Improving Axonal Regeneration: Side-to-Side Bridges Coupled with Local Delivery of Glial Cell Line-Derived Neurotrophic Factor (GDNF)

Transcription

1 Improving Axonal Regeneration: Side-to-Side Bridges Coupled with Local Delivery of Glial Cell Line-Derived Neurotrophic Factor (GDNF) by MARIA CECILIA ALVAREZ VERONESI B.Sc., University of Calgary, 2011 A thesis submitted in conformity with the requirements for the degree of MASTER OF APPLIED SCIENCE Institute for Biomaterials and Biomedical Engineering University of Toronto Copyright by Maria Cecilia Alvarez Veronesi, 2013

2 Improving Axonal Regeneration: Side-to-Side Bridges Coupled with Local Delivery of Glial Cell Line-Derived Neurotrophic Factor (GDNF) M. Cecilia Alvarez Veronesi Master of Applied Science (MASc) Institute of Biomaterials and Biomedical Engineering University of Toronto 2013 Abstract Chronic denervation and chronic axotomy present independent barriers for axonal regeneration. Chronic denervation occurs when nerves are no longer connected to their neuronal cell bodies; chronic axotomy occurs when neurons are not connected to their end-organs for prolonged periods of time. The harmful effects of chronic denervation can be addressed by the side-to-side bridge surgical technique. Additionally, the negative effects of chronic axotomy can be reversed by GDNF delivery to the nerve. The experiments in this thesis were designed to evaluate nerve regeneration in a rat model of chronic injury after treatment with local GDNF delivery, side-to-side bridge protection, or both. The GDNF delivery system consisted of poly(lactic-co-glycolic acid) microspheres embedded in fibrin for controlled delivery of GDNF. Overall, the side-to-side bridges technique was effective in protecting against the negative effects of chronic denervation in several measures regardless of treatment with or without GDNF. Local delivery of GDNF did not increase axonal regeneration or functional recovery. ii

3 Acknowledgements First and foremost, I would like to thank my supervisors, Dr. Gregory Borschel and Dr. Tessa Gordon. Thank you for your patience and support during the past two years, and for sharing your knowledge, advice, and constructive criticism for the completion of this thesis. It has truly been a learning and rewarding experience. I must also thank my thesis committee members: Dr. Molly Shoichet and Dr. Michael Salter. Thank you for contributing to the project with your expertise and insight. I would like to thank my lab mates - and friends - at the Borschel lab for these past two years. Christine Lafontaine, Eva Placheta, Matthew Wood, and Steve Kemp, thank you for welcoming me to the lab when I first started. Thank you for your patience while I was learning all the techniques that we use in the lab. As well, Jennifer Chang, Cameron Chiang, Mike Hendry, Edward Liu, and Mike Willand: you guys always made coming in to the lab more fun. Thank you for the encouragement, advice, and all the laughs we shared. Joseph Catapano and Kasra Tajdaran, best of luck for the next two years! This thesis work could not have been completed without the help of several people. Matthew Wood and Christine Lafontaine, thank you for teaching me rat surgeries and most of the techniques I used for outcome measures. Special thanks to Matt, your advice throughout this project was invaluable. Eva Placheta, I learnt a lot from your meticulous work ethic during surgeries and completion of experiments. Cameron Chiang, thank you for the countless hours you spent imaging and analyzing the histomorphometry data for this project, it was truly essential to this work. Mike Willand, thank you for your help with the iii

4 electrophysiology experiments. It really provided valuable insight for my project and I could not have done it without your expertise. Steve Kemp, thank you for your help with the immunohistochemistry experiments. Thank you to the Shoichet lab for letting me use their equipment for the completion of my work. Especially, I would like to thank Jackie Obermeyer and Ying Fang Cheng, who helped me carry out the GDNF bioactivity assays. I would like to thank my friends, new and old, for always reminding me to live life to the fullest. Anup, thank you for your companionship and friendship during this endeavour. Lastly, I would like to thank my family. Mom and dad: you have always given me your full support in everything I do. Most importantly, you ve taught me to not only strive for success, but also to enjoy the process. Thank you for leading by example. iv

5 Table of Contents Abstract... ii Acknowledgements... iii Table of Contents... v List of Figures... viii List of Tables... x Chapter 1. Introduction Overview Peripheral nerve anatomy and function Peripheral nerve injuries Neural response to injury Acute response: Wallerian degeneration Chronic nerve injury: Effects of chronic axotomy and denervation Surgical Techniques for Nerve Repair Nerve transfer End-to-side nerve coaptations Side-to-side nerve bridges Growth Factor Support Glial cell line-derived neurotrophic factor (GDNF) GDNF delivery systems Nerve guidance conduits (NGCs) Fibrin glue and affinity-based delivery systems (ABTS) Poly(lactic co-glycolic acid) (PLGA) microspheres Outcome Measures Axon counts Retrograde labeling Muscle force and mass Electrophysiology Motor unit number estimation Summary and Research Goals v

6 1.9.1 Hypothesis Objectives Chapter 2. Long-Term Delivery Experiments Abstract Introduction Methods Experimental design Preparation of GDNF microspheres and delivery system Surgical procedures Force analysis and motor unit number estimation (MUNE) Retrograde labeling Histomorphometry Statistical analysis Results Force analysis, MUNE, and wet muscle weights Retrograde labeling Histomorphometry Discussion Chapter 3. Short-Term Delivery Experiments Abstract Methods Experimental Design Preparation of microsphere delivery system GDNF microspheres characterization In vitro GDNF release DRG neurite outgrowth assay for GDNF bioactivity Surgical procedure Electrophysiology Retrograde labeling Histomorphometry Immunohistochemistry Statistical analysis Results vi

7 3.3.1 Microsphere characterization DRG bioactivity assay Functional outcome measures Retrograde labeling Histomorphometry Immunohistochemistry analysis Discussion In vitro delivery system characterization In vivo experiments Chapter 4. Summary and Conclusions Chapter 5. References vii

8 List of Figures Figure 1-1: Peripheral nerve connective tissue anatomy... 4 Figure 1-2: Myelinated axon anatomy... 5 Figure 1-3: Changes in cell membrane potential during an action potential... 6 Figure 1-4: Chronic axotomy and chronic denervation Figure 1-5: Techniques of microsurgical repair of injured nerves Figure 1-6: End-to-side nerve repair sprouting mechanisms Figure 1-7: GDNF receptor system for RET activation Figure 1-8: Fabrication of PLGA microspheres using a double emulsion process Figure 1-9: Twitch and tetanic isometric contractions Figure 2-1: Schematic of experimental design Figure 2-2: Timeline of experimental treatments Figure 2-3: Cumulative release of GDNF from various microsphere formulations in vitro.. 48 Figure 2-4: Surgical pictures for groups 1 and 3 no distal stump protection Figure 2-5: Surgery pictures for groups 2 and 4 distal stump protection Figure 2-6: Recording of muscle force and motor unit forces in vivo for the determination of reinnervated motor units Figure 2-7: Retrograde labeling surgery and nerve collection for histomorphometry Figure 2-8: EDL muscle force measurements, MUNE, and absolute muscle weights 5 months following delayed nerve repair Figure 2-9: TA muscle force measurements, MUNE, and absolute muscle weights 5 months following delayed nerve repair Figure 2-10: Gastrocnemius isometric twitch force measurements and absolute muscle weights 5 months following delayed nerve repair Figure 2-11: Number of retrograde labeled CP motoneurons in the spinal cord and sensory neurons in the dorsal root ganglia counted 5 months after immediate or 4 month delayed nerve repair Figure 2-12: Number of retrograde labeled CP motoneurons counted in the ventral horn of spinal cords 2 months after 2 month delayed nerve repair, with or without empty MS treatment viii

9 Figure 2-13: Representative sections of axons growing in the distal CP stump 5 months after nerve repair Figure 2-14: Histomorphometric analysis of CP nerves 5 months after nerve repair Figure 3-1: Timeline of experimental treatments Figure 3-2: Schematic of experimental design Figure 3-3: Recording of CMAP and MUNE in vivo Figure 3-4: Retrograde labeling procedure Figure 3-5: GDNF in vitro release profile Figure 3-6: Morphology of neurite outgrowth from DRGs Figure 3-7: Effect of GDNF on DRG neurite outgrowth length Figure 3-8: DRG neurite outgrowth images Figure 3-9: Functional outcome measures recorded from the tibialis anterior (TA) muscle 2 months following 2 months delayed nerve repair Figure 3-10: CMAP amplitude and templates for MUNE calculations Figure 3-11: Number of retrograde labeled motoneurons counted in the spinal cords 2 months after immediate or 2 month delayed nerve repair Figure 3-12: Pictures of retrograde labeled motoneurons as observed under fluorescent microscopy Figure 3-13: Representative sections of axon morphology from distal CP nerve cross sections collected 2 months following nerve repair Figure 3-14: Histomorphometry analysis of CP nerves 2 months after immediate or 2 month delayed nerve repair Figure 3-15: Number of axonal profiles in the CP nerve ix

10 List of Tables Table 1-1: Seddon s classification system for nerve injuries (Seddon 1943)... 9 Table 1-2: Sunderland's classification system for nerve injuries (Sunderland, 1978) (Sunderland 1951) Table 2-1 Control and experimental groups Table 3-1 Control and experimental groups x

11 Chapter 1. Introduction 1.1 Overview This work aimed to elucidate the protective effects of the side-to-side bridge surgical technique when coupled with the delivery of glial cell-line derived neurotrophic factor (GDNF) for the improvement of peripheral nerve regeneration. It was previously shown that both chronic denervation and chronic axotomy present independent major barriers for axonal regeneration (Fu & Gordon 1995b; Fu & Gordon 1995a). Chronic denervation occurs when nerves lose continuity with their neuronal cell bodies; chronic axotomy occurs when neurons are not connected to their target end-organs for prolonged periods of time (refer to section 1.4.2: Chronic nerve injury). The harmful effects of chronic denervation can be addressed by inserting nerve grafts connecting the side of an intact donor nerve to the side of a denervated recipient nerve stump (a side-to-side bridge ). This technique was previously implemented by Ladak and colleagues and was found to increase the number of motoneurons that regenerated their axons after a period of chronic denervation (Ladak et al. 2011). Additionally, the negative effects of chronic axotomy can be reversed with long-term continuous treatments of GDNF (Boyd & Gordon 2003a; Wood, Gordon, et al. 2012; Wood, Kim, et al. 2012; Wood et al. 2013). Wood et al. demonstrated that GDNF delivered at the 1

12 2 site of repair improved nerve regeneration and functional outcomes following delayed repair (Wood, Gordon, et al. 2012; Wood et al. 2013). The GDNF delivery system in that case consisted of GDNF encapsulated in poly(lactic co-glycolic acid) (PLGA) microspheres, placed within a fibrin gel for controlled delivery. Although both of these treatments (nerve bridging and GDNF microspheres) were previously evaluated individually, they had not yet been tested simultaneously to determine whether chronic axotomy and chronic denervation can be addressed concomitantly. The first study performed (presented in Chapter 2) evaluated the effects of the coupled treatment of side-to-side bridge protection with local GDNF delivery in long-term experiments. A rat model of chronic axotomy and denervation was created by surgical transection of the common peroneal (CP) nerve with and without side-to-side nerve graft protection of the denervated distal stump. After 4 months of chronic injury, the CP nerve was repaired and the GDNF delivery system was placed surrounding the repaired nerves in the experimental groups. The local GDNF delivery system released GDNF for a period of 6 weeks in vitro. After 5 months of nerve regeneration, the functional recovery of muscles innervated by the CP nerve was evaluated using force analysis and motor unit number estimation (MUNE). Motor and sensory neuron regeneration was assessed using retrograde labeling techniques and histology and morphometric evaluation of nerve sections. The second study performed (presented in Chapter 3) was introduced to elucidate whether results obtained in the first study were replicated with shorter time points and shorter span of GDNF delivery. These shorter time points were chosen because the long-term GDNF delivery system was found to be detrimental to regeneration. The same experimental groups

13 3 were utilized as those presented in Chapter 2 with some alterations. The release of GDNF from the delivery system was characterized in vitro for a period of 2 weeks, and the bioactivity of the GDNF released was assessed by neurite extension from rat embryonic dorsal root ganglionic neurons. The period of chronic axotomy and denervation was shortened from 4 months to 2 months, similar to the previous study evaluating the effects of GDNF on nerve regeneration after delayed repair (Wood, Gordon, et al. 2012; Wood et al. 2013). Two months after the repair surgery, reinnervation of the tibialis anterior muscle was evaluated using electrophysiological measures, including compound muscle action potential (CMAP) and motor unit number estimation (MUNE). Motoneuron regeneration was assessed using retrograde labeling techniques and histology and morphometric evaluation of nerve sections. The introduction chapter that follows provides an overview of peripheral nerve injury and regeneration. In the course of this chapter I present the current knowledge of the mechanisms which underlie nerve regeneration, and, explain, in part, the frequent disappointing outcomes of clinical nerve repair surgeries. I discuss the current experimental treatments for peripheral nerve injures, including growth factor delivery, and the rationale for the studies performed in this work. 1.2 Peripheral nerve anatomy and function Peripheral nerves consist of the axons and the Schwann cells of motor, sensory, and autonomic fibres. Motoneurons, located in the spinal cord or brainstem, transmit signals from the central nervous system to skeletal muscles in the periphery. Sensory neurons, located in

14 4 the dorsal root ganglia (DRG) or cranial ganglia, receive information from receptors in the periphery, including pain, thermal, and tactile receptors. The peripheral nerve trunk contains sensory and/or motor axons and is structured by connective tissue layers. The outer layer, the epinerium, surrounds the nerve trunk and serves as a protective layer binding several fascicles together. Fascicles are defined by the perineurium, and contain myelinated and unmyelinated axons that are surrounded by endoneurium (Figure 1-1). Figure 1-1: Peripheral nerve connective tissue anatomy A peripheral nerve trunk consists of the outer layer, the epineurium, binding fascicles together. Each fascicle is enveloped by the perineurium and contains the endoneurium which surrounds myelinated and unmyelinated axons. Figure adapted from (Biazar et al. 2010). Myelinated axons are surrounded by a series of Schwann cells within the basal lamina (Figure 1-2). Schwann cells produce myelin, an electrical insulator that wraps around

15 5 the axon and facilitates electrical conduction. Myelin increases impulse conduction velocity by decreasing the capacitance of the Schwann cell membranes, and this velocity is dependent on the fiber diameter and myelin thickness (Goldman & Albus 1968). The nodes of Ranvier are located between adjacent Schwann cells and are not myelinated, and have a high concentration of voltage gated sodium channels (Dugandgija-novakovic et al. 1995). They increase conduction velocity through the axon by allowing action potential propagation by saltation (Hodler et al. 1952). Figure 1-2: Myelinated axon anatomy The myelinated axon is surrounded by a series of Schwann cells. The figurative illustration of a motoneuron is adapted from (Waldram 2003). The cell body of a neuron (or soma) receives electrical signals from other neurons through its dendrites. The dendrites, which are cellular extensions branching from the soma, receive the electrochemical stimuli via synapses and transmit an electrical stimulus to the cell body. When the summation of synaptic inputs (excitatory and inhibitory) reach a triggering threshold, an action potential is initiated and conducted down the axon in an all-or-none response (Eyzaguirre & Kuffler 1955). The resting axon membrane potential is between -60 to -70 mv, with a net positive charge outside the axon and a net negative charge inside the axon. The inside of the cell has a higher concentration of potassium (K + ) and

16 6 organic ions, while the outside has a higher concentration of sodium (Na + ) and chloride (Cl - ) ions. An electric impulse is propagated when the voltage-gated Na + ion channels are opened and Na + moves down its concentration gradient into the axon and passes the threshold potential. The influx of Na + ions causes a brief reversal of charge, with the inside of the axon becoming positively charged and the outside negatively charged. This process is referred to as the rising phase of the membrane potential (Figure 1-3). The Na + channels are then inactivated, and passive K + channels allow K + to leak out of the axon returning the membrane potential to its resting state (falling phase). The brief reversal of charge between the inside and outside of the axon propagates and depolarizes the adjacent segment of membrane in a similar fashion. After the repolarization, the Na + /K + pump actively restores the ionic concentration to original levels, pumping K + into the cell and Na + out of the cell. Figure 1-3: Changes in cell membrane potential during an action potential Phases of the action potential in an axon segment. Figure adapted from (Sherwood & Kell 2009).

