurements using the reticle were converted to millimeters. Sample size was n = 10 with 2 deaths per group.

23 Spinal Cord Transection: Spinal cord transection was performed on anesthetized tadpoles

(0.02g/200 mL MS-222) by fully transecting the spinal cord between the 3rd and 4th vertebra and passing the tips of fine forceps through the injury site. Tadpoles were fully immobile posterior to the injury site, but movement still occurred anterior to the injury site. Tadpoles showing any movement posterior to the injury site one day after spinal cord transection were discarded. Dox treatment (50 ug/mL) began in all experimental groups on the day of spinal cord transection and maintained for the entire 21-day observation period. T3 treatment (1 nM changed every 2 days) began 48 hours after spinal cord transection. Recovery of tadpole behavioral movements were tracked for three weeks. At the end of each one-week interval, tadpoles were assessed for the following behavior characteristics: volitional swimming - tadpoles move voluntarily in a particular direction, fictive swimming - movement of the distal tail while hovering or floating to stay in place, righting reflex - the ability of the tadpole to maintain its correct posture while swimming. Behaviors were assigned a score on a scale of 1-3: 1 (absent), 2

(weakly present), or 3 (moderately present to normal). A score for each behavioral category was assigned to a tadpole based on movement induced after touching with a pipette 3-4 times (Gibbs et al., 2011). Samples size was n = 9 with 0-2 deaths per treatment.

Statistical Methods: Significant differences in morphology measurements between wild type and transgenic tadpoles were determined using a two-way repeated measures ANOVA in IBM SPSS.

Pairwise comparisons were conducted for each day of measurement. Differences in gene expression of thrb and klf9 across treatment groups were analyzed using a one-way ANOVA followed by Tukey Honestly Significant post hoc analysis in R. For behavioral data, pairwise comparisons were made across each treatment group through the Mann Whitney U-test, a statistical test for non-parametric ranked data in R.

24 Results

Neuron-specific inducible expression of a dominant negative thyroid hormone receptor

The NbT:TRDN (inducible) transgene constitutively expresses the rtTA protein only in neurons (Fig. 1A) (REF). In the presence of Dox, rtTA binds to the tetO promoter inducing the expression of TRDN, a dominant negative form of the T3 receptor. A faint GFP fluorescence is also expressed in the neurons of these tadpoles because of the GFP fused to TRDN. To better visualize transgene expression in neurons, double transgenic tadpoles were reared from pDRTREG x NbT:TRDN (inducible) adults. The pDRTREG transgene consists of a tetO promoter controlling unfused GFP, which fluoresces much brighter than GFP-TRDN (Fig. 1A)

(REF). Transgenic tadpoles harboring the pDRTREG also constitutively express DsRed2, a red fluorescent protein, in the eyes as a transgenic marker.

Morphological studies showing effect of TRDN on brain remodeling: To readily visualize the effect of neuron-specific TRDN expression on T3-induced metamorphosis, we used transgenic animals with Dox-inducible expression of TRDN and GFP in neurons and treated them with T3

(Fig. 2A). We first treated double transgenic tadpoles (Fig. 1) with Dox three days to induce

TRDN and GFP. Dox removal allows TRDN mRNA and protein to degrade as well as GFP mRNA. However, GFP protein is stable for weeks and so persists to label the nervous system.

These animals were the -TRDN group and received no more Dox. The +TRDN group was also double transgenic and was treated with Dox for 1 day, as well as during the subsequent T3 treatment. Both groups were then treated with T3 for 6 days to induce metamorphic changes

(Fig. 2B). At 3 days after T3 treatment, gills began to resorb in both -TRDN and +TRDN animals. By 6 days, Meckel’s cartilage began to protrude and gills were fully resorbed in both treatment groups at day 6 and olfactory nerve and brain length were much reduced. To quantify

25 differences in T3 response between -TRDN and + TRDN tadpoles, we measured changes in olfactory nerve length, brain length, brain width, and head width on wild type and transgenic tadpoles (Fig. 3). Significant differences in changes in olfactory nerve length and head width were observed where +TRDN animals showed a delay in changes (two way repeated measures

ANOVA: olfactory nerve - F = 79.82 p = <.001, head width – F = 75.191 p = <.001).