17 7 Given the length of nerve fibers, axoplasmic transport of mitochondria and materials from the cell body and along the axons is essential to provide materials for the axon membranes and the nerve terminals. Axoplasmic transport is categorized based on direction of transport: anterograde transport occurs from the cell body to the periphery and retrograde transport from the periphery to the cell body. Transport is also characterized as fast or slow, based on transport speed. Fast transport can cover up to 400 mm/day or 300 mm/day in the anterograde and retrograde direction, respectively, and slow transport can cover up to 4 mm/day (Brushart 2011). These processes allow for transportation of mitochondria, lysosomes, neurotransmitters, growth factors and their receptors, and materials necessary for the maintenance of the plasma membrane and cytoskeletal structures, including lipids, glycoproteins, actin, and microtubules (reviewed in Grafstein & Forman 1980). 1.3 Peripheral nerve injuries Injured neurons in the peripheral nervous system have the ability to regenerate their axons robustly in the presence of growth supportive Schwann cells (Hall 1986b; Hall 1986a; Gulati et al. 1995). However, the extent of nerve regeneration depends heavily on the severity of the injury. Several techniques have been developed over the years to address the poor outcomes observed after surgical repair but none were found to return sensory and motor function to pre-injury states (Kline 1990). This is particularly true for injuries requiring long regeneration times and distances, or of injuries presenting late and undergoing delayed repair (Hudson & Hunter 1977; Kline & Hudson 1995).

18 8 Nerve injuries caused by lacerations and other penetrating wounds are the most common, and include injuries resulting from gunshot wounds, automotive and sport accidents, and glass or knife wounds (Waldram 2003; Kline 1990). Due to the nature of the accidents, nerve injuries are missed sometimes or are diagnosed too late to achieve beneficial nerve regeneration through standard microsurgical repair of the divided nerve (Robinson 2004). Timing of the repair is crucial due to the deleterious effects of chronic axotomy and denervation (see below Section 1.4.2: Chronic nerve injury) that arise when nerve repair is delayed (Fu & Gordon 1995b; Fu & Gordon 1995a). After an injury, neurons are in a state of chronic axotomy after losing continuity with their target end-organs. At the same time, Schwann cells in the distal stump are in a state of chronic denervation, where they are no longer connected with neuronal cell bodies and viable axons. Even under circumstances where nerve repair is performed immediately after the nerve insult, lengthy periods of months and years are required for axons to regenerate long distances at a rate of 1 to 4 mm per day, limited by the rate of slow axonal transport (Grafstein 1995). Because injuries in older patients also result in suboptimal outcomes, age is an additional and important factor that that must be considered for functional outcome after peripheral nerve repair surgery (reviewed by Verdú et al. 2000). Much of the knowledge in peripheral nerve injury and repair was collected during periods of war, given that peripheral nerve injuries occurred in great numbers. Descriptions of nerve anatomy and nerve repairs, including grafting procedures, were developed during the First and Second World Wars (Sunderland 1951; Seddon 1943).

19 9 There are currently two major peripheral nerve injury classification systems. The first was developed by Seddon, in 1943, and the second by Sunderland, in It is useful to classify nerve injuries as often treatments are assigned based on the injury type. Seddon s system classifies nerve injuries into three categories, as presented in Table 1-1. Neurapraxia and axonotmesis do not cause disruption of the perineurium or epineurium and the injured nerves undergo spontaneous regeneration. On the other hand, neurotmesis involves damage to the whole nerve trunk, usually requiring surgical intervention for nerve regeneration to proceed (Seddon 1943). Table 1-1: Seddon s classification system for nerve injuries (Seddon 1943) Nerve Injury Type Characteristics of the nerve injury Neurapraxia Nerve compression No damage to the axon or perineurium No Wallerian degeneration Axonotmesis Damage to the axon No disruption of the endoneurial tube, perineurium, or epineurium Neurotmesis Transection of nerve trunk Repair of nerve required for recovery Sunderland developed a similar system but extended it to a five category classification (1-5). A first-degree injury is similar to Seddon s neurapraxia, a second-degree injury is axonotmesis, and third- to fifth- degree injuries represent different histologicallybased grades of injury as presented in Table 1-2 (Sunderland 1951; Waldram 2003; Wood et al. 2011).

20 10 Table 1-2: Sunderland's classification system for nerve injuries (Sunderland, 1978) (Sunderland 1951) Nerve Injury Type Characteristics of the nerve injury 1 Seddon s neurapraxia 2 Seddon s axonotmesis 3 Damage to the axon and endoneurial basal lamina tubes No disruption of the perineurium and epineurium 4 Damage to the axons, endoneurium, and perineurium No disruption of the epineurium 5 Transection of nerve trunk 1.4 Neural response to injury Acute response: Wallerian degeneration Following a peripheral nerve transection injury, the neuronal cell bodies undergo chromatolysis, and the proximal and distal nerve stumps undergo Wallerian degeneration. During chromatolysis, the nucleus swells and shifts from a central position to the periphery, and the Nissl bodies in the neuron disintegrate (Reviewed in Torvik 1976; Engh & Schofield 1972). During Wallerian degeneration, Schwann cells and macrophages phagocytose and remove myelin and axonal debris (Stoll et al. 1989). The proximal stump undergoes Wallerian degeneration for a small segment (one or more nodes of Ranvier) (Friede & Bischhausen 1980). In the distal nerve stump, Wallerian degeneration occurs in the whole stump. In the first few days after injury before macrophages reach the area Schwann cells are the main contributors to phagocytosis (Perry et al. 1995). Afterwards, when macrophages infiltrate the distal nerve stump from blood vessels, they become the main participants in phagocytosis and complete the process after days in a rat (Martini et al. 2008; Avellino

21 11 et al. 1995; Perry et al. 1995). Schwann cell proliferation occurs at the time of macrophage invasion (Perry et al. 1987). Macrophages are known to release Schwann cell mitogens, including transforming growth factor-β (TGF-β) and epidermal growth factor (EGF) (reviewed in Fu & Gordon 1997). The dividing Schwann cells line themselves as the Bands of Bungner to guide regenerating axons across the suture site and through the endoneurial tubes. If there is no surgical gap, or there is a minimal gap between the proximal and distal nerve stumps, regenerating axons travel through endoneurial tubes towards denervated targets. This is dependent on the availability of a guiding structure and a growth supportive environment through the distal nerve stump, as is demonstrated by the importance of basal lamina tubes (Ide et al. 1983). Otherwise abortive sprouting at the transection site can develop into a painful neuroma, an uncontrolled localized growth of immature fibers and connective tissue (Badalamente et al. 1985) Chronic nerve injury: Effects of chronic axotomy and denervation Axotomy occurs in the proximal stump when neurons from the injured nerve are no longer in contact with their target end-organs. Denervation occurs in the distal stump where the resident Schwann cells are no longer in contact with neuronal cell bodies and viable axons (Figure 1-4). Under circumstances in which the nerve gap is long or nerve repair is delayed, the neuronal cell bodies are in a state of chronic axotomy. At the same time, the distal nerve stump may be denervated for months and in many cases, years (Figure 1-4). The states of chronic axotomy of the neurons and chronic denervation of the Schwann cells have been associated with negative effects on axonal regeneration (Fu & Gordon 1995b; Fu & Gordon 1995a).

22 12 Figure 1-4: Chronic axotomy and chronic denervation Chronic axotomy occurs in the proximal stump when neurons are no longer in contact with their target end-organs. Chronic denervation occurs in the distal stump whereas Schwann cells are no longer connected with neuronal cell bodies and viable axons. It is known that following injury, neurons change their phenotype to a growth-permissive mode; however, the ability of neurons to regenerate their axons effectively is lost progressively with time. The effects of chronic axotomy on nerve regeneration, independent of chronic denervation, were evaluated by Fu and Gordon (1995b). In the study, a proximal nerve stump was axotomized for various periods of time and subsequently sutured to a freshly denervated distal stump. They found that the total number of reinnervated motor units decreased progressively with prolonged axotomy, with only 35% of motoneurons reinnervating muscle after only 3 months of chronic axotomy. Another study by Boyd & Gordon (2002) reported that the number of motoneurons that regenerated their axons was significantly lower after chronic axotomy than after immediate repair. A reason for this poor regeneration after axotomy may be due to the lack of contact with the distal stump that normally provides neurotrophic factors that act on the cell body. For instance,

23 13 delivery of growth factors to the regenerating axons has been found to reverse the negative effects of chronic axotomy (Boyd & Gordon 2003a; Boyd & Gordon 2002). With the progress of chronic denervation there is a substantial decrease in the number of axons that regenerate due to the progressive atrophy experienced by the Schwann cells in the distal stump (Röyttä & Salonen 1988). The cause of the progressive loss of Schwann cell support for regeneration has been studied extensively in the past (You et al. 1997; Li et al. 1997; Midha et al. 2005; Höke et al. 2002; Vuorinen et al. 1995; Fu & Gordon 1995a; Sulaiman & Gordon 2009). It is now known that the expression of several genes and receptors is altered following nerve injury. Schwann cells change from a myelinating phenotype to a growth-supportive phenotype (Reviewed by Fu and Gordon, 1997). Genes that are associated with myelination are downregulated, and growth-associated genes are upregulated, such as the genes for neural cell adhesion molecule (N-CAM), laminin, neurotrophins, growth factors, and their receptors. However, the upregulation response is transient and levels return to normal within relatively short periods of time, regardless of whether or not contact with axons is re-established. You et al. found that the early expression of the p75 neurotrophin receptor and cell adhesion proteins such as N-CAM was not maintained 2 months after injury, and hypothesized that Schwann cells provide the most suitable environment for axonal regeneration up to the first month after injury (You et al. 1997). Similarly, Li and colleagues found that the rate of Schwann cell proliferation and the expression of c-erbb receptors decreased progressively with time in denervated stumps, correlating with a decrease in axonal regeneration (Li et al. 1997). An important study for the basis of this thesis was performed by Hoke and colleagues, in which it was demonstrated that

24 14 the progressive decline in GDNF expression in denervated distal nerve stumps was associated with impaired regeneration (Höke et al. 2002). The importance of growth factors and GDNF specifically for nerve regeneration will be discussed in depth in the next section. The importance of Schwann cells and Schwann cell migration for distal stump axonal regeneration was demonstrated in several studies. Hall (1986) investigated axonal regeneration through acellular autografts where Schwann cell proliferation from the proximal stump was inhibited via injection of anti-mitotic agent mitomycin C. Even after 5 weeks of regeneration, 70-90% of basal lamina tubes remained empty of myelinating axons (Hall 1986b). Similarly, Enver and Hall (1994) evaluated axonal regeneration into muscle autografts in the absence of Schwann cells. Axonal regeneration was inhibited in autografts with no Schwann cells for up to 3 weeks, after which the Schwann recovered from the antimitotic effects of mitomycin C and migrated to support axonal regeneration (Enver & Hall 1994). Furthermore, the lack of axonal regeneration in acellular grafts longer than 40 mm was linked to the limited capacity of Schwann migration (Nadim et al. 1990; Hall 1986a; Enver & Hall 1994; Reviewed in Fu & Gordon 1997). Fu and Gordon (1995) performed a study in which they isolated the effects of chronic axotomy and chronic denervation. To evaluate the effects of chronic denervation alone they sutured a freshly transected proximal stump to a distal stump that had been denervated for varying periods of time. They found that with longer periods of denervation, axonal regeneration exponentially decreased. Furthermore, although axonal regeneration decreased with increased denervation periods, muscle end plates were still reinnervated successfully, indicating that muscles are still able to be innervated following denervation (Fu & Gordon

25 a). It is thus important to maintain a growth permissive environment in the distal stump to prevent the changes that occur after prolonged denervation periods, such as the collagenization and fragmentation of the endoneurial tubes (Sunderland & Bradley 1950). 1.5 Surgical Techniques for Nerve Repair The preferred surgical intervention following a nerve injury consists of the alignment and suture of the transected proximal and distal nerve stumps in an end-to-end fashion (Figure 1-5B). However, this technique is not an option when nerve gaps are too large since tension introduced in the nerve cables inhibits regeneration (Sunderland et al. 2004). When this is the case, surgeons resort to nerve grafting procedures in which a donor nerve is placed between the injured nerve stumps (Figure 1-5C) (Millesi et al. 1972). The grafts guide regenerating axons with their resident Schwann cells and basal lamina tubes. Although nerve grafting has been used as a gold standard for nerve gap repair, it requires the use of a healthy donor nerve, leading to donor site morbidity and the need for multiple surgical sites (Deumens et al. 2010). Additionally, the distal end of the graft can also be affected by the negative effects of chronic denervation, especially if the distance for regeneration is large (Koller et al. 1997). Nerve guidance conduits (NGCs) were introduced to avoid the sacrifice of a donor nerve required by nerve grafting. However, NGCs are only effective in short-gap injuries which limits their use (reviewed by Deumens et al., 2010). Despite the advancements in microsurgical techniques in these standard nerve repair procedures, the recovery of nerve function remains poor. Alternative techniques are thus being pursued, especially those targeting the negative effects of chronic denervation.

26 16 Figure 1-5: Techniques of microsurgical repair of injured nerves Schematic of (A) intact nerve, (B) end-to-end microsurgical nerve repair, (C) end-to-end nerve grafts placed between the proximal and distal ends of a transacted nerve, (D) nerve transfer of a freshly transected nerve to the distal stump of the injured nerve (E) end-to-side connection of the injured distal stump to an intact nerve, and (F) side-to-side bridge repair in which nerve grafts are connected from the side of an intact donor nerve to the distal stump of an injured nerve Nerve transfer The nerve transfer technique consists of the coaptation of the proximal stump of a freshly transected donor nerve to the injured distal nerve stump, in an end-to-end fashion (Figure 1-5D). Donor axons regenerate into the injured nerve denervated stump and serve as protection until the injured proximal stump is available for resuture. Bain and colleagues (2001) used sensory transfer nerve protection of a distal nerve stump to improve reinnervation after delayed nerve repair. They found significantly less muscle atrophy and increased compound muscle action potential (CMAP) in groups that had the protection (Bain et al. 2001). This positive effect was attributed to the neurotrophic effect that sensory axons growing in the distal stump had on the muscle fibers. Midha et al. (2005) evaluated the effects of sensory and motor nerve transfer protection of a denervated stump. They found

27 17 that denervated stumps that had been protected with a motor branch resulted in significantly more motoneurons regenerating their axons than motoneurons regenerating axons within pathways that were protected with a sensory nerve (Midha et al. 2005). This study was consistent with the study performed by Brenner et al. that reported that motor nerve regeneration was enhanced to a greater degree when the inserted grafts were motor rather than sensory grafts (Brenner et al. 2006). Whilst the technique of sacrificing either a motor or sensory nerve to protect a chronically denervated distal nerve stump is useful in research purposes, it is often not feasible for clinical use, involving the total sacrifice of a donor nerve. Specifically, motor nerves are usually not available for sacrifice due to their importance in their current functions to directly or indirectly control the muscles they innervate End-to-side nerve coaptations End-to-side (ETS) coaptation is a surgical technique performed after nerve injury that consists of the suture of the injured distal stump to the side of an intact nerve (Figure 5-1E). This technique was introduced as an alternative surgical method in injuries requiring long regeneration distances: the distal nerve is protected in an ETS manner until the proximal stump is close enough for resuture with the distal stump. However, there remain many controversies about its efficacy in clinical practice (Dvali & Myckatyn 2008). In this technique axons grow from the intact nerve into the denervated stump, thereby providing protection against denervation effects (reviewed extensively by Lykissas 2011; Tos et al. 2009; Bontioti & Dahlin 2009). The proposed mechanisms by which axons grow into the injured stump include collateral sprouting and terminal regenerating sprouting. Collateral

28 18 sprouting occurs when a healthy axon in the intact nerve sprouts an extra branch that grows into the injured nerve, without sacrificing its original innervation (Figure 1-6A). Terminal regenerating sprouting occurs when an injured donor axon grows into the protected end-to-side injured stump but sacrifices its original innervation for the new pathway (Figure 1-6B). Retrograde labeling of the donor and the recipient nerves after ETS repair have shown double-labeling of both motor and sensory neurons indicating the existence of collateral sprouting (Bontioti et al. 2005). Another mechanism that is thought to induce sprouting is the interneuronal signalling that occurs between injured and non-injured neurons in the central nervous system. If the injured neurons transfer their growth promoting signals to the intact neurons, sprouting may be induced in the intact nerve (Bajrović et al. 2002). Terminal regenerating sprouting requires the sacrifice of connections from the donor nerve, and is dependent on the extent of the donor nerve injury (Hayashi et al. 2008). Potential injuries induced in the donor nerve include epineurial and perineurial windows, and partial nerve neurectomy. Hayashi et al. (2008) found that more axons sprout into an ETS distal stump when there is injury or compression to the donor nerve. Furthermore, in the case of an atraumatic injury, only sensory fibers sprout into the recipient stump although this sprouting is insufficient for clinical relevance (Hayashi et al. 2008). Although the ETS technique is promising, it requires a secondary surgery for the reconnection of the proximal and distal stumps. Furthermore, if the distal stump lesion is found very proximally, the distal portion of the distal stump is still under the stress of chronic denervation during the regeneration period.