Significant differences were not observed for brain length and head width.

Gene Expression Studies: For these studies, wild type (WT) and NbT:TRDN (inducible) transgenic premetamorphic tadpoles (NF stage 54) were used and treated with or without 2 nM

T3 for 24 hrs. The expression of klf9 and thrb from tail and brains of tadpoles from WT-T3,

WT+T3, and TRDN+T3 tadpoles were quantified using quantitative PCR (Fig. 4). In tail samples, klf9 expression was induced to the same degree in TRDN+T3 and WT+T3 samples

(one-way ANOVA – F = 7.47, p = 0.003). For thrb expression in tails, it was significantly induced by T3 in WT animals, but its expression was intermediate in TRDN transgenic animals

(one-way ANOVA – F = 3.59, p = 0.046). In brain samples, both klf9 and thrb expression increased in WT+T3 tadpoles but was not significantly induced in TRDN transgenic animals

(klf9 one-way ANOVA – F = 11.6147, p < 0.001; thrb one way ANOVA – F = 12.79, p <

0.001).

Behavioral Recovery: For these studies, WT and NbT:TRDN (inducible) single transgenic premetamorphic tadpoles were again used in WT-T3, WT+T3, and TRDN+T3 groups.

Immediately following spinal cord transection, tadpoles in WT-T3, WT+T3, and TRDN+T3 groups were completely paralyzed posterior to the injury site and sank to the bottom without any signs of swimming. Movement including breathing and other head movements were still present anterior to the injury site. Immobile tadpoles were also immediately treated with 50 ug/mL Dox

26 and two days later 1 nM T3 treatments began. Preliminary studies used varying T3 concentrations to determine the optimal T3 concentration to minimize tadpole deaths. Most tadpoles in all treatment groups at 7 dpt remained completely paralyzed in swimming ability.

The tadpoles that did show any sign of movement at 7 dpt swam sharply upwards in circular patterns for a short period of time before sinking to the bottom.

At 14 dpt, WT-T3 and TRDN+T3 both experienced a dramatic increase in both fictive and volitional swimming, whereas WT+T3 was delayed in recovery. Righting reflex is the most impaired at 14 dpt as most tadpoles alternated between dorsal side down and up while swimming showing difficulties in maintaining the correct posture while swimming. Tadpoles that were able to recover close to the full extent before spinal cord transection were able to swim in more defined directions while also regaining the ability to remain afloat within a particular area.

WT+T3 tadpoles continued to move in sporadic patterns typical of volitional swimming at 7 dpt, but most appeared to remain stunted in the ability to swim in a well-defined direction. WT+T3 tadpoles that displayed signs of volitional swimming at 14 dpt also seemed unable to remain afloat for the extended period of time seen in WT-T3 tadpoles, a sign of severely inhibited fictive swimming. At 21 dpt, more WT-T3 and TRDN display relatively normal swimming patterns.

Many tadpoles still alternate between dorsal side up and down while swimming. Surprisingly,

WT+T3 tadpoles recovered close to the extent that WT-T3 and TRDN+T3 tadpoles were able to at 21 dpt. Results from the Wilcoxon Mann Whitney U-test were as follows: Volitional –

TRDN+T3:WT+T3 U=12 p = 0.0367; WT+T3:WT-T3 U=8, p = 0.0119; Fictive –

TRDN+T3:WT+T3 U=12 p = 0.0365; WT+T3:WT-T3 U=8 p = 0.0348). [Justin - explain what these numbers tell us.]

27

Discussion:

We sought to expand on previous studies by assessing the role of T3 inhibition in maintaining spinal cord regeneration ability in-vivo. Differences in premature metamorphosis and gene expression in WT and transgenic tadpoles were assessed by observing differential recovery from spinal cord transection. We used transgenic animals overexpressing TRDN inducible under control of an NbT promoter which impairs T3 signaling in neurons. We found that transgenic animals expressing TRDN in neurons treated with T3 did not differ from WT control animals in behavioral recovery from complete spinal cord transection. Surprisingly, WT animals treated with T3 were also able to recover movement but after a significant delay.