29 19 Figure 1-6: End-to-side nerve repair sprouting mechanisms Collateral sprouting of a healthy axon (A), and terminal regenerating sprouting of an injured axon (B). Figure adapted from (Bontioti & Dahlin 2009a) Side-to-side nerve bridges The side-to-side nerve bridge protection technique consists of nerve grafts ( bridges ) placed between the sides of an intact donor nerve and the sides of a denervated recipient stump (Figure 5-1E). In this technique, axons grow from the injured nerve through the bridges into the denervated stump, in a manner similar to the sprouting that occurs during ETS repair. Ladak and colleagues previously found that this technique protected against the negative effects of chronic denervation in a rat model (Ladak et al. 2011). They found that the number of motoneurons that regenerated their axons significantly increased after protection of the denervated distal stump with side-to-side nerve grafts. As discussed in the previous section (1.4.3 Chronic nerve injury), Schwann cells residing in the distal stump are important due to the trophic support that they provide for nerve regeneration. The donor axons growing through the bridges are thought to provide axon-schwann cell interactions that keep Schwann cells from becoming quiescent or atrophic. It has also been shown that

30 20 after injury Schwann cells migrate along with their regenerating axons from the proximal stump for a maximum distance of 40 mm (Nadim et al. 1990; Hall 1986a; Enver & Hall 1994). For instance, a study by Hall (1986) found that migration of Schwann cells precedes axonal regeneration into acellular grafts. Therefore, co-migration of Schwann cells with donor axons growing through the grafts into the denervated stump might serve as a further protective mechanism. The side-to-side repair technique is attractive due to its ability to protect denervated stumps over long distances as opposed to only one end as is the case with ETS repair. Multiple grafts can be placed throughout the length of autografts or distal stumps, providing maximum protection. Furthermore, the protection is enhanced given that axons grow both ways (proximally and distally) into the denervated stump (Gordon et al., unpublished data). If applied in a clinical setting, this repair technique requires only one surgery since the grafts can be applied at the same time of nerve repair. 1.6 Growth Factor Support Growth factors play a main role in the development of the nervous system and the nerve regeneration process after injury. The main candidates for use in the treatment of nerve injuries include the nerve growth factor (NGF), ciliary neurotrophic factor (CNTF), brain-derived neurotrophic factor (BDNF), neurotrophin-3 (NT-3), neurotrophin-4/5 (NT-4/5), and glial cell-line derived neurotrophic factor (GDNF) (reviewed by Boyd and Gordon, 2003a). It has been shown that these factors are crucial for motoneuron survival during development (Klein et al. 1993; Henderson et al. 1994; Yan et al. 1995). For example, inactivation of the receptors for BDNF and NT-4/5 in neonatal mice results in 35% loss of

31 21 spinal motoneurons and up to 70% loss of facial motoneurons (Klein et al. 1993). These numbers confirm the importance of BDNF and NT-4/5 during development but also indicate that motoneurons also respond to other sources of survival-inducing molecules. GDNF is one of these sources of survival, and was first isolated and identified as a growth factor promoting the survival of dopaminergic neurons that degenerate in Parkinson s disease (Lin et al. 1993). Since then it has been widely studied for its effect on motoneuron survival and nerve regeneration. For instance, GDNF was shown to be more potent than recombinant rat BDNF and CNTF for motoneuron survival in neonatal rats (Henderson et al. 1994; Yan et al. 1995). BDNF has shown beneficial and inhibitory effects on nerve regeneration, depending on the dosage (Boyd & Gordon 2002; Boyd & Gordon 2003a). When high doses of BDNF are administered, the binding of BDNF to the low-affinity p75 receptors mediates inhibitory effects. This was demonstrated by Boyd and Gordon (2002), where they reversed the inhibitory effects of the high BDNF doses by preventing the binding of BDNF and the p75 receptor. Unlike BDNF, GDNF does not bind to the p75 receptor, and thus does not show an inhibitory effect in the same manner (Boyd & Gordon 2003a). Although GDNF is potent for motoneuron survival, it can only exert its effects on a subset of sensory and autonomic neurons. This subset consists of the nociceptive neurons that are able to bind to the isolectin B4 and express the receptors for GDNF, GFRα and RET (Takaku et al. 2013). Growth factors have complex roles in peripheral nerve regeneration and are not yet fully understood. GDNF was chosen for use in this study due to its potency in motoneuron survival and regeneration and its extensive characterization in the past. The role of GDNF in the regeneration process will be discussed in more detail in the following section.

32 Glial cell line-derived neurotrophic factor (GDNF) GDNF belongs to the transforming growth factor-β (TGF-β) superfamily and is a protein of 134 amino acids, containing 7 cysteine residues that are characteristic of the TGF-β family. GDNF is N-glycosylated at 2 amino acid residues and possesses a cysteine knot that is also characteristic of the TGF-β proteins (Eigenbrot & Gerber 1997). The cysteine knot, formed by three cysteines, permits homodimerization of GDNF, conferring biological activity (Saarma & Sariola 1999). Downstream signalling of GDNF occurs via various signalling pathways, most of which require the binding of GDNF to the co-receptors GFRα1 (GDNF family receptor α1) and the tyrosine kinase RET (Figure 1-7). Signalling can also occur without the binding to RET: binding of GDNF to GFRα1 can activate the sarcoma kinase SRC promoting cell survival, and binding of GDNF to N-CAM can activate the SRC family kinase FYN, promoting neurite outgrowth (extensively reviewed by Saarma and Sariola, 2003 and Brushart, 2011).

33 23 Without RET Figure 1-7: GDNF receptor system for RET activation Dimeric GDNF (A) binds to the GDNF family receptor α1 (GFRα1) (B) and brings together two tyrosine kinase RET molecules (C). The co-receptors are joined and initiate autophosphorylation and downstream signalling. Signalling without RET also takes place. Figure adapted from (Sariola & Saarma 2003).

34 24 GDNF has been shown to promote cell survival, cell proliferation, and neurite outgrowth in vitro and has been shown to be a potent growth factor for motoneurons in vivo (Saarma & Sariola 1999; Henderson et al. 1994; Yan et al. 1995; Takaku et al. 2013; Trupp et al. 1995). For example, a study performed by Henderson and colleagues showed that after a neonatal facial nerve injury and chronic axotomy, treatment with GDNF at the proximal stump retained 92% of motoneurons, whereas treatment with vehicle retained only 29% of their facial motoneurons (Henderson et al. 1994). Furthermore, treatment with GDNF prevented the axotomy-induced shrinkage of motoneuron soma observed with treatment with BDNF and NT-4/5 (Yan et al. 1995; Henderson et al. 1994). These studies encouraged further research into the role of GDNF in the peripheral nerve regeneration process. After an injury, denervated Schwann cells in the distal stump increase the expression of GDNF and GFRα1, while the expression of RET remains constant (Höke et al. 2002). At the same time, axotomized motoneurons in the spinal cord increase the expression of the GFRα1 and RET receptors (Boyd & Gordon 2003b). Application of GDNF at the nerve injury site in adult rats has shown that GDNF is retrogradely transported to the spinal cord and prevents the decrease of acetylcholine transferase (AChT) levels normally observed after axotomy (Yan et al. 1995). It is thus assumed that following axotomy the regenerating axonal growth cones that express the RET coreceptor take up the GFRα1 and GDNF expressed by the Schwann cells and thus GDNF is able to exert its effects. A more recent study demonstrated that impaired nerve regeneration after chronic injury was correlated to the downregulation of the endogenous GDNF expression (Höke et al. 2002). Studies evaluating treatment of nerve injuries undergoing immediate nerve repair found no enhanced recovery

35 25 with GDNF treatment (Jubran & Widenfalk 2003; Boyd & Gordon 2003a). It is thought that the endogenous upregulation of GDNF that is observed at the early stages of nerve injury is sufficient to promote nerve regeneration and thus exogenous GDNF delivery has no effect at this stage. However, after chronic axotomy, GDNF is able to increase axonal regeneration and reverse the negative effects of chronic axotomy. There have been various studies replicating these results in various models, with GDNF improvement of nerve regeneration following chronic nerve injury (Barras et al. 2009; Barras et al. 2002; Boyd & Gordon 2003a; Wood, Gordon, et al. 2012; Wood, Kim, et al. 2012). It is important to note that multiple studies found that GDNF delivery did not have any beneficial effects if applied immediately after nerve repair (Barras et al. 2009; Eggers et al. 2008; Tannemaat, Eggers, Hendriks, de Ruiter, et al. 2008; Eggers et al. 2013; Boyd & Gordon 2003a; de Boer et al. 2012). It is possible that immediately after injury the transient upregulation of endogenous trophic support may reach saturating levels, which may explain, in part, the lack of a beneficial effects obtained with exogenous GDNF support during this time (Boyd & Gordon 2003a; de Boer et al. 2012). The focus of this thesis is to deliver GDNF at a time when endogenous GDNF expression is no longer upregulated in order to sustain axonal regeneration by targeting the negative effects of chronic axotomy. Different methods of GDNF delivery and the several challenges to be addressed will be discussed in the next section, as well as the choice for GDNF delivery used in this study.

36 GDNF delivery systems It was demonstrated that GDNF is an important factor that aids nerve regeneration following chronic injury (Höke et al. 2002). There are several methods used for the delivery of growth factors in research, including systemic and local delivery. Systemic administration of growth factors, although simple to implement, is not preferred since it can lead to systemic toxicity and unwanted side effects. For instance, in a clinical study involving injections of NGF for treatment of nerve neuropathies, NGF administration led to site pain and hyperalgesia in 67.2% of the patients compared to 11.6% of those receiving placebo (Schwartz et al. 2000). Another issue with systemic administration is that penetration of growth factors into nervous tissue from systemic delivery is poor and might be non-specific. Local administration addresses these concerns and is thus preferred. One of the requirements for effective local growth factor delivery is the ability to prevent the fast degradation that occurs in vivo. Several methods have been developed to address this issue by sequestering the growth factors and preventing the fast diffusion into the extracellular environment. Mini-osmotic pumps that deliver growth factors at a constant rate have been successfully used experimentally, although their permanent implantation can lead to inflammatory rejection (Aprili et al. 2009). Additionally, the eventual removal of the device is surgically problematic due to the scarring that occurs around the nerve and can risk neural damage. Other local delivery methods, including release from fibrin matrices, affinity-based delivery systems, nerve guidance conduits, and poly(lactic co-glycolic acid) (PLGA) microspheres will be discussed in the following section.

37 Nerve guidance conduits (NGCs) Nerve guidance conduits (NGCs) were initially designed for the guidance of axons through nerve gaps and isolation of the nerve regenerative environment (reviewed by Pfister et al. 2007). Since then, NGCs have been extensively studied and modified in an attempt to replace the need for autografts in nerve repair. They consist of hollow conduits filled with different materials, such as silicone, ethylene-vinyl acetate (EVA), poly(lactic-co-glycolic acid) (PLGA), collagen, chitosan, chitin, fibronectin, and fibrin. Several studies were succesful in controlling the delivery of growth factors from NGCs. For instance, microspheres incorporated in NGCs allowed for controlled the delivery of epidermal growth factor (EGF) and maintenance of EGF activity for a period of 14 days in vitro (Goraltchouk et al. 2006). Similarly, incorporation of GDNF releasing microspheres into NGCs allowed for sustained delivery of bioactive GDNF for >50 days in vitro and improved functional outcomes that were comparable to those of isografts in vivo (Kokai et al. 2010; Kokai et al. 2011). Wood and colleagues developed a fibrin-based NGC delivery system for GDNF in vitro that was then shown to improve peripheral nerve regeneration when compared to empty channels in vivo (Wood, Borschel, et al. 2009; Moore et al. 2010). However, the group treated with GDNF regenerated significantly lower number of fibers when compared to the isograft control group. Similarly, Fine and colleagues designed a NGC releasing GDNF across a 15 mm gap (Fine et al. 2002). The study found that axonal regeneration through the GDNF filled conduit was significantly increased when compared to conduits filled with BSA. However, no comparison to regeneration through an autograft was

38 28 made, so it is unclear whether this technique is an improvement to current nerve repair practices. Despite the advances made in NGCs there are still limitations to the clinical use of NGCs due to their limited efficacy in long gaps and the failure to improve recovery when compared to biological iso- and autografts. Additionally, the lack of compatibility set it behind other drug delivery methods, since most conduits studied are not biodegradable which may pose a problem for clinical use. For instance, a study evaluating the effects of GDNF delivery from NGCs found that half of the channels implanted became disconnected due to scratching (Barras et al. 2002). Many NGCs are still in the development phase and have only been tested in vitro and when NGCs were tested in vivo, most did not match the regenerative potential of biological isografts (Moore et al. 2010; Barras et al. 2002) Fibrin glue and affinity-based delivery systems (ABTS) Fibrin glue is a biocompatible and biodegradable material that has been tested for the controlled delivery of several growth factors, including GDNF, NGF, and NT-4/5 (Cheng et al. 1998; Yin et al. 2001; Jubran & Widenfalk 2003). It consists of a fibrinogen solution and a thrombin solution that is rich in calcium. When the two solutions are mixed they form fibrin, which is then crosslinked with factor XIIIa and forms a gel. Fibrin glue is an attractive biomaterial since it slows down the release of growth factors while also gluing tissues together, aiding in the repair of nerves (Spicer & Mikos 2010). A study by Cheng and colleagues measured the delivery of GDNF from fibrin glue and found that it was indeed released over prolonged periods of time (Cheng et al. 1998). The release of the GDNF from

39 29 fibrin is mainly governed by the disintegration of the fibrin network. The fibrinolytic system, including plasmin, is responsible for the lysis of the exposed fibrin network. Lysis occurs in several steps, initiated by the accumulation of plasminogen on the surface of the delivery system and followed by the fragmentation and subsequent collapse and dissolution of the fibrin network. This leads to the delivery of GDNF as the fibrin is gradually lysed. Despite the promising results, GDNF was released at a fast rate, with a half-life of around 3 days (Cheng et al. 1998). This fast GDNF release was attributed to the degradation of the fibrin network and weak binding of fibrin with GDNF. Fibrin glue is an attractive biomaterial since many of its properties can be altered by the concentration of its components. For example, pore size and time required for fibrin gelling can be controlled by the thrombin concentration (Spicer & Mikos 2010). Affinity-based delivery systems (ABTS) were developed to enhance the fibrin glue drug delivery properties. Wood and colleagues developed a fibrin gel with increased affinity for GDNF. They did this by crosslinking a bidomain peptide containing a transglutamine and a heparin-binding domain into the fibrin network (Wood, Borschel, et al. 2009). The addition of heparin changed the release kinetics of the delivery system, and resulted in retention of GDNF for longer periods. This ABTS was then introduced in a nerve guidance conduit model (NGC) to test the in vivo effects of GDNF on axonal regeneration and was found to improve results after sciatic nerve injury (Wood, Moore, et al. 2009; Moore et al. 2010).