We first verified the functionality of neural specific TRDN expression in transgenic tadpoles by analyzing morphological and gene expression changes in response to T3 as seen previously with this transgenic line (Marsh-Armstrong et al., 2004). We expected neural-related delays in response T3 but no delays in other tissues. Indeed, we found a significant delay in olfactory nerve shortening which may be caused by reduced T3 action on olfactory neurons blocking their normal T3-induced change. The significant delay in head width reduction is not clear but may relate to indirect effects on gill remodeling from reduced T3 signaling in neurons.

The delay in T3 response in olfactory neurons in transgenic tadpoles are consistent with our gene expression studies. It is expected that T3 response gene induction between transgenic and WT tadpoles treated with T3 would differ in brains but not tails. Indeed, klf9 and thrb gene induction occurred in tails and not brains in T3-treated NbT:TRDN (inducible) tadpoles. Tails are expected to have few neurons relative to other cell types and thus blockade of T3 action in

28 neurons would not have an effect on T3 response gene induction. klf9 and thrb are induced by T3 outside neurogenic zones in non-proliferating cells, i.e., in differentiated or differentiating neurons expressing NbT (Denver & Williamson, 2009; Hoopfer et al., 2002). Thus, inhibition of

T3 responses in neurons overexpressing TRDN is expected to significantly reduce klf9 and thrb expression in brain. The induction of T3 response genes in tail and not brain confirms neural- specific inhibition of T3 responses in Nbt:TRDN (inducible) transgenic tadpoles.

Our results showing significant impairment in behavioral recovery from spinal cord transection in WT tadpoles treated with T3 is consistent with previous studies showing T3- dependent loss of regenerative ability. However, the cell type(s) responsible for this loss of regeneration were not known. Using transgenic overexpression of TRDN that blocks T3 signaling specifically in neurons, we showed that neurons are a cellular target of T3 capable of blocking regeneration. In addition, all other cell types were subjected to T3 signaling in our transgenic system indicating T3 activation in non-neuronal cells might not be sufficient to inhibit behavioral recovery. These results suggest that T3 acted on axotomized hind brain neurons to achieve a developmental state where they no longer had the potential to extend axons and regenerate. Previous studies in tadpoles used high T3 doses that are lethal to tadpoles within about 7 days and thus recovery beyond 7 days of T3 treatment was not known. Our experiments used low T3 doses which enabled long-term survival and surprisingly allowed for behavioral recovery albeit with a delay. Because young tadpoles were used, they were still growing and developing, and thus neural stem cells were still present and producing more neurons (REF).

Thus, the recovery observed in the presence of T3 may be due to neural stem cells, stimulated or not by T3 to proliferate, differentiating and reinnervating targets posterior to the lesion site. The

29 delay thus would come from the time required to go through neuron differentiation process and targeting which would take longer than regrowth of cut axons.

What our studies do add on compared to studies that assessed intrinsic axon regeneration from lack of T3 signaling is showing the capability of regaining functionality in locomotion close to pre spinal cord transection conditions. By conducting behavioral studies in-vivo, we also show the potential capability of recovering movement through lack of T3 signaling in neurons.

Our studies may also confirm previous hypotheses that developmental loss of regeneration of neurons is explained at least in part by intrinsic factors in neurons. This is consistent with evidence of neurons capable of overcoming external barriers as shown from studies on embryonic tissue grafted into adult CNS which shows how embryonic axonal tissues were still able to proliferate despite the inhibitory environment of adult CNS (Fawcett, 2020; Syková et al.,

2006). Our studies provide evidence of lack of intrinsic T3 signaling in neurons to be sufficient to allow for improvement in spinal cord regeneration in an in-vivo situation.