40 Poly(lactic co-glycolic acid) (PLGA) microspheres Polymer microspheres have been previously used for the delivery of various medications, including synthetic drugs, proteins, and antibiotics (reviewed by Freiberg & Zhu 2004). Microspheres are of great utility in situations requiring controlled delivery of drugs, and can be used for oral or local administration. For the purpose of nerve injury and repair, local delivery is preferred, given that oral administration necessitates higher doses to ensure efficacy, which apart from being expensive can lead to systemic side-effects (Schwartz et al. 2000). Microspheres are an attractive option for the local delivery of growth factors for nerve regeneration. Microspheres made of polymers such as poly(lactic coglycolic acid) (PLGA) have been the focus of most studies due to their biodegradable and biocompatible properties, and their extensive use in local drug delivery (Shive & Anderson 1997). The polymer s biodegradation eliminates the need for removal of the delivery system after treatment completion, and its biocompatibility prevents an immune rejection by the body. PLGA is made of monomers that are natural in the body, lactic and glycolic acids, so it does not pose toxicity issues in the body. Furthermore, PLGA can stabilize the encapsulated proteins by delaying the fast-degradation that occurs in vivo (Garbayo et al. 2008). For instance, Wood and colleagues delivered GDNF for a period of 2 and 4 weeks using PLGA microspheres, and found this delivery to be beneficial to nerve regeneration after delayed nerve repair (Wood, Gordon, et al. 2012; Wood et al. 2013). Release from PLGA microspheres can be tailored by adjusting several chemical and physical characteristics. For instance, release rates are faster in microspheres with increased porosity, smaller diameters, and made with low molecular weight polymers (Ravivarapu et al. 2000; Bezemer et al. 2000;

41 31 Yang et al. 2000; Klose et al. 2006). Different fabrications techniques can be used to alter these properties, such as varying the fabrication temperature, stirring rates, concentrations, and chemical make-up, including the choice of polymers. Given that GDNF is a water soluble protein, the PLGA microspheres used in this study were made using a double emulsion process, also known as the solvent evaporation method (Freiberg & Zhu 2004; Wood, Gordon, et al. 2012; Garbayo et al. 2008). The process consists of two main steps (Figure 1-8). During the first emulsion (W/O), the water phase containing GDNF is dispersed in the oil phase containing the PLGA and organic solvents (dichloromethane (DCM) and acetone) (Figure 1-8A). The second emulsion (O/W) is formed from the dispersion of the contents of the first emulsion in an aqueous medium containing poly(vinyl alcohol) (PVA), which acts as a stabilizer (Figure 1-8B). The microspheres are formed as the organic solvents (DCM/acetone) evaporate and the polymer hardens.

42 32 A B Figure 1-8: Fabrication of PLGA microspheres using a double emulsion process Schematic of the first and second emulsions created during GDNF microsphere fabrication. The first emulsion (W/O) consists of water dispersed in an oil medium containing organic solvents. The second emulsion (O/W) consists of the first emulsion dispersed in an aqueous medium containing a stabilizer. Subsequently, evaporation of the organic solvents and hardening of the polymer leads to microsphere formation. The rate of drug release from microspheres depends on degradation and diffusion rates. Release via degradation occurs by the formation of pores in the polymer matrix as the polymer degrades, whereas release from diffusion occurs by the dissolution of the drug into the bulk solution led by concentration gradients through the aqueous channels in the polymer matrix. Most often microsphere release profiles display an initial burst that is representative of the release of drugs at the surface of the microspheres (reviewed by Freiberg & Zhu 2004; Yeo & Park 2004). This burst release is followed by a more constant release, controlled by polymer degradation and diffusion. If high MW polymers are used, the release is usually initially slow, followed by an increased constant release rate (Makino et al. 2000). The initial

43 33 slow release is thus controlled by the diffusion, which is later followed by a faster release rate controlled by PLGA degradation. It is important to note that PLGA microspheres undergo degradation by bulk erosion rather than surface erosion, since water rapidly imbibes into the microparticles (Klose et al. 2006). Apart from GDNF, other growth factors that have been encapsulated in PLGA microspheres include NGF and BDNF (Mittal et al. 1994; Aubert-Pouëssel et al. 2004; Garbayo et al. 2008; Wood, Kim, et al. 2012; Krewson et al. 1996). Garbayo et al. and Pouessel et al. successfully encapsulated bioactive GDNF in PLGA microspheres for the sustained delivery over 40 days in vitro (Garbayo et al. 2008; Aubert-Pouëssel et al. 2004). Many of the in vivo studies applying GDNF microspheres have been focused on delivery to the spinal cord or brain for the treatment after stroke injury or Parkinson s disease. For the purpose of nerve regeneration, the use of GDNF microspheres is relatively new. Microspheres have been used in isolation or in addition to already developed growth factor delivery methods, such as NGCs and fibrin delivery systems (Baumann et al. 2009; Stanwick et al. 2012; Wood, Kim, et al. 2012; de Boer et al. 2012). Microspheres releasing GDNF incorporated into NGCs were used by de Boer et al. after nerve injury and repair (de Boer et al. 2012). Although they found an initial beneficial effect of the treatment, they did not show a long term improvement in regeneration in terms of electrophysiology, functional outcomes, and histological and morphometry measures. These findings were consistent with the studies performed previously by Boyd and Gordon, which demonstrated no beneficial effect on regeneration by GDNF application after immediate nerve repair (Boyd & Gordon 2003a). Similarly, Kokai et al. developed a NGC that incorporated GDNF microspheres and was implanted in vivo (Kokai et al. 2010). GDNF delivery from microspheres increased

44 34 regeneration in terms of increased axonal numbers and restoration of muscle force when compared to empty conduits in a rat sciatic nerve gap, indicating that GDNF can potentially be incorporated in current NGC treatments. However, wet muscle weights were not comparable from empty nerve guides. More recently, Wood et al. evaluated various GDNF microsphere formulations in vitro and in vivo for the application in delayed nerve repair (Wood, Gordon, et al. 2012; Wood, Kim, et al. 2012; Wood et al. 2013). GDNF releasing microspheres embedded in fibrin significantly increased the number of motoneurons that regenerated their axons and improved muscle force following delayed repair. These studies concluded that a shorter term delivery of GDNF for a period of 2 weeks following delayed repair is preferential compared to longer treatment (4 weeks) for the improvement of muscle mass and axonal regeneration. Together, these studies demonstrate the efficacy of GDNF microspheres for enhanced nerve regeneration. Furthermore, these biodegradable systems are ideal for translation into clinical use due to their ease of application at the time of nerve repair surgeries and their use of FDA-approved materials. 1.8 Outcome Measures The following section will present the outcome measures that were used in this thesis work to evaluate the extent of nerve regeneration in vivo. Several measures were included to obtain a comprehensive evaluation of the regeneration process. All these techniques were reviewed in depth by Wood et al. (2011) and Brushart (2011).

45 Axon counts Myelinated axon numbers and axon morphology can be evaluated by examination of thin cross-sections of nerve tissue. The nerve tissue is osmicated and counterstained with toluidine blue dye, which stains the myelin blue. Images are taken at 1,000x magnification and are then analyzed with software to determine the total axon count, axon diameter, and fiber diameter. Secondary measures, such as the fiber area, axon area, myelin thickness, and g-ratio can be calculated afterwards. The measure of axon counts is widely used and serves as a good evaluation of regeneration as long as the procedure is carried out in the same manner across all groups being compared. However, a limitation of this technique is that it does not distinguish axons that originated from the same parent neuron. This is particularly an issue when experimental conditions increase axonal sprouting but do not increase the number of neurons that regenerated their axons. For instance, studies found that delivery of GDNF following nerve repair promoted sprouting and resulted in higher myelinated axon counts, however it did not result in an increase in the number of motoneurons that regenerated their axons (de Boer et al. 2012; Boyd & Gordon 2003a). This measure is thus better used as a complimentary technique to ensure that appropriate conclusions on the regeneration process are made Retrograde labeling Retrograde labeling is a technique used to count the number of neurons that regenerated their axons. Dyes are placed in the periphery and axons retrogradely transport the dye to the parent neuron. Common dyes used for this purpose include FluoroGold,

46 36 FluoroRuby, FluoroEmerald, and Fast Blue (Richmond et al. 1994). The dye can then be visualized in the spinal cord or dorsal root ganglia under fluorescence microscopy and the number of neurons can be counted. This labeling technique is powerful if performed in the same manner across all groups tested. Furthermore, the labeling of the parent neurons eliminates the confounding factor of axonal sprouting during regeneration. The technique used in our laboratory isolates the nerve and limits the exposure of the dye to the nerve of interest. This prevents the contamination of surrounding tissue that occurs with intramuscular or intradermal injections (Hayashi et al. 2007) Muscle force and mass The measure of isometric muscle force is performed at the end of experiments since it is an invasive procedure involving damage to the muscle tendons. In this procedure, the patella is immobilized and the tendons of the muscle innervated by the nerve of interest are tied to a strain gauge. The nerve is stimulated and the muscle length is adjusted for maximum twitch force production. The maximal twitch force that results from short duration stimuli to the nerve provides insight on the degree of muscle fiber activation in response to a single action potential (Figure 1-8A). Measure of the tetanic force involves the stimulation of the nerve at a high frequency. If the frequency of stimulation is not high enough, the twitch contractions will oscillate (Figure 1-8B). Once a threshold frequency is reached, muscle fibers have no time to relax after individual twitch contractions and the muscle reaches a peak force, staying constant at a plateau (Figure 1-8C). The magnitude of twitch and tetanic forces is dependent on the number of muscle fibers innervated by each motor unit. Reinnervation after injury can lead to the enlargement of motor units, with regenerating axon

47 37 innervating more than one muscle fiber. However, each regenerated axon only innervates up to 3-5-fold the number of motor fibers in the rat, so low muscle forces often translate to fewer axonal reinnervation (Fu & Gordon 1995a; Fu & Gordon 1995b; Wood et al. 2011). Muscle mass is also correlated to the number of motor fibers that are reinnervated, and is a good measure of muscle atrophy. Figure 1-9: Twitch and tetanic isometric contractions Twitch force response from a short duration stimuli to the nerve (A). Stimulation of the nerve at a frequency below the threshold, leading to the oscillating twitch contractions (B). Stimulation of the nerve after the threshold frequency is reached: muscle fibers have no time to relax and the muscle reaches a peak force, staying constant at a plateau (C).

48 Electrophysiology Compound muscle action potentials (CMAP) are the summation of individual motor unit action potentials, and are obtained by electrically stimulating the nerve and recording the electromyography (EMG) response in the muscle in mv. The amplitude of the CMAP, or the area under the curve, are generally used to evaluate the extent of functional recovery (Archibald et al. 1991). CMAP is dependent on the number and size of the innervated motor units and the different axon conduction velocities. The more synchronized the axon velocities, the higher the CMAP (Madison et al. 1999) Motor unit number estimation Motor unit number estimation (MUNE) can be performed using force analysis or CMAP and EMG measurements. MUNE evaluation using force analysis is performed by first recording several isometric twitch responses over a range of stimulation amplitudes up to ~60% of the maximum twitch force (Major et al. 2007). Afterwards, twitch responses are selected and they are subtracted from the next largest response. The mean motor unit (MU) force is then calculated as the average of the motor unit response samples obtained from the subtractions. MUNE is the ratio of the maximum muscle twitch force and the mean MU force. In this method, less-than-linear summation of motor unit forces can occur, leading to lower or higher estimations, depending on muscle type (Drzymała-Celichowska et al. 2010). Furthermore, one of the limitations of the force analysis method is that it does not detect the smallest motor unit potentials, since that is dependent on the sensitivity of the strain gauge (Major et al. 2007)

49 39 The EMG method used to evaluate MUNE is performed by first measuring the maximum CMAP elicited by supramaximal stimulation of the nerve, representing the sum of the individual motor unit potentials. Afterwards, templates are recorded after gradual increases of the stimulus from threshold, causing distinct increments in EMG amplitudes. The absolute area under the CMAP curve of each template is calculated and subtracted from the next largest. The average motor unit size is then calculated as the average of the CMAP template subtractions. The estimation of the number of reinnervated motor units (MUNE) is obtained by the ratio of the maximum CMAP and the average size of a motor unit (Willand et al. 2011). This metric is a good evaluation that is independent of the size of the reinnervated fibers (Wood et al. 2011). 1.9 Summary and Research Goals Targeting the negative effects of chronic axotomy and denervation and maintaining an optimal nerve regenerating environment for extending axons are key to improving nerve regeneration after injury. The use of side-to-side bridges successfully targets chronic denervation in a rat model of delayed nerve repair, and long-term continuous treatments with exogenous sources of glial cell-line derived neurotrophic factor (GDNF) reverse the effects of chronic axotomy. The objective of this study is to evaluate the combined effects of the side-to-side bridges in conjunction with GDNF application in a rat model of chronically axotomized and denervated neurons. These studies will be important for the translation of these techniques to human surgery in cases where regenerative distances are long and side-to-side bridges may successfully protect chronically denervated nerve.

50 Hypothesis Combination therapy targeting chronic denervation with side-to-side bridges and chronic axotomy using locally applied GDNF-loaded microspheres will improve axonal regeneration and muscle functional recovery compared to either treatment alone Objectives 1. Evaluate functional recovery in denervated muscles by force analysis and motor unit number estimation (MUNE) in a rat model of chronic axotomy and denervation after local GDNF delivery, side-to-side bridge protection, or both. 2. Measure the amount of axonal regeneration using retrograde labeling and histomorphometry in a rat model of chronic axotomy and denervation after local GDNF delivery, side-to-side bridge protection, or both.

51 Chapter 2. Long-Term Delivery Experiments This chapter presents the results of the first experiment evaluating the effects of GDNF delivery and side-to-side nerve graft protection of denervated distal stumps for the improvement of regeneration after delayed nerve repair. The periods of chronic injury and subsequent regeneration were 4 and 5 months respectively, for consistency with previous studies using the side-to-side bridge technique (Ladak et al. 2011). Local delivery of GDNF was performed using poly(lactic-co-glycolic) acid (PLGA) microspheres embedded in fibrin for GDNF release for up to 6 weeks. Outcome measures evaluated were force analysis and motor unit number estimation (MUNE) of the reinnervated muscles as well retrograde labeling and histology and morphology analysis of the regenerated nerves. Based on the outcomes of this study a second set of experiments were performed, presented in Chapter Abstract Following nerve injury, chronic denervation of Schwann cells and chronic axotomy of neurons present major barriers to axonal regeneration and functional recovery. The objective of this study was to evaluate the extent of functional recovery and axonal regeneration in a rat model of delayed nerve repair after treatment with local GDNF delivery, side-to-side 41

52 42 bridge protection, or both. A delayed nerve repair model was created by transection of the common peroneal (CP) nerve and suture of the proximal and distal nerve stumps to innervated muscle. Nerve grafts obtained from the contralateral side were used as side-to-side nerve grafts in the experimental groups. After 4 months the CP nerves were repaired, and the GDNF delivery system was placed surrounding the repaired nerves in the experimental groups. After 5 months of nerve regeneration, functional recovery of the muscles was evaluated using force analysis and motor unit number estimation (MUNE). Additionally, the regenerated nerves were evaluated by retrograde labeling and histomorphometry analysis of nerve sections. Comparing the treatments with side-to-side nerve grafts with and without GDNF revealed no statistical difference in twitch force, tetanic force, and motor unit numbers. There was no significant difference between groups that did not receive distal stump protection, regardless of GDNF treatment, in any of these metrics. Furthermore, the presence of side-to-side nerve grafts, regardless of GDNF treatment, led to statistical improvement in twitch, tetanic force, and MUNE compared to groups without bridge protection of the denervated distal stumps. Retrograde labeling and histomorphometry measures revealed no significant differences between groups with and without side-to-side bridges. Furthermore, axonal regeneration was significantly impaired in groups that received GDNF treatment. Overall, the side-to-side bridging technique was effective in protecting against the negative effects of chronic denervation regardless of GDNF treatment based on force and MUNE analysis, but showed no improvement in the number of motoneurons that regenerated their axons. Local delivery of GDNF for a period of 6 weeks did not increase functional recovery and based on the numbers of motoneurons that regenerated their axons impaired axonal regeneration.

53 Introduction In contrast to injured neurons in the central nervous system, injured neurons in the peripheral nervous system have the ability to regenerate their axons robustly in the presence of growth-supportive Schwann cells (Enver & Hall 1994; Hall 1986b; Nadim et al. 1990). However, the effectiveness of this regeneration depends on several factors, including the extent of the injury and time of surgical repair (Kline & Hudson 1995; Hudson & Hunter 1977). Nerve injuries are sometimes missed or are diagnosed too late to achieve beneficial nerve regeneration through standard microsurgical techniques (Robinson 2004). Injuries that require nerve regeneration over long distances often lead to poor functional outcomes due to the long regeneration period that depends on the slow axoplasmic transport of 1 to 4 mm per day (Grafstein & Forman, 1980). For example, after injuries to the sciatic nerve or brachial plexus it can take months and even years for regenerating axons to reach their target organs (Kim et al. 2003). During the long regeneration process neurons are not connected to their target end-organs and are in a state of chronic axotomy. At the same time, Schwann cells in the distal stump are not connected to neuronal cell bodies through viable axons, and are in a state of chronic denervation. It was shown previously that both chronic axotomy and chronic denervation present independent major barriers for axonal regeneration (Fu & Gordon 1995b; Fu & Gordon 1995a). Höke et al. (2002) demonstrated that the progressive decline in GDNF expression in denervated distal nerve stumps is associated with impaired regeneration. Studies were then performed to elucidate whether the maintenance of elevated levels of GDNF during the regeneration period may prevent the progressive decline in regeneration potential. Indeed, the negative effects of chronic axotomy were reversed with

54 44 long-term continuous treatments of GDNF (Boyd & Gordon 2003a; Wood, Gordon, et al. 2012; Wood, Kim, et al. 2012; Wood et al. 2013). Additionally, the harmful effects of chronic denervation can be addressed by inserting nerve grafts connecting the side of an intact donor nerve to the side of a denervated recipient nerve stump (a side-to-side bridge ) (Ladak et al. 2011; Midha et al. 2005). This study aimed to elucidate whether combination therapy targeting chronic denervation with side-to-side bridges and chronic axotomy using locally applied GDNF-loaded microspheres enhanced nerve regeneration as compared to either treatment alone. Functional recovery of muscles was evaluated using force analysis and motor unit number estimation (MUNE) and axonal regeneration was evaluated using retrograde labeling and histomorphometry analysis of nerve sections. 2.3 Methods Experimental design Sixty-four female Sprague-Dawley rats ( g) were divided equally into 4 groups in which all rats undergo delayed nerve repair (refer to Figure 2-2 for timelines). Each group was evaluated by retrograde labeling and histomorphometry (n = 8) and by muscle force analysis and motor unit number estimation (MUNE) (n = 8), as shown in Table 2-1. In the first surgery, all rats (groups 1-4) underwent a common peroneal (CP) nerve transection. The proximal and distal nerve stumps were sutured to innervated muscle for a period of 4 months to prevent axon regeneration into the distal CP nerve stump (groups 1 and 3, Figure 2-1A). Additionally, nerve grafts obtained from the contralateral side

55 45 were used as autologous side-to-side nerve grafts connecting the injured CP nerve to the intact tibial (TIB) nerve in groups with the bridge protection (groups 2 and 4, Figure 2-1B). Following 4 months of chronic axotomy and denervation, a nerve repair surgery was performed on all the animals (Figure 2-1C-F). At this time the repaired CP nerves of groups 3 and 4 were surrounded by a GDNF delivery system (Figure 2-1E, F). Local delivery of GDNF was performed using poly(lactic-co-glycolic acid) (PLGA) microspheres embedded in fibrin allowing for non-invasive controlled delivery for up to 6 weeks. Five months after CP nerve repair, nerve regeneration and muscle reinnervation were evaluated using force analysis, motor unit number estimation (MUNE), retrograde labeling, and histomorphometric analysis. Table 2-1 Control and experimental groups Nerve regeneration was evaluated in experimental groups by retrograde labeling, histomorphometry analysis of nerve sections, and motor unit number estimation (MUNE). Delayed nerve repair Group Treatment Outcome measure n 1 No protection + Saline Retrograde labeling, histomorphometry 8 Force analysis, MUNE 8 2 Bridge protection + Saline Retrograde labeling, histomorphometry 8 Force analysis, MUNE 8 3 No protection + GDNF Retrograde labeling, histomorphometry 8 Force analysis, MUNE 8 4 Bridge protection + GDNF Retrograde labeling, histomorphometry 8 Force analysis, MUNE 8 Total 64 Immediate nerve repair Treatment Outcome measure n 5 None Retrograde labeling, histomorphometry 8 Total 8

56 Figure 2-1: Schematic of experimental design. All rats underwent a common peroneal (CP) nerve transection, in which the proximal and distal CP nerve stumps were sutured to innervated muscle for 4 months (A). Additionally, half of the rats received side-to-side bridge protection of the distal CP nerve stump (B). After 4 months, the CP nerve was repaired in all animals in groups 1-4 (C-F), and the CP nerves of animals in groups 3 and 4 were surrounded by the GDNF delivery system (E-F). 46

57 47 Figure 2-2: Timeline of experimental treatments During the first surgery, the CP nerves were transected and sutured to innervated muscle, with or without side-to-side bridge protection. After 4 months of chonic axotomy and denervation, the CP nerves were repaired and half the animals received no treatment (NT) or GDNF MS embedded in fibrin. Outcome measures were evaluated 5 months after the repair surgery Preparation of GDNF microspheres and delivery system Poly(lactic-co-glycolic acid) (PLGA) (50/50; inherent viscosity 75% , 25% dl/g) microspheres loaded with GDNF were prepared through a double emulsion process (Wood et al. 2013; Wood, Gordon, et al. 2012). The aqueous phase consisted of 100uL of ddh 2 O, 12.5 mg of heparin, and 500 µg GDNF. The solvent phase was comprised of 230 mg of PLGA dissolved in 1 ml solution of dichloromethane/acetone (75%/25%) and 12.5 mg of MgCO 3. The aqueous and solvent phases were emulsified under sonication (45 s; 3 mm probe; 30% amplitude) and then added to 25 ml of 2.5% polyvinyl alcohol (PVA)/10% NaCl solution and homogenized (60 s; 6,000 rpm). The resulting mixture was then poured into a 250 ml bath of 0.25% PVA/10% NaCl and stirred at 125 rpm for 3 h for microsphere hardening and evaporation of organic solvent. The microspheres were then washed with 1 L of ddh 2 O by centrifugation (500G, 5 min, at least 5 cycles). The resulting microspheres were pooled into a conical tube with 4 ml ddh 2 O, flash frozen in liquid nitrogen, and lyophilized for 3 days. Microspheres were stored at -20ºC until use for 3 months (encapsulation efficiency of 80 ± 4%).

58 48 The delivery system was constructed by mixing 80 µl of fibrin with 5 mg of microspheres (containing ~8 µg GDNF). Fibrin was formed by mixing 40 µl of fibrinogen ( mg/ml) with 40 µl thrombin (5 U/mL) (Tisseel, Baxter, Deerfield, IL). The time release of GDNF from the microspheres was determined by Matthew Wood previously (Wood, Gordon, et al. 2012). Microspheres were found to have a sustained release of GDNF projected to 6 weeks (Formulation 3, Figure 2-3). Figure 2-3: Cumulative release of GDNF from various microsphere formulations in vitro. The microspheres used for the experimental groups in this study were formulation GDNF MS (6 wk) (Wood, Gordon, et al. 2012) Surgical procedures All surgical procedures were approved by the Hospital for Sick Children s Laboratory Animal Services Committee and were done in an aseptic manner. Rats were anaesthetized with isoflurane and the surgical sites were shaved and cleaned using Betadine and isopropyl alcohol. The three major branches of the sciatic nerve the CP, sural, and TIB nerves were exposed through a gluteal-splitting incision and separated. The CP nerve was

59 49 transected approximately 5 mm distal to the point of trifurcation. The proximal and distal CP nerve stumps were then sutured to innervated muscle to prevent axon regeneration (Figure 2-4A). Additionally, in groups with distal stump protection, grafts serving as the side-to-side bridges were prepared from the CP nerve in the contralateral hindlimb (3 nerve grafts ~ 4 mm each) (Figure 2-5A). The grafts were secured with biodegradable fibrin glue (Tiseel, Baxter Healthcare) between epineural openings (windows) made in the intact donor TIB nerve and in the recipient denervated CP distal nerve stump. Following the surgery all animals were allowed to convalesce for a period of 4 months (Ladak et al. 2011). After 4 months of chronic axotomy and denervation, the transected CP nerve was exposed (Figures 2-4B; 2-5B). Approximately 1 mm of nerve was resected from each CP stump and repair was performed by coaptation of the proximal and distal CP nerve stumps using 10-0 nylon sutures (Figures 2-4E; 2-5D) (Wood, Kim, et al. 2012). Groups 1 and 3, which required GDNF treatment, were surrounded by the delivery system at the suture site (Figures 2-4E, F; 2-5E, F). The animals were allowed to convalesce for a period of 5 months after which regeneration was evaluated using different metrics.

60 50 A B C CP proximal CPdistal TIB D E F Figure 2-4: Surgical pictures for groups 1 and 3 no distal stump protection. The CP nerve was transected and sutured back to innervated muscle (A). After 4 months, the CP nerve was exposed (B), dissected (C, D), and repaired using 10-0 nylon sutures. Additionally, the CP nerves of animals in group 3 were surrounded by a fibrin GDNF delivery system at the suture site (E, F). A B C CP proximal CPdistal TIB D E F Figure 2-5: Surgery pictures for groups 2 and 4 distal stump protection. The CP nerve was transected and sutured back to innervated muscle, and three side-to-side bridges were placed between the donor TIB nerve and the recipient CP distal stump (A). After 4 months, the CP nerve was exposed (B), dissected (C, D), and repaired using 10-0 nylon sutures. Additionally, the CP nerves of animals in group 4 were surrounded by a fibrin GDNF delivery system at the suture site (E, F).

61 Force analysis and motor unit number estimation (MUNE) Five months after the nerve repair surgeries animals were anesthetised and tendons of some of the muscles innervated by the CP nerve the tibialis anterior (TA) and extensor digitorum longus (EDL) were tied to a force transducer. Isometric twitch and tetanic muscle contractions were recorded in response to stimulation of the CP nerve after determining the optimal muscle length and stimulating amplitude. Furthermore, motor units were counted using motor unit number estimation (MUNE) (Major et al. 2007). Briefly, several evoked twitch responses over a range of stimulation amplitudes of up to ~60% of the maximum twitch force were recorded (Major et al. 2007). Afterwards, twitch responses were selected and each was subtracted from the next largest evoked response. The mean motor unit (MU) force was then calculated as the average of the motor unit response samples obtained from the subtractions. The estimation of the number of motor units that were reinnervated was obtained by the ratio of the maximum recorded muscle twitch force and mean motor unit (MU) isometric forces (Figure 2-6). Following the force analysis and MUNE procedures the muscles were harvested and weighted to evaluate the extent of muscle atrophy in the different groups.

62 52 Figure 2-6: Recording of muscle force and motor unit forces in vivo for the determination of reinnervated motor units Muscles innervated by the CP nerve were tied to a force transducer to record isometric twitch and tetanic muscle contractions in response to stimulation of the CP nerve. The number of reinnervated motor units was obtained by the ratio of the maximum muscle force and average force of a motor unit (Major et al. 2007) Retrograde labeling Five months after the repair surgery, the CP nerve was exposed and transected approximately 12 mm distal to the suture site. A 4% FluoroRuby dye solution was applied at the tip of the transected CP nerve in a Vaseline well for 1 h. Additionally, the side-to-side nerve grafts of groups 2 and 4 were cut to prevent backlabeling of the TIB motoneurons that grow through the side-to-side branching (Figure 2-7). Animals were allowed to recover for 7 days, to allow for retrograde transport of the dye to the cell bodies before animal perfusion and tissue harvest. ANIMAL PERFUSION Seven days after the retrograde surgery, rats were anaesthetized by injection of sodium pentobarbital (Euthanyl) for transcardial perfusion. A detailed methodology for the

63 53 procedure was presented by Gage et al. (2012) previously. Briefly, an incision was made in the diaphragm using blunt scissors and the incision was continued along both sides of the ribcages to expose the lungs and heart. The tip of the sternum was clamped with a hemostat and placed over the rat s head to provide a clear view of the heart. An 18G blunt needle was passed from the apex of the heart through the aorta, and clamped in place with another hemostat. The right atrium was then cut, and 180 ml of 0.9% saline was slowly injected into the aorta with a syringe, followed by 180 ml of cold paraformaldehyde (4% paraformaldehyde in 0.1 M phosphate buffer). After finishing the perfusion, the lumbar enlargement of the spinal cords, and L4 and L5 dorsal root ganglia were harvested and kept in 4% paraformaldehyde solution overnight. The tissues were then moved to a cryoprotective 30% sucrose solution and left overnight, after which the tissues were embedded and frozen with liquid nitrogen and stored at -80ºC. MOTOR AND SENSORY NEURON COUNTS Sagittal sections of the spinal cords and DRGs were cryosectioned at 50 µm and 20 µm, respectively. Motor and sensory neurons that regenerated their axons into the CP nerve stump were fluorescently labelled, and were counted at nm excitation with a 10x objective (100x overall magnification, Leica). For spinal cord motoneuron counts, all motoneurons in the sections were counted and the Abercrombie correction factor was applied to correct for repetitive counting of the same motoneurons in multiple sections (Abercrombie 1946). Similarly, for DRG sensory neurons counts all neurons in every fifth section were counted as performed previously (Wood et al. 2013; Barras et al. 2002; Barras et al. 2009; Bennett et al. 2000).

64 54 FluoroRuby Figure 2-7: Retrograde labeling surgery and nerve collection for histomorphometry A 4% FluoroRuby dye solution was applied at the tip of the transected CP nerve in a Vaseline well for 1 h. In groups with side-to-side bridge protection of the distal stump, the side-to-side bridges were cut to prevent backlabeling of TIB neurons that grow through the grafts. Motoneurons that regenerated their axons fluoresce in the spinal cord sagittal sections, and were counted using a fluorescent microscope. At the time of retrograde labeling surgery, sections of distal TIB and CP nerves were collected histomorphometry analysis Histomorphometry At the time of the retrograde labeling surgery, sections of the distal CP nerve (~3 mm) were collected and placed in glutaraldehyde nerve fixative solution (Figure 2-7). The nerves were post-fixed with osmium tetroxide, ethanol dehydrated, and embedded for sectioning (0.6 µm). Nerve sections were then stained with toluidine blue for viewing under light microscopy. Entire nerve cross sections were captured at 1000X magnification (Leica DM2500) to count regenerated axons, measure myelin thickness and fibre diameter, and calculate G-ratio using a semi-automated Matlab program (More et al. 2011).