30

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34

Figure Legends

A. Transgenic line 1 – NbT:TRDN (inducible) Transgenic line 2 – pDRTREG-HS4

NbT rtTA tetO TRDN GFP X tetO EGFP CRY DsRed

B. WT Double transgenic

Bright Field GFP Bright Field GFP

No Dox

Dox

Figure 1 – Tissue-specific inducible control of TRDN in neurons. A. To achieve high TRDN

(dominant negative thyroid hormone receptor) and GFP (green fluorescent protein) expression in neurons, two transgenic lines of X. laevis were crossed. First, the NbT/TRDN (inducible) line harbors a transgene with the neuron-specific promoter NbT (neural beta tubulin) controlling the

Dox-activated engineered transcription factor rtTA and a tetO promoter controlling TRDN fused to GFP (REF). Because GFP is dim when fused to TRDN (not shown), the first transgenic line was crossed with a second transgenic line harboring the TRE (tetracycline response element) promoter controlling GFP and a lens-specific crystallin promoter driving DsRed fluorescent protein constitutively in the eyes (REF). B. In double-transgenic offspring, Dox added to the rearing water of premetamorphic tadpoles binds to rtTA expressed only in neurons which then

35 induces expression of TRDN-GFP and GFP from the very similar tetO and TRE promoters. In the absence of Dox, no fluorescence is expressed except the constitutive red fluorescence in the eyes (not shown). Wild-type (WT) tadpoles show no fluorescence with or without Dox.

A. Days -6 -5 -4 -3 -2 -1 1 3 6 Dox ✓ ✓ ✓ ------TRDN T3 ------✓ ✓ ✓ Dox - - - - - ✓ ✓ ✓ ✓ +TRDN T3 ------✓ ✓ ✓ B. Day 1 Day 3 Day 6

-TRDN

+TRDN

Figure 2 – Neuron-specific expression of TRDN delays thyroid hormone-induced brain remodeling. A. To achieve two groups of tadpoles with high neuron-specific GFP expression that differ in TRDN expression, double transgenic animals (described in Fig. 1) were treated with different Dox regimes. For the -TRDN group (n = 5), tadpoles were treated with 50ug/mL Dox

36 for 3 days followed by three days in the absence of Dox to induce GFP and allow time for

TRDN-GFP mRNA to decay while GFP protein remains due to its long half-life. For the

+TRDN group (n = 5), tadpoles were treated with 50ug/mL Dox for 1 day to induce TRDN-GFP and GFP. For both groups, 10nM T3 was then added to induce metamorphic changes. B. Images were taken after, 1, 3, and 6 days of T3 treatment. The effect of T3 on gill resorption (bracket) was observed at Day 3 and complete by Day 6. Changes in brain size (arrow head) and olfactory nerve length (arrow) can also be seen on Days 3 and 6.

A. Olfactory Nerve B. Brain Length 3 6 WT TRDN WT TRDN 5

2 4 * mm 3 mm 1 * 2

* 1

0 0 0 1 2 3 4 5 6 0 1 2 3 4 5 6 Days After T3 Treatment Days After T3 Treatment C. Brain Width D. Head Width 3 WT TRDN 10 WT TRDN 8 2 * * 6 * mm mm 4 1 2

0 0 0 1 2 3 4 5 6 0 1 2 3 4 5 6 Days After T3 Treatment Days After T3 Treatment

Figure 3 – Quantification effect of neuron-specific expression of TRDN on T3-induced brain remodeling. Changes in (A) olfactory nerve length, (B) brain length, (C) brain width, and (D) head width were measured daily during 6 days of treatment with 50ug/mL Dox and 10nM T3 in wild-type (WT) and transgenic tadpoles (NbT:TRDN (inducible)) . (TRDN+T3: N = 10, 1 death at day 5, 1 death at day 6; WT+T3: N = 10, 2 death at day 5). Each anatomical feature was

37 measured using the reticle of a Leica stereo dissecting microscope and then coverting to millimeters. Mean measurements ± SE were tabulated for each day of treatment. Asterisks above bars indicate significant difference between WT and transgenic tadpoles for the indicated day using from two-way repeated measures ANOVA.