65 Statistical analysis Results are reported as mean ± standard error of the mean. Differences among groups were evaluated using a one-way analysis of variance (ANOVA) followed by a post-hoc multiple comparisons of means (p 0.05). 2.4 Results Force analysis, MUNE, and wet muscle weights Functional recovery following delayed nerve repair with and without placement of 3 side-to-side bridges, and treatment with or without GDNF, was evaluated by isometric twitch force, tetanic force, and motor unit number estimation (MUNE) in the extensor digitorum longus (EDL) and tibialis anterior (TA) muscles 5 months after delayed nerve repair. Results for the TA and EDL muscles were similar (Figure 2-8, Figure 2-9). Comparing treatments of side-to-side bridge protection with and without GDNF revealed no statistical difference in twitch force (GDNF MS Bridges: 0.35 ± 0.04 N; Bridges: 0.32 ± 0.03 N), tetanic force (GDNF MS Bridges: 1.35 ± 0.1 N; Bridges: 1.43 ± 0.1 N), number of reinnervated motor units (MUNE) (GDNF MS Bridges: 26 ± 4; Bridges: 32 ± 2) in the EDL muscles (Figure 2-8A-C). Likewise, there was no significant differences in twitch and tetanic forces produced in the TA muscle between groups that received side-to-side bridge protection of the distal CP stumps with and without GDNF treatment (Figure 2-9A, B). Treatments without side-to-side bridge protection, regardless of GDNF treatment, had no effect in muscle reinnervation, with no differences detected in twitch force

66 56 (GDNF MS: 0.16 ± 0.04 N; No treatment (NT): 0.21 ± 0.03 N), tetanic force (GDNF MS: 0.78 ± 0.18 N; NT: 1.0 ± 0.15 N), and MUNE (GDNF MS: 18 ± 3; NT: 22 ± 2) in the EDL muscle (Figure 2-8A-C). Results from the TA muscle followed the same pattern (Figure 2-9). After force analysis and MUNE procedures were complete, EDL and TA muscles were excised and weighed. Muscle mass is commonly used as a measure of reinnervation since denervated muscle atrophies through proteolytic degradation of muscle fibers (Gutmann & Zelena 1962). Protection of the CP nerves with side-to-side bridges, regardless of GDNF treatment, resulted in higher EDL and TA muscle mass than those that did not receive protection (Figure 2-8D, Figure 2-9D). Interestingly, when the CP nerves did not receive protection of the distal stump, GDNF treatment led to a significant loss of EDL muscle mass compared to the group where the CP nerves received no treatment (GDNF MS: 63 ± 9 mg; NT: 96 ± 7 mg). This loss was no longer significant in the TA muscle (GDNF MS: 241 ± 27 mg; NT: 314 ± 21 mg). As can be observed from the values presented, the presence of side-to-side nerve grafts, regardless of GDNF treatment, led to statistically significant improvement in twitch force, tetanic force, and MUNE compared to groups without distal nerve stump protection. Furthermore, there was significantly less muscle mass loss in groups that received side-to-side bridge protection of the distal nerve stumps. These results complemented the study performed previously by Ladak et al. where the number of regenerated motoneurons was significantly increased in groups with side-to-side bridge protection (Ladak et al. 2011). However, the lack of an effect observed from GDNF is not consistent with previous studies using the microsphere delivery system, although the formulation used here was for a longer release, which may account for the contradicting results (Wood, Gordon, et al. 2012).

67 57 A B EDL Twitch Force (N) * EDL Tetanic Force (N) * No Treatment Bridges GDNF MS GDNF MS Bridges No Treatment Bridges GDNF MS GDNF MS Bridges C D * EDL MUNE * EDL Absolute Muscle Weight (mg) No Treatment Bridges GDNF MS GDNF MS Bridges No Treatment Bridges GDNF MS GDNF MS Bridges Figure 2-8: EDL muscle force measurements, MUNE, and absolute muscle weights 5 months following delayed nerve repair. Tendons of the EDL muscles were tied to a force transducer and twitch and tetanic muscle contractions as well as MUNE were recorded in response to CP nerve stimulation. Absolute muscle weights were measured following the procedure. Comparing side-to-side nerve grafts with and without GDNF revealed no statistical difference in twitch force (A), tetanic force (B), MUNE (C), and muscle weights (D). However, the presence of side-to-side nerve grafts, regardless of GDNF treatment, led to statistical improvement in all metrics compared to groups without bridge protection. Normal values for uninjured nerves are represented by dashed lines.

68 58 A B TA Twitch Force (N) * TA Tetanic Force (N) * No Treatment Saline Bridges GDNF MS GDNF MS Bridges No Treatment Bridges GDNF MS GDNF MS Bridges C D 50 * 600 TA MUNE TA Absolute Muscle Weights (mg) * 0 0 No Treatment Bridges GDNF MS GDNF MS Bridges No Treatment Bridges GDNF MS GDNF MS Bridges Figure 2-9: TA muscle force measurements, MUNE, and absolute muscle weights 5 months following delayed nerve repair. Tendons of the TA muscles were tied to a force transducer and twitch and tetanic muscle contractions as well as MUNE were recorded in response to CP nerve stimulation. Absolute muscle weights were measured following the procedure. Comparing side-to-side nerve grafts with and without GDNF revealed no statistical difference in twitch force (A), tetanic force (B), and muscle weights (D). However, MUNE was lower in the GDNF treated groups (C). The presence of side-to-side nerve grafts in the most part, led to statistical improvements in twitch and tetanic force (A, B), and muscle weights (D), compared to groups without bridge protection. Normal values for uninjured nerves are represented by dashed lines. Force analysis was also performed on the gastrocnemius muscle, innervated by the donor TIB nerve, to evaluate the extent of donor nerve injury after the side-to-side nerve

69 59 bridge protection procedure. The muscles were also excised and weighed. There was no significant difference in the twitch force produced by the gastrocnemius when the TIB nerve was stimulated in the different groups (Figure 2-10A). However, the group with side-to-side bridge protection appears to have lower twitch forces, which is significantly different when compared solely with the group with no treatment. This decrease in force production can be attributed to the small injury created in the TIB nerve in the initial surgery to encourage axon sprouting into the grafts. There was no significant differences in gastrocnemius muscle mass in the different groups (Figure 2-10B). A B Gastrocnemius Twitch Force (N) No Treatment Bridges GDNF MS GDNF MS Bridges Gastrocnemius Absolute Muscle Weight (mg) No Treatment Bridges GDNF MS GDNF MS Bridges Figure 2-10: Gastrocnemius isometric twitch force measurements and absolute muscle weights 5 months following delayed nerve repair. Tendons of the gastrocnemius muscle were tied to a force transducer and twitch muscle contractions were recorded in response to TIB nerve stimulation (A). Absolute muscle weights were measured following the procedure (B). There were no significant differences in twitch force or absolute muscle weights between the groups. The group with the side-to-side bridge protection appears to have lower twitch forces which is significantly different when compared solely with the group receiving no treatment. This decrease can be attributed to the initial injury to the donor TIB nerve that is performed for axons to sprout into the side-to-side nerve grafts.

70 Retrograde labeling A retrograde labeling procedure was performed 5 months after the nerve repair surgery to quantify the number of motoneurons that regenerated their axons into the denervated CP nerve stumps in all the groups. FluoroRuby dye was applied 20 mm distal to the injury site for 1 h during the procedure. Spinal cords and dorsal root ganglia were harvested 7 days later and sectioned for analysis under fluorescence microscopy. The groups that received side-to-side nerve graft protection of the CP stumps did not have high numbers of labelled motoneurons, which does not agree with previous results based on the force analysis and MUNE data. In this technique, the nerve grafts were cut to prevent backlabeling of TIB axons growing into the distal CP nerve, to ensure that no TIB motoneurons that grew axons through the bridges were backlabelled. This can account for the differences with the previous results, since the force analysis methodology did not exclude the effects of the donor TIB axons in muscle production. Comparing with the number of motoneurons that regenerated their axons in the immediate repair groups (as a percentage), groups receiving no treatment (NT) regenerated 64% of motoneurons, compared to only 36% in the group with side-to-side bridges. Furthermore, groups receiving GDNF treatment had worsened regeneration (GDNF: 27%; GDNF Bridges: 11%) compared to the groups without GDNF treatment (Figure 2-11A). The results obtained for sensory neuron counts followed the same pattern as the motoneurons (Figure 2-11B). In this set of experiments there was no control group included in the study using empty microspheres with the 6 week release formulation. This was based on the previous studies that found no difference in regeneration between groups treated with saline and those

71 61 with empty microspheres placed in fibrin (Wood, Kim, et al. 2012). In contrast, when microspheres alone were placed directly on the nerve, the regeneration was significantly decreased potentially due to PLGA degradation by-products and the formation of an acidic environment (Shive & Anderson 1997; Sung et al. 2004). Fibrin degrades with time just as the microspheres do. The longer release formulation used in these experiments might lead to a situation in which the microspheres are directly in contact with the nerve once the fibrin has degraded and provide no buffer for the PLGA acidic by-products. This can lead to potential negative effects. In order to find out whether this was the case, 3 groups were added to the study two with empty microspheres (6 week and 2 week release formulation), and one that was repaired without the delivery system (n = 8 per group). These groups were denervated for a period of 2 months, and retrograde labeling was performed 2 months following repair. The number of motoneurons that regenerated their axons was significantly decreased following treatment with empty microspheres of a long term release, indicating that there was a negative effect associated with the long-term release delivery system which may account for the decreased regeneration observed in the GDNF treated groups (Figure 2-12). There was no toxicity effect of the short-term delivery system, consistent with previous results (Wood, Kim, et al. 2012).

72 62 A B Retrograde Labeled Motoneurons * *** Retrograde Labeled Sensory Neurons * *** 0 0 No Treatment Bridges GDNF GDNF Bridges Immediate Repair No Treatment Bridges GDNF GDNF Bridges Immediate Repair Delayed Repair Delayed Repair Figure 2-11: Number of retrograde labeled CP motoneurons in the spinal cord and sensory neurons in the dorsal root ganglia counted 5 months after immediate or 4 month delayed nerve repair. The number of fluorescently labeled motoneurons were counted in spinal cord sections (50 µm sections, all sections counted, correction factor = 0.6 (Abercrombie 1946)) and DRG sections (20 µm sections, every fifth section counted). The GDNF delivery system worsened motor and sensory neuron regeneration regardless of side-to-side bridge protection (A, B). Side-to-side bridge protection of distal stumps did not increase the number of regenerating motor and sensory neurons when compared to the group receiving no treatment (A, B). Normal values for uninjured nerves are represented by a dashed line. 250 Retrograde Labeled Motoneurons * 0 Empty MS 6 wk release Empty MS 2 wk release No MS Figure 2-12: Number of retrograde labeled CP motoneurons counted in the ventral horn of spinal cords 2 months after 2 month delayed nerve repair, with or without empty MS treatment. CP nerves treated with empty microspheres of a long release formulation had significantly decreased numbers of motoneurons that regenerated their axons into the CP distal stump. Short-term release microspheres did not demonstrate a negative effect on regeneration.

73 Histomorphometry Distal CP nerve sections were harvested at the time of retrograde labeling 5 months after delayed nerve repair. Examination of the images obtained under light microscopy (Figure 2-13) indicated that groups receiving the GDNF delivery system treatment had significantly affected nerve morphology, with significantly less axons present in the nerve cross-sections. All experimental groups differed from the normal control nerve in morphology. A B C No treatment Bridges GDNF MS D E F GDNF MS + Bridges Immediate Repair Normal Figure 2-13: Representative sections of axons growing in the distal CP stump 5 months after nerve repair. (A) No treatment (NT); (B) Side-to-side bridges; (C) GDNF delivery; (D) GDNF microspheres and side-to-side bridges; (E) immediate repair; (F) normal nerve. Groups with no GDNF treatment (No treatment; Bridges) demonstrate more similar nerve architecture to the immediate repair group as compared to the groups with the GDNF delivery system (GDNF MS; GDNF MS + Bridges). Histomorphometry measures, including myelinated axon number (Figure 2-14A), myelin thickness (B), fiber diameter (C), and G-ratio (D), were quantified for all groups.

74 64 There was no significant difference in the number of axons that regenerated in the delayed repair groups with no GDNF treatment, with and without bridges (NT: 3028 ± 271; Bridges: 2639 ± 270). Furthermore, there was no significant difference in the number of axons that regenerated into the CP nerves treated with GDNF (GDNF MS: 1349 ± 270; GDNF MS Bridges: 1256 ± 205). The groups receiving the GDNF delivery system had a significantly lower number of axons regenerating in the distal CP nerve stumps. This is indicative of a negative effect of GDNF treatment, which could be a combined effect of microsphere toxicity or a negative effect of an excessive dose of GDNF leading to coil formation at the site of GDNF delivery (Piquilloud et al. 2007; Eggers et al. 2013; Tannemaat, Eggers, Hendriks, de Ruiter, et al. 2008). There was no significant differences in myelin thickness and fiber diameter in all the groups, and all were statistically equivalent to the immediate repair group. The G-ratio, calculated as the ratio of axon diameter and total fiber diameter, was statistically equivalent for all groups, and approached the value of for maximum conduction velocity (Waxman 1980).

75 65 A B Myelinated Axon Number * Myelin Thickness ( µm) No Treatment Bridges GDNF MS GDNF MS Bridges Immediate Repair No Treatment Bridges GDNF MS GDNF MS Bridges Immediate Repair Delayed Repair Delayed Repair C D Fiber Diameter ( µm) 10 5 G-ratio No Treatment Bridges GDNF MS GDNF MS Bridges Immediate Repair No Treatment Bridges GDNF MS GDNF MS Bridges Immediate Repair Delayed Repair Delayed Repair Figure 2-14: Histomorphometric analysis of CP nerves 5 months after nerve repair. Nerves were harvested prior to the retrograde labeling procedure for processing and staining with toluidine blue. (A) Number of myelinated axons; (B) myelin thickness; (C) fiber diameter; (D) G-ratio. The groups with no GDNF treatment had increased myelinated axon counts compared to the groups with the GDNF delivery system. No groups exhibited significant differences in myelin thickness, fiber diameter, and G-ratio, but all were below the values of normal uninjured nerves (demonstrated by the dashed line). 2.5 Discussion The side-to-side bridging technique was effective in protecting against the negative effects of chronic denervation on nerve regeneration, regardless of treatment with or without GDNF in terms of muscle force and muscle mass outcome measures. This was not reflected

76 66 in the backlabeling data, which showed a significantly lower number of motoneurons and sensory neuron counts in the groups receiving CP nerve protection with side-to-side bridges, with and without GDNF treatment. Histomorphometry analysis of nerve sections showed a decrease in the number of regenerated axons after treatment with side-to-side bridge protection and GDNF, although this decrease was not observed without the GDNF treatment. This indicated that the protection effects of the side-to-side bridges were compromised by the GDNF delivery system. Although the number of axons regenerating through the CP nerve was statistically equivalent in groups with and without side-to-side bridge protection, the number of functional axon connections made with muscle fibers was significantly increased in groups with the side-to-side nerve bridge protection, as indicated by the improvement in force, MUNE, and muscle weight outcome measures. These positive results imply that the side-to-side nerve grafts provided protection against denervation. However, the positive results observed in the muscle reinnervation measures may include the effects of the axons that grow through the side-to-side bridges. The lack of improvement in the number of CP motoneurons that regenerated their axons through protected distal stumps indicate that this might be the case. One of the ways in which the protective mechanism of the side-to-side bridges may work is by maintaining the Schwann cells (SCs) in the distal CP stump in a regenerative state. Axons that sprout from the TIB nerve through the resident SCs in the grafts grow into the denervated stump and may through contact with the SCs residing in the distal stump prevent them from changing into quiescent or atrophic states. For instance, SC proliferation is mediated by the binding of erbb receptors at the SC surface with neuregulin expressed by the growth cones of regenerating axons (Pellegrino & Spencer 1985; Li et al. 1997). Current

77 67 studies in our laboratory are trying to determine the phenotype of resident SCs in CP distal stumps in groups with and without protection, and whether neuregulin expression regulates the positive effect of the bridges. SC migration is another possible mechanism by which the bridges exert their protective effects. SCs might migrate from the side-to-side nerve grafts along with regenerating axons to populate the denervated distal stump. Local delivery of GDNF for a period of 6 weeks did not increase axonal regeneration 5 months after delayed repair based on force analysis, and seemed to worsen regeneration based on the backlabeling data. Histomophometry analysis of nerve sections did not reveal any differences in the number of regenerated axons between groups with no GDNF treatment, and furthermore revealed that groups treated with GDNF had significantly impaired regeneration. Because no empty microspheres were used in the negative control groups it was impossible to identify whether the negative effects observed from the delivery system were due to a toxicity effect from the microspheres alone, or whether it was also from an overdose of GDNF at the delivery site. We introduced two groups of animals to evaluate whether there was a toxicity effect associated with the delivery system with the longer release rate and degradation time. The number of retrograde labeled motoneurons in groups treated with empty MS formulation in fibrin (6 week release) was significantly lower compared to groups receiving no treatment. This indicated that there was indeed negative effects associated with the delivery system that influenced the results. Sung and colleagues (2004) studied the effects of PLGA degradation on cell viability. They found that the acidic ph occurring from fast degradation of PLGA resulted in lower cell viability (Sung et al. 2004). Based on studies performed in our laboratory, when microspheres with a 2 week release formulation were placed in direct contact with regenerating nerves, regeneration was