16 7 A. Tail klf9 B. Tail thrb 14 6

12 5

expression 10 expression 4 klf9 klf9 8

thrb 3 6 2 4 B B A B AB A 2 1 normalized rel. rel. normalized

0 rel. normalized 0 WT-T3 WT+T3 TRDN+T3 WT-T3 WT+T3 TRDN+T3 100 40 C. Brain klf9 35 D. Brain thrb 80 30 expression 60 expression 25 klf9 klf9

thrb 20 40 15 A 10 20

normalized rel. rel. normalized A B A 5 A B normalized rel. rel. normalized 0 0 WT-T3 WT+T3 TRDN+T3 WT-T3 WT+T3 TRDN+T3

Figure 4 – Neuron-specific expression of TRDN inhibits thyroid hormone-induced gene expression in brain but not tail. A-B) Tails (n = 8-10) and (C-D) brains (n = 9) were harvested from tadpoles at NF stage 54 (premetamorphosis) treated as follows: wild type (WT) control,

WT with 2 days of 2 nM T3, transgenic (NbT:TRDN (inducible) with 2 days of 2 nM T3. All tadpoles were treated with 50ug/mL Dox for 48 hours, given a water change, and then treated with ± T3 and more Dox (50 ug/mL) for 24 hrs before tissue harvest. RNA was extracted, followed by cDNA synthesis, quantitative PCR for klf9, thrb, rpl8 (reference gene). Bars indicate mean ± SE of klf9 and thrb normalized to rpl8 relative to a control sample, and letters indicate

38 pairwise significant differences letters determined by one-way ANOVA followed by Tukey

Honestly Significant post hoc analysis (a = 0.5).

A. Volitional C. Righting 3 * 3 WT WT+T3 TRDN+T3 * WT WT+T3 TRDN+T3

2 2 Score Score 1 1

0 0 1 dpt 7 dpt 14 dpt 21 dpt 1 dpt 7 dpt 14 dpt 21 dpt B. Fictive 3 Scoring System * WT WT+T3 TRDN+T3 * 1 - Immobile

2 2 - Present but abnormal 3 - Normal Score 1

0 1 dpt 7 dpt 14 dpt 21 dpt

Figure 5 – Neuron-specific expression of TRDN prevents T3-dependent delay in behavioral recovery from spinal cord transection (TRDN+1 nM T3: N=9, 1 death at 7 dpt, 1 death at 21 dpt;

WT+1 nM T3: N = 9, 1 death at 7 dpt, 1 death at 14 dpt; WT-T3: N = 9). Spinal cord transection was performed on wild-type (n = 18) and NbT:TRDN (inducible) (n = 9) transgenic animals, and

Dox (50 ug/mL) treatment began the day of transection to induce neuron-specific TRDN expression. Treatment of 1nM T3 began 48 hrs after spinal cord transection in transgenic animals and half of the WT tadpoles. Dox and T3 treatments were maintained for the 3-week observation period. Behavioral recovery of tadpoles was assessed by scoring A) volitional, B) fictive, and C) righting as 1 (absent), 2 (weakly present), or 3 (moderately present to normal). Mean scores +/-

SE were determined for each 1week interval for three weeks after spinal cord transection. (dpt –

39 days post transection) Brackets indicate significant differences for pairwise comparisons from

Mann Whitney U-test for each time point.

Chapter 3

40

How to induce regenerative capabilities in CNS neurons has been of great interest in modern medicine. Understanding how to achieve such a feat has been complicated from the various neuronal internal and external barriers to regeneration that must be overcome (Fawcett,

2020; Lee-Liu et al., 2013). In many vertebrates, regenerative ability as development progresses

(Geoffroy et al., 2016). In amphibians, the developmental loss of regeneration occurs in the transition from froglet to tadpole (Gaete et al., 2012; Gibbs et al., 2011). The developmental transition between froglet to tadpoles is analogous to the transition from embryo to birth in mammals including humans which is when loss of regeneration occurs (Buchholz, 2015; Gibbs et al., 2011). Various model organisms have been used in understanding the mechanisms of regeneration and how it is lost. In these studies, regeneration was facilitated by manipulating a wide range of external and internal neural factors leading to discussions on potential therapeutic targets to allow for regeneration (Goldberg et al., 2002; Jerry Silver, Martin E. Schwab, 2015;

Mills et al., 2015). Here I discuss the implications of these studies and previous hypotheses as well as what our current studies tell us as well as possible future directions in manipulating neurons into axon regrowth after injury.

What do we target?

Many initial studies on neural regeneration have been focused on the external environment of neurons as the external environment of PNS and CNS neurons differ explaining differences in regenerative ability between the two (Lang et al., 1995; Reier et al., 2017; Zhang et al., 2017). Recent studies have been focused on the intrinsic properties of neurons leading to developmental loss of regeneration in as it has also been found that removing the inhibitory

41 activities only allows for limited regeneration (Fang & He, 2011). While it has been shown the modifying the intrinsic properties by neurons by inhibiting T3 response or knocking out certain developmental genes allows for increased axon growth, many of these studies were conducted in an in-vitro environment and thus do not account for the possibility of the external factors also affecting neural regeneration in vivo (Avci et al., 2012; Moore et al., 2011). We have shown for the first time in-vivo inhibition of T3 in neurons facilitating increased recovery from spinal cord injury. We did not test however whether or not spinal cord neurons can be explained by the inherent properties of axons rather than the known external barriers. On the other hand, therapeutic bone marrow transplants in patients with spinal cord injury show the capability of neurons to proliferate in such an environment. Previous studies showing how administration of

Epothilone B, a drug known to reorganize neural cytoskeleton of neurons and reduce scarring, also shows that neurons do possess some intrinsic capability to overcome an inhibitory environment (Ruschel et al., 2015).

Treatment of Epothilone B in some model organisms shows that reducing glial scar formation at the injury site can facilitate regeneration (Griffin & Bradke, 2020; Ruschel et al.,

2015). In non-regenerative organisms, inflammatory response after spinal cord injury are formed through extracellular matrix components (ECM) components secreted by astrocytes and microglial cells (Yiu & He, 2003). Secreted ECM components form an intricate network of signaling pathways and molecular mediators that triggers the action of immune cells forming glial scars. Among these ECM components, the family of chondroitin sulfate proteoglycans

(CSPGs) has been of great interest (García-Alías & Fawcett, 2012). The application of bacterial enzyme chondroitinase ABC (ChABC) in mice has allowed for recovery from spinal cord injury by targeting CSPGs (Bloom, 2014). How ChABC works to alter CSPGs has not been extensively

42 studied. It has been suggested however that CSPGs interfere with cell-to-cell adhesion formation of neurons which is a pre-requisite for synapse formation (Crespo et al., 2007). Treatment of

ChABC would then allow for cell-to-cell adhesion to occur (Crespo et al., 2007). However, other researchers have noted that treatment through ChABC does not allow for long distance axon regeneration and functional recovery is very limited (Fang & He, 2011). We are unsure how spinal cord neurons in our tadpoles from our behavioral studies were able to recover functionality. It is also not known how altering the intrinsic properties of neurons can overcome the inhibitory environment created by glial cells. Addressing these questions could be of great interest for future studies in understanding how neurons can regain regenerative ability.

Recent advances in our understanding of neural regeneration

Recent studies have shown klf9 expression as a major intrinsic factor inhibiting regeneration (Galvao et al., 2018; Hu et al., 2016; Knoedler & Denver, 2014). KLF9 is a developmental protein that induces developmental loss of regeneration by mediating neural signaling pathways that differentiate neurons into their mature form (Avci et al., 2012).

Knocking out klf9 in neurons reverses these effects as seen from previous studies on organotypic brain slice culture from mice (Avci et al., 2012). The dramatic increase in length and area of axon growth across the lesion site from klf9 knockouts shows klf9 as a crucial neural intrinsic factor preventing proliferative ability of axons after injury (Avci et al., 2012). As klf9 is positively expressed from T3, transgenics with neural specific inhibition of T3 from our studies have also shown decreased klf9 expression in brain. The significant decrease in klf9 expression in

TRDN+T3 and WT-T3 brain samples from our studies seems to correlate with the increase in recovery observed at 14 dpt in our studies. The decrease in klf9 expression also correlates to the

43 decrease in trb expression as trb is positively expressed from KLF9. The klf9/trb axis is crucial for neural development programs as trb autoinduction is increased during post embryonic development in X. laevis tadpoles (Bonett et al., 2011; Denver et al., 1997). The autoinduction of trb in tadpoles is analogous to the molecular developmental pathways that are induced during development in humans before birth (Buchholz, 2015). While we did not test directly the effects of inhibiting klf9 expression in neurons, it is reasonable to hypothesize that inhibiting T3 in spinal cord neurons also inhibited the positive feedback expression of klf9 leading to enhanced regeneration in our transgenic tadpoles.