78 68 inhibited. However, the negative effects were reversed after incorporation of microspheres in fibrin (Wood et al. 2013). The addition of fibrin may prevent the fast degradation of PLGA, and thus the negative effects. However, because fibrin degrades with time, the microsphere formulation used in this study for a longer release period (6 week), might lead to a situation in which the microspheres are no longer protected from fast degradation once the fibrin has degraded. This may account for the negative effects observed. However, in force, MUNE, and muscle weight measures, the side-to-side bridges were able to improve regeneration regardless of the delivery system. The formulation of microspheres used in these experiments (6 week release) was not previously tested in vivo. Because of the longer regeneration periods in this study, the longer period of GDNF release was deemed appropriate. Wood and colleagues (2012) used GDNF microspheres with release formulations of 2 and 4 weeks (Wood, Gordon, et al. 2012). They evaluated nerve regeneration in a 2 month chronic injury model followed by repair and a 1 or 3 month regeneration period. In our study, the period of chronic axotomy and denervation was 4 months, and outcome measures were evaluated 5 months following nerve repair. The longer period of axotomy and denervation (4 months) ensured sufficient time for the growth of donor axons from the TIB nerve into the denervated CP distal stump, given the staggered growth across each suture site. Furthermore, it ensured a state of chronic injury, as most growth-associated genes cease to be upregulated by this time (Boyd & Gordon 2003b). The longer period of regeneration was chosen to ensure axonal connection with muscle fibers, given the slow regeneration of axons at 1-4 mm per day (Grafstein & Forman 1980). It is possible that by this time the positive effects of GDNF may have levelled out as has been previously observed (de Boer et al. 2012; Piquilloud et al. 2007). De Boer and colleagues

79 69 previously found that delivery of GDNF in microspheres resulted in increased number of myelinated fibers when compared with the saline control at an early timepoint of 6 weeks; however, at 16 weeks, this difference was no longer observed (de Boer et al. 2012). Piquilloud et al. (2007) found that GDNF delivery induced an overgrowth of nerve fibers at the site of growth factor delivery, which increased progressively with higher GDNF concentrations. However, there was no difference in the number of axons regenerating in the distal segment of the nerves in all groups (Piquilloud et al. 2007). Additionally, the speed of recovery was decreased in animals receiving GDNF treatment, although 3 months later the level of recovery was similar in all groups. Another issue with the use of this delivery system is that although it was tested in vitro, the release kinetics are very likely to differ in an in vivo environment. It was previously shown that GDNF administered at a rate of 0.1 µg/day was sufficient to improve regeneration (Boyd & Gordon 2003a). Our in vitro studies suggested that the amount of GDNF released per day met the daily suggested dose; however, the in vivo release may have resulted in a GDNF concentration that was suboptimal. There is also a possibility that a longer release of GDNF may have led to the decreased axonal regeneration into the distal nerve stump by causing the axons to coil at the site of GDNF delivery. Several investigators have shown that an overdose of exogenous GDNF can inhibit nerve regeneration (Eggers et al. 2008; Tannemaat, Eggers, Hendriks, de Ruiter, et al. 2008; Eggers et al. 2013). Overall, the side-to-side bridging technique was effective in protecting against the negative effects of chronic denervation based on muscle force and MUNE analysis. Local delivery of GDNF for a period of 6 weeks did not increase functional recovery and based on retrograde labeling data impaired axonal regeneration. There are many questions that remain

80 70 to be answered: 1) is the impaired regeneration after GDNF treatment observed in retrograde labeling the result of axon coiling or delivery system toxicity? 2) Is the lack of functional recovery differences between groups with and without GDNF treatment due to the outcome measurements being performed really late into the regeneration process? 3) Are the discrepancies from the previous studies due to different timepoints? To answer these remaining questions further experiments were performed using the same experimental groups as in this study with some alterations to the experimental design (Refer to Chapter 3). The period of axotomy and denervation was shortened from 4 months to 2 months, similar to previous studies performed with GDNF (Boyd & Gordon 2003b; Wood, Gordon, et al. 2012; Wood et al. 2013). Furthermore, GDNF delivery was performed using GDNF with a 2 week release formulation as opposed to a 6 week release formulation, and empty microspheres were introduced as a control. Outcome measures including retrograde labeling, histomorphometry analysis of nerve sections, and MUNE were performed 2 months following the delayed nerve repair.

81 Chapter 3. Short-Term Delivery Experiments The experiments presented in this chapter were introduced based on the results of the long-term experiments presented in Chapter 2. In those studies, the glial cell-line derived neurotrophic factor (GDNF) delivery system released GDNF for a period of 6 weeks and was found to impair regeneration when compared to groups that did not receive the GDNF treatment. This observation was consistent with several studies that found excessive doses of GDNF to be detrimental to nerve regeneration as a result of axon coiling at the site of GDNF delivery (Eggers et al. 2013; Eggers et al. 2008; Tannemaat, Eggers, Hendriks, de Ruiter, et al. 2008; Blits et al. 2004; Georgievska et al. 2002). However, it was undetermined whether the negative effects were due to the longer release of GDNF or solely a result of delivery system toxicity. In this study a GDNF delivery system that released GDNF for a period of 2 weeks was used to prevent the potential putative coiling from occurring. The same experimental groups were utilized as those presented in Chapter 2 with some alterations. The period of axotomy and denervation was shortened from 4 months to 2 months, similar to the previous studies with GDNF, and the period of regeneration was shortened from 5 months to 2 months (Boyd & Gordon 2003a; Wood, Gordon, et al. 2012; Wood, Kim, et al. 2012). Outcome measures for regeneration included compound muscle action potential, motor unit number estimation, retrograde labeling, and histomorphometry analysis of the nerves. 71

82 Abstract The objective of this study was to evaluate the peripheral nerve regenerative effects of treatment with exogenous GDNF delivery, side-to-side bridge protection, or both, in a model of chronic axotomy and denervation. A delayed nerve repair model was created by transection of the common peroneal (CP) nerve and suture of the proximal and distal nerve stumps to innervated muscle. Nerve grafts obtained from the contralateral side were used as side-to-side nerve grafts in the experimental groups. After 2 months the CP nerves were repaired, and the delivery system containing microspheres loaded with GDNF (termed GDNF MS ) or double distilled water (ddh 2 O empty MS ), was placed surrounding the repaired nerves. Axonal regeneration and muscle reinnervation were assessed 2 months after delayed nerve repair by several modalities including compound muscle action potential (CMAP), motor unit number estimation (MUNE), muscle weights, retrograde labeling, and histomorphometry. Comparing groups receiving treatments with and without GDNF revealed no statistical difference in retrograde labeled neurons, regenerated axons, CMAP area, and tibialis anterior (TA) muscle weight. The presence of side-to-side nerve grafts, regardless of GDNF treatment, led to a significant increase in the number of regenerated axons, CMAP, and TA muscle mass. However, no differences were found in the number of motoneurons that regenerated their axons through protected and unprotected CP stumps. We failed to see an impact in regeneration with GDNF delivery, but showed that side-to-side nerve grafts can be used to improve several aspects of regeneration following delayed repair.

83 Methods Experimental Design Thirty-six female Sprague-Dawley rats ( g) were divided into 4 groups of delayed nerve repair as shown in Table 3-1 (groups 1-4). Additionally, a fifth group undergoing immediate nerve repair served as a positive control group (n = 8). Each group was evaluated by retrograde labeling and histomorphometry, as well as compound muscle action potential (CMAP), motor unit number estimation (MUNE), and muscle weight. In the first surgery, all animals (groups 1-5) underwent a common peroneal (CP) nerve transection (refer to Figure 3-1 for a timeline). The CP nerve of animals in group 5 was immediately repaired using two 10-0 microsutures. In groups 1-4 the proximal and distal nerve stumps were sutured to innervated muscle for a period of 2 months to prevent axon regeneration into the distal CP nerve stump (groups 1 and 3 Figure 3-2A). Additionally, nerve grafts obtained from the contralateral side were used as side-to-side nerve grafts (4 mm each) connecting the injured CP nerve to the intact tibial (TIB) nerve in groups with the bridge protection (groups 2 and 4, Figure 3-2B). Following 2 months of chronic axotomy and denervation, a nerve repair surgery was performed on groups 1-4 (Figure 3-2C-F). At this time the repaired CP nerves were surrounded by a delivery system. The delivery system consisted of poly(lactic-co-glycolic acid) (PLGA) microspheres loaded with GDNF or double distilled water (ddh 2 O) embedded in fibrin for non-invasive controlled delivery. Two months after the CP nerve repair surgery outcome measures were evaluated.

84 74 Figure 3-1: Timeline of experimental treatments During the first surgery, the CP nerves were transected and sutured to innervated muscle, with or without side-to-side bridge protection. After 2 months of chronic axotomy and denervation, the CP nerves were repaired and the animals were treated with either empty MS or GDNF MS embedded in fibrin. Outcome measures were evaluated 2 months after the repair surgery. Table 3-1 Control and experimental groups Rats were divided into 5 groups of delayed and immediate nerve repair. Nerve regeneration was evaluated using retrograde labeling, histomorphometry, muscle mass, and motor unit number estimation (MUNE). Delayed nerve repair Group Treatment Outcome measure n 1 No protection + Empty MS Retrograde labeling, histomorphometry, muscle mass, MUNE 2 Bridge protection + Empty MS Retrograde labeling, histomorphometry, muscle mass, MUNE 3 No protection + GDNF MS Retrograde labeling, histomorphometry, muscle mass, MUNE 4 Bridge protection + GDNF MS Retrograde labeling, histomorphometry, muscle mass, MUNE Immediate nerve repair Treatment Outcome measure n 5 None Retrograde labeling, histomorphometry 8

85 Figure 3-2: Schematic of experimental design. All animals underwent a common peroneal (CP) nerve transection, in which the proximal and distal CP nerve stumps were sutured to innervated muscle for 2 months (A). Additionally, half of the animals received side-to-side bridge protection of the distal CP nerve stump (B). After 2 months, the CP nerve was repaired in all animals in groups 1-4 (C-F), and the CP nerves of animals in groups 1 and 2 were surrounded by an empty MS delivery system and groups 3 and 4 (C-D) were surrounded by a GDNF delivery system (E-F). 75

86 Preparation of microsphere delivery system In this experiment, a delivery system releasing GDNF for 2 weeks was used for comparison with delivery of GDNF for 6 weeks that was found to impair regeneration in the previous study, conceivably because of an excess source of GDNF or delivery system toxicity (Refer to Chapter 2). The technique for microsphere fabrication is identical to that presented in Chapter 2 (section 2.3.2) except that in this experiment (2 week release), PLGA with a lower molecular weight was used (5,000 Da). For the previous experiment (6 week release), PLGA of higher molecular weights were used (6,700 Da and 12,900 Da), to maintain GDNF delivery for longer periods GDNF microspheres characterization To quantify GDNF encapsulation efficiency 5 mg of GDNF microspheres were dissolved in 0.5 ml dichloromethane and 0.5 ml ddh 2 O. The solution was vortexed for 10 min and centrifuged at 1,000 rpm for 5 min. The supernatant was then collected. The process was repeated three more times and the pooled aqueous portion was assayed using an ELISA as per manufacturer s instructions (R & D systems). The encapsulation efficiency was calculated as the ratio of the mass of the retrieved GDNF in the aqueous phase compared to the theoretical mass of GDNF from 5 mg of microspheres (5 µg). The theoretical mass of GDNF in the microspheres was determined based on the initial loading ratio of 1 µg GDNF / 1 mg PLGA during the microsphere fabrication process.

87 In vitro GDNF release The release of GDNF from the delivery system was evaluated for 15 days and expressed in terms of cumulative release as described previously (Wood, Gordon, et al. 2012). Briefly, the delivery system was placed in 2 ml low-retention Eppendorf tubes and incubated in 1 ml of release media (1% BSA in PBS) at 37ºC for two weeks under gentle agitation. The release media was collected daily and replaced. Samples were stored at -20ºC until evaluation for GDNF quantity using an ELISA assay as per manufacturer s instructions (R & D systems) DRG neurite outgrowth assay for GDNF bioactivity Bioactivity of GDNF released from the delivery system was evaluated using a dorsal root ganglia (DRG) neurite outgrowth assay with the help of Jackie Obermeyer from the Shoichet laboratory. DRGs were dissected from day 15 rat embryos (E15 female Sprague- Dawley rats) and placed in modified neurobasal media (NBM) with 2 vol. % B-27 serumfree supplement, 1 vol.% penicillin-streptomycin, and 1 vol.% L-glutamine (Stanwick et al. 2012). The DRG were then placed in a 24-well plate with 12 mm diameter glass cover slips that had been pre-coated with poly-d-lysine (50 µg/ml) overnight and laminin (5 µg/ml) for 2 hr at 37ºC. The wells were treated with 0.5 ml of modified NBM media and 0.5 ml of release samples from in vitro release samples collected at day 3, 7, and 14 of the release period. For controls, 0.5 ml of 1% BSA with GDNF concentrations of 0, 10, 50, and 100 ng/ml was added to the wells. These concentrations were chosen based on previous studies that found neurite outgrowth to be maximal at these GDNF concentrations (Garbayo

88 78 et al. 2008; Garbayo et al. 2007; Wood, Borschel, et al. 2009). Neurite outgrowth was measured after 48 h and quantified using the method described by Herbert and colleagues (1996) in two consecutive experiments, for a total of 10 DRG per group. Average neurite lengths were obtained by calculating the width of an annulus with area equal to the area of the neurite zone (from the edge of the cluster of cell bodies to the wavefront of the growing neurites).this was calculated as L = (1/π) 1/2 [(A cell cluster + A area of neurites ) 1/2 (A cell cluster ) 1/2 ], where L = average neurite length of a circular ganglion, A cell cluster = area of the cell cluster of the ganglion, and A area of neurites = area of the neurite zone. The average neurite outgrowth was normalized to the neurite extension of controls with GDNF at 100 ng/ml, a concentration that had consistent results between the two consecutive experiments (Wood, Borschel, et al. 2009) Surgical procedure Surgical procedures were carried out in the same manner as presented in Chapter 2 (section 2.3.3), with alterations in time points. All animals had a CP nerve transection surgery with or without side-to-side bridge protection, and 2 months later the nerve repair surgeries were performed with either empty (ddh 2 O) or GDNF loaded microspheres embedded in fibrin, depending on treatment group. Animals were allowed to convalesce for a period of 2 months after which outcome measures were evaluated (Refer to Figure 3-1, 3-2 for timelines and experimental design schematic).

89 Electrophysiology Two months after the CP nerve repair surgeries rats were anesthesized with isofluorane. The nerve and the tibialis anterior muscle (TA) were exposed using aseptic techniques. Recording electrodes were placed in the TA muscle, and stimulating electrodes were placed in the distal CP nerve (Figure 3-3). Electrophysiological parameters were evaluated as performed previously (Willand et al. 2011). First, the maximum compound muscle action potential (CMAP) was recorded, representing the sum of individual motor unit potentials. Twenty templates were then recorded by gradually increasing the stimulus from threshold, causing distinct increments in CMAP amplitude. The responses were visually inspected and identical responses were deleted. The average size of a motor unit was calculated using a Matlab script that ranked the templates by absolute area (voltage time) and subtracted each template from the next largest. The number of reinnervated motor units was then calculated by the ratio of the maximum CMAP and the average size of a motor unit. A Stimulating Electrodes B CP nerve TIB nerve Recording Electrodes Figure 3-3: Recording of CMAP and MUNE in vivo (A) Schematic of experimental setup for electrophysiology recordings. (B) Stimulation of the CP nerve in vivo and recording the response from the TA muscle in an aseptic manner to ensure survival for retrograde labeling evaluation a week later.

90 Retrograde labeling After the electrophysiology procedures were performed, the distal CP nerve was transected 20 mm distal to the repair site. A 4% FluoroGold dye solution was applied at the tip of the transected CP nerve in a Vaseline well for 1 h. Additionally, the TIB nerve of the groups with side-to-side bridge protection was tied to prevent backlabeling of TIB axons that grow through the side-to-side branching (Figure 3-4). Animals were allowed to recover for 7 days, to allow for retrograde transport of the dye to the cell bodies prior to animal perfusion and tissue harvest. Animal perfusion and motoneuron counts were performed as described previously in Chapter 2 (section 2.3.5). At the time of sacrifice, TA muscles were weighed and CP nerves were harvested for immunohistochemistry.

91 81 A DYE CP nerve SPINAL CORD TIB nerve B C TIB nerve CP nerve Figure 3-4: Retrograde labeling procedure (A) Schematic of retrograde labeling procedure. The distal CP nerve was transected 20 mm distal to the repair site and a dye was applied at the tip of the nerve in a Vaseline well for 1 h. In groups with side-to-side bridges the TIB nerve was tied. (B) Labeled motoneurons in a spinal cord section under fluorescent microscopy. (C) In vivo surgical procedure Histomorphometry CP nerve sections were harvested at the time of retrograde labeling and analyzed for histomorphometry as described previously in Chapter 2 (section 2.3.6) Immunohistochemistry At the time of sacrifice, 7 days after the retrograde labeling surgery, CP nerves were harvested and fixed in 4% paraformaldehyde and cryoprotected in 30% sucrose at 4ºC overnight. Tissue samples were embedded in tissue freezing medium (Electron Microscopy

92 82 Science) and cryosectioned (20 µm sections). Staining techniques were performed as previously described (Kemp et al. 2009). Briefly, slides were washed (1% Triton-X in PBS) and blocked for 1 h at room temperature (0.1% BSA/10% donkey serum in PBS). Slides were then incubated with monoclonal mouse antibody neurofilament-160 (Sigma, 1:200) overnight at 4ºC. The next day, slides were washed and incubated with donkey anti-mouse AlexaFluor 568 (Sigma, 1:500) for 2 h at room temperature. After the last washing step, cover slips were mounted and imaged under fluorescence microscopy (Leica DM2500). Normal CP nerves served as positive controls and negative controls omitted the primary antibody. Analysis of axonal regeneration through the nerve was analyzed similarly to the method used by Kemp and colleagues (Kemp et al. 2009). Longitudinal sections of CP nerve were stained with neurofilament and the axonal profiles were counted at three points in the CP nerve: at the suture site, slightly after the suture site, and distally. Axon profiles crossing a perpendicular line in the middle of the field in each section were counted by a blinded observer Statistical analysis Results are presented as means ± standard error of the means. Statistical analysis was evaluated using a one-way analysis of variance (ANOVA) followed by a post-hoc multiple comparison of means when appropriate (p 0.05).

93 Results Microsphere characterization Microspheres used in this study were formulated to release GDNF for a period of 2 weeks, since results obtained in the first study (Chapter 2) indicated that a longer release period may be detrimental to regeneration. The microspheres were constructed as described previously and yielded an encapsulation efficiency of 59±7% (Wood, Gordon, et al. 2012). The release profile of the GDNF delivery system (microspheres embedded in fibrin) was observed in vitro for a period of 2 weeks. Eighty percent of the encapsulated GDNF was released at the end of the 15 day period (Figure 3-5). This release profile was similar to that previously observed (Wood, Gordon, et al. 2012) and the microspheres released 0.1 µg/day starting at day 5. This concentration was previously determined to be effective at promoting nerve regeneration following delayed nerve repair using a mini-osmotic pump (Boyd & Gordon 2003a). Figure 3-5: GDNF in vitro release profile Release of GDNF from microspheres embedded in fibrin gels was observed for a period of 2 weeks. The GDNF delivery system released 0.1 µg/day starting at day 5, and released 80% of the encapsulated GDNF by day 15.

94 DRG bioactivity assay The bioactivity of GDNF released from the delivery system was evaluated based on neurite outgrowth from rat embryonic (E15) DRGs. DRGs contain a subpopulation of sensory neurons which expresses the receptor GFRα1, and are thus responsive to GDNF (Gavazzi et al. 1999). The release media from the delivery system collected at day 3, 7, and 14 of the release duration was diluted in media where rat E15 DRGs were cultured. After 48 h of culture, brightfield images of the DRGs and their projected neurites were captured. The length of the neurites was quantified by the method described by Herbert and colleagues, and was normalized to the neurite extension of DRGs at 100 ng/ml (Herbert et al. 1996). The morphology of DRG neurite outgrowths was similar in all samples tested. Non-neuronal cells and neurites advanced to form a starburst pattern, with neurites growing in advance of non-neuronal migrating cells (Figure 3-6). This type of growth was observed previously in chick embryonic DRGs (Herbert et al. 1996). Previous use of this GDNF delivery system demonstrated improved regeneration after delayed repair, so it was predicted that the bioactivity of GDNF was maintained throughout the delivery period. The results demonstrated that the GDNF released from the delivery system was not significantly different from the positive control (media supplemented with 100 ng/ml GDNF). Furthermore, neurite extension from DRGs treated with GDNF from the positive control and release media was significantly higher than the negative control (release media from blank microspheres), demonstrating that the release delivery system did not have a negative effect on GDNF bioactivity (Figure 3-7, Figure 3-8).

95 85 Figure 3-6: Morphology of neurite outgrowth from DRGs Images taken 48 h after tissue culture of DRGs, showing non-neuronal cells (nn, black arrows) and neurites (n, yellow arrows). (A) Image of neurites growing from the DRG. (B) Image at the mid-section of the neurite outgrowth zone showing neurites and non-neuronal cells. (C) Image focused at the end of neurites, with a lower number of non-neuronal cells present. Normalized neurite extension * Control 0 ng/ml Control 100 ng/ml day 3 release day 7 release day 14 release Figure 3-7: Effect of GDNF on DRG neurite outgrowth length DRG neurite extension was evaluated 48 h after incubation in vitro. Neurite extension of DRGs supplemented with GDNF released from the delivery system was not significantly different than the positive control (100 ng/ml). Furthermore, neurite extension of DRGs supplemented with GDNF (both positive control and GDNF delivery system release medium) was significantly different from the negative control, at day 3 and 7.

96 86 Figure 3-8: DRG neurite outgrowth images Images taken 48 h after tissue culture of DRGs with (A) media enriched with 100 ng/ml GDNF, (B) GDNF delivery system release sample, and (C) release sample from blank microspheres (0 ng/ml GDNF). DRGs treated with GDNF enriched medium (A, B) had visually longer neurites extending from the cell clusters, compared to the negative control group (C) Functional outcome measures Electrophysiological parameters were characterized in the TA muscle 2 months following delayed nerve repair, including CMAP, MUNE, and motor unit size. Muscle weights were also recorded 7 days after the retrograde labeling procedure. The area under the CMAP curve for each group was determined because it is a better estimation of axon reinnervation than CMAP amplitude alone (Archibald et al. 1991; Brushart 2011). The groups receiving side-to-side bridge protection of the distal CP stump treated with GDNF MS (301 ± 48 mv s) or with empty MS (394 ± 20 mv s) had significantly higher CMAP values compared to groups treated with only GDNF (119 ± 17 mv s) or empty MS (187 ± 22 mv s). Furthermore, treatment with GDNF resulted

97 87 in no significant difference in CMAP values in groups with and without side-to-side bridge protection, although the groups treated with GDNF appeared to be lower (Figure 3-9A). A Area of Max CMAP (mv s) * B TA MUNE * 0 0 Empty MS Empty MS Bridges GDNF MS GDNF MS Bridges Empty MS Empty MS Bridges GDNF MS GDNF MS Bridges C D Motor Unit Size (mv s) * TA Absolute Muscle Weight (mg) * 0 0 Empty MS Empty MS Bridges GDNF MS GDNF MS Bridges Empty MS Empty MS Bridges GDNF MS GDNF MS Bridges Figure 3-9: Functional outcome measures recorded from the tibialis anterior (TA) muscle 2 months following 2 months delayed nerve repair. (A) Area under the curve of the maximum CMAP wave elicited. (B) Estimate of the number of functional motor units (MUNE). (C) Size of each motor unit as determined by the ratio of the max CMAP and the number of motor units. (D) Absolute weight of the TA muscle 7 days following the retrograde labeling procedure. MUNE was performed to evaluate the number of motor units that were reinnervated in the TA muscle (Figure 3-9B). The size of the motor units was determined by dividing the

98 88 area of the maximum CMAP by the number of reinnervated motor units (MUNE) (Figure 3-9C). The group with only empty MS treatment had a significantly higher number of reinnervated motor units (94 ± 9) than any other group, and also had the lowest size of motor units (2 ± 0.2). In this study the number of reinnervated motor units was evaluated using electrophysiology. This technique had not yet been optimized when electrophysiology testing of the first group (empty MS) was performed, which may account for its unexpectedly high numbers. The high number of reinnervated motor units obtained in the TA muscles in the empty MS group (94 ± 9) were not significantly different than normal uninjured muscle, which indicates an error in the MUNE methodology since a significant decrease from normal is expected after chronic axotomy and denervation (Wood et al. 2013). Changes in the experimental setup and optimization that occurred after the first experimental session may have affected the results. Apart from the first group tested (empty MS), the rest of the groups follow the same patters as those observed in the CMAP and muscle weight outcome measures. The group with side-to-side bridge protection of the distal CP stump and empty MS treatment had 55 ± 11 reinnervated motor units, and the size of the motor units (15 ± 6) was higher than any other group. Figure 3-10 shows typical CMAP potentials and the corresponding templates for TA muscles in the empty MS treatment group and the group receiving side-to-side bridge protection of the injured CP stump. The group receiving empty MS treatment had consistently lower CMAP amplitudes compared to groups with side-to-side bridge protection but the templates obtained were lower in magnitude than those of the groups with side-to-side bridge protection, corresponding to smaller muscle units (Note the difference in scales in the figure). For the GDNF treated groups, MUNE and the size of the motor units were higher in the group with side-to-side bridge protection (MUNE:

99 89 49 ± 3; Size: 6.4 ± 3 1.2) than in the group without protection (MUNE: 36 ± 4; Size: 3.0 ± 0.4). This indicated that although GDNF impaired regeneration to some extent, the side-to-side bridges were able to counteract some of the negative effects. Absolute TA muscle mass was recorded 7 days after the last retrograde surgery procedure to evaluate muscle atrophy (Figure 3-9D). The groups receiving side-to-side bridge protection of the distal CP stumps and treatment with GDNF MS (239 ± 17 mg) or empty MS (238 ± 12 mg) had significantly decreased muscle atrophy than groups treated with only GDNF (170 ± 11 mg) or empty MS (164 ± 10 mg). These results were consistent to the trends observed with CMAP area. A CMAP Empty MS B templates Empty MS C Bridges Empty MS D Bridges Empty MS voltage (mv) time Figure 3-10: CMAP amplitude and templates for MUNE calculations CMAP amplitudes for groups without (A) and with (C) side-to-side bridge protection. Templates recorded by gradually increasing the stimulus from threshold and causing distinct increments in CMAP amplitude in groups without (B) and with (D) side-to-side bridge protection.

100 Retrograde labeling To quantify the number of motoneurons that regenerated their axons into the denervated CP nerve stumps in the different groups, Fluorogold dye was applied 20 mm distal to the injury site 2 months following delayed nerve repair. Spinal cords were harvested 7 days later and sectioned for analysis under fluorescent microscopy (Figure 3-12). Half the number of CP motoneurons regenerated their axons after 2 months of regeneration in the groups that received side-to-side bridge protection of the distal CP stump, when compared to the group that received immediate nerve repair. The group with only GDNF treatment at the injury site regenerated 28% of motoneurons (compared to the immediate repair group) which increased to 41% with the addition of the side-to-side bridge protection. No significance was found amongst any of the groups, apart from the immediate repair group. These results were consistent with the estimation of the number of reinnervated motor units using MUNE analysis, where no differences were found between the groups apart from the empty MS group.

101 * Retrograde Labeled Motoneurons Empty MS Empty MS Bridges GDNF GDNF Bridges Immediate Repair Delayed Repair Figure 3-11: Number of retrograde labeled motoneurons counted in the spinal cords 2 months after immediate or 2 month delayed nerve repair Fluorescently labeled CP motoneurons were counted to evaluate the number of motoneurons that regenerated their axons into the denervated CP nerve stump in the different experimental groups. No differences were found amongst the groups undergoing delayed nerve repair.

102 92 A B C D E Figure 3-12: Pictures of retrograde labeled motoneurons as observed under fluorescent microscopy. Representative slides of motoneurons labeled in the following groups: (A) Empty MS; (B) Empty MS + Bridges; (C) GDNF MS; (D) GDNF MS + Bridges; (E) Immediate repair Histomorphometry Nerve sections were harvested for nerve morphometry analysis 2 months after delayed or immediate nerve repair. Sections were imaged under light microscopy and qualitatively examined for differences (Figure 3-13). As was observed previously, axons growing through the CP nerve receiving GDNF treatment were distributed in distinct bundles as opposed to being evenly distributed throughout the nerve (Figure 3-13C, D) (Wood et al. 2013). Groups receiving side-to-side nerve bridge protection of the distal CP stump closely resembled the nerve morphology of an immediately repaired nerve (Figure 3-13B, D, E). All groups were visually different from the normal control nerve where the axons were

103 93 uniformly distributed, demonstrated no axonal bundles, and had larger diameters (Figure 3-13F). A B C Empty MS Empty MS + Bridges GDNF MS D E F GDNF MS + Bridges Immediate Repair Normal Figure 3-13: Representative sections of axon morphology from distal CP nerve cross sections collected 2 months following nerve repair. (A) Empty MS; (B) Side-to-side bridges and Empty MS; (C) GDNF MS; (D) Side-to-side bridges and GDNF MS; (E) immediate repair; (F) normal nerve. Groups with side-to-side bridge protection demonstrate nerve architecture that more similarly resembles the immediate repair group. Histomorphometric measures, including myelinated axon number, fiber diameter, myelin thickness, and G-ratio were quantified for all groups (Figure 3-14A-D). Myelinated axon numbers were significantly higher in groups with side-to-side bridge protection. Groups with empty MS or GNDF MS alone were not statistically different from one another in numbers which has been previously reported in the use of this delivery system (Wood et al. 2013).

104 94 A B * 2.5 Myelinated Axon Number Myelin Thickness ( µm) Empty MS Empty MS Bridges GDNF GDNF Bridges Immediate Repair 0.0 Empty MS Empty MS Bridges GDNF GDNF Bridges Immediate Repair C Delayed Repair D Delayed Repair Fiber Diameter ( µm) 10 5 G-ratio Empty MS Empty MS Bridges GDNF GDNF Bridges Immediate Repair Empty MS Empty MS Bridges GDNF GDNF Bridges Immediate Repair Delayed Repair Delayed Repair Figure 3-14: Histomorphometry analysis of CP nerves 2 months after immediate or 2 month delayed nerve repair. Nerves were harvested prior to the retrograde labeling procedure for processing and staining with toluidine blue. (A) Number of myelinated axons; (B) myelin thickness; (C) fiber diameter; (D) G-ratio. The groups that received side-to-side bridge protection of the distal CP stump, regardless of GDNF treatment, had increased number of myelinated axon counts compared to the groups with no protection. There was no significant differences in myelin thickness, fiber diameter, and G-ratio between treatment groups. Normal values for uninjured CP nerves are presented as dashed lines.

105 Immunohistochemistry analysis Immunostaining with neurofilament antibody was performed on longitudinal sections of the CP nerves that received GDNF MS or empty MS treatments to evaluate axonal regeneration through the nerve. This measure was introduced to identify whether axon coiling was occurring in CP nerves treated with GDNF at the repair site. Axonal profiles were counted at three points in the CP nerve: at the suture site, and at the distal stump at two locations, proximally and distally (Figure 3-15). Axon profiles crossing a perpendicular line in the middle of the field in each section were counted by a blinded observer (Kemp et al. 2009). We failed to observe a coiling effect in the nerves, with no significant differences found between treatment groups in the sections of the nerves analyzed (Figure 3-15). Neuronal Profiles # of NF Profiles Empty MS GDNF MS 0 SUT PROX DIST Location in Nerve CP nerve SUT PROX DIST Figure 3-15: Number of axonal profiles in the CP nerve Quantitative analysis of axonal profiles in the CP nerve at the suture site, the proximal end of the distal stump, and distally. No significant differences were found between the groups, indicating no axonal coiling effects in response to GDNF treatment.