We have focused our studies to understand how lack of T3 response could allow for recovery from spinal cord injury because of the likelihood of axon growth to differ between cerebral and spinal cord neurons. The functional recovery of our transgenic X. laevis shows the capability of the spinal cord in tadpoles to regenerate. Inhibiting T3 as a therapeutic target may be difficult as T3 differentiates neurons into a non-pluripotent form. Thus, inhibiting T3 function after spinal cord injury in organisms that already lack regenerative ability would not allow for recovery. Inhibiting T3 shows that maintenance of larval features is required for spinal cord injury recovery in tadpoles however, so de-differentiating neurons into their early developmental state may be the most viable approach. In perinatal rats, Schwann cells after neural injury revert to their immature form and dedifferentiate into axons at the injury site. Blocking de- differentiation of Schwann cells stops the healing process.

Implications of facilitating regeneration?

Whether or not dedifferentiating neurons may have significant side effects is unknown as there are hypothesized evolutionary costs associated with regenerative ability. In lizards, tail

44 autotomy comes with a significant energetic cost as the metabolic rate of lizards increases significantly during tail synthesis (Naya et al., 2007). Some organisms such as fiddler crabs have a fidelity cost (Bely & Nyberg, 2010). In this case, regeneration is often imperfect and imperfect structures might harm the organism more than the amputated structure (Bely & Nyberg, 2010).

Imperfect spinal cord regeneration was observed from non T3 WT tadpoles at the end of the recovery period from our behavioral studies as many tadpoles maintained an irregular posture while swimming. Despite the imperfect regeneration, however, such a scenario would still be much more favorable than complete paralysis from spinal cord injury. It has also been hypothesized that sustaining memory and the ability to function in social situations are dependent on the neurons remaining in a non-pluripotent state. Such functions would be lost from neural remodeling. Tradeoffs of regeneration in X. laevis, as well as other model organisms with a closer evolutionary distance to humans, have not been studied extensively, but such studies can provide insight into loss of regeneration through evolutionary time as well as other physiological tradeoffs to consider in inducing regeneration in humans.

Concluding Remarks

Numerous studies focusing on understanding the barriers to regeneration and how to awaken regenerative potential in neurons have pointed to a wide range of extrinsic and intrinsic factors that can be targeted. It was initially believed that external factors such as formation of glial scars were the main impediment to regeneration. Recent studies however have now shown the intrinsic abilities of neurons to regenerate (He & Jin, 2016; Lemmon & Goldberg, 2009).

Previous in-vitro studies show that knocking out critical developmental genes including klf9 or stopping positive feedback expression of klf9 through T3 inhibition has allowed for dramatic

45 increase of axon growth (Avci et al., 2012). We have expanded on these findings by studying T3 function in X. laevis tadpoles in-vivo and assessing if normal swimming behaviors would return after spinal cord injury.

Our findings confirm that inhibiting T3 function in neurons does facilitate recovery. We also show that lack of T3 signaling in neurons allows for functional recovery, while also confirming other findings that premature metamorphic T3 treatment on tadpoles accelerated developmental loss of regeneration. How our findings could lead to new therapeutic approaches for spinal cord regeneration may be difficult as T3 was inhibited in transgenic tadpoles at a developmental stage where regeneration is still possible. From previous conclusions that maintenance of larval features in X. laevis is required for regeneration, it is likely that future studies will be focused more on the intrinsic ability of axons to regenerate. Inhibiting T3 response in transgenics at the froglet stage would unlikely allow for recovery. Potential therapeutic approaches would most likely be focused on reversing the developmental effects of

T3. Our recent advances in understanding the mechanisms of inhibiting spinal cord regeneration provide exciting insight into how we can cure spinal cord regeneration in humans.

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