Histological analyses are useful tools in PN regeneration research being an essential complement to the clinical, functional and electrophysiological investigation techniques (Vleggeert-Lankamp, 2007; Carriel et al., 2014b; Geuna, 2015). In this study, light microscopy and TEM histology were crucial to demonstrate an active PN regeneration process at the middle portion of all substitutes. However, differences in the amount, regenerative tissue distribution pattern and host tissue response to the implanted grafts were detected. Autograft group histology confirmed the presence of newly formed PN fascicles with moderate myelination (MCOLL and TEM results) along the connective tissue layers and especially, at the intrafascicular level. A comparable regeneration process was observed in NFABNS groups, although it was restricted to the connective tissue covering the implanted substitutes and not through the substitute layers. Surprisingly, tissue regeneration in filled conduits was considerably less abundant and poorly myelinated than the in autograft and NFABNS groups. In fact, the nerve tissue regeneration was restricted to the intraluminal area just around the intraluminal surface of the NFABNS. In summary, histology demonstrated that NFABNS – used alone or as intraluminal filler – did not promote nerve tissue regeneration through its biomaterials layers. Nevertheless, the direct use of NFABNS in PN repair resulted to be an efficient physical platform to keep both nerve stumps connected, supporting and guiding nerve tissue regeneration through its surrounding connective tissue, thus reaffirming the clinical, functional and EMG results.  In relation to the less favorable results obtained with filled NeuraGen R conduits, our histological analyses suggest that this finding could be related to physical factors (Daly W. et al., 2012; Carriel et al., 2014a). It is well accepted that nerve conduits provide a close and controlled protective microenvironment that guide and support nerve tissue regeneration in non-critical nerve gaps (Daly W. et al., 2012; Carriel et al., 2013, 2014a; Krarup et al., 2017; Pedrosa et al., 2017; Ronchi et al., 2018). In this regard, we hypothesize that the combined use of an external conduit filled with a highly dense NFABNS may reduce the area needed for an optimal tissue regeneration, and this would explain the poor functional and electrophysiological recovery observed with this strategy. In fact, it was experimentally demonstrated that low-density intraluminal fillers enhance regeneration process and functional recovery (Daly W. et al., 2012; Daly W.T. et al., 2012; Lee et al., 2012; Carriel et al., 2013, 2014a, 2017c; Gu et al., 2014; Wieringa et al., 2018). On the other hand, highly dense and slowly degrading intraluminal fillers can reduce the area available for tissue regeneration, thus delaying or even inhibiting tissue regeneration (Labrador et al., 1998; Yao et al., 2010; Daly W. et al., 2012; Daly W.T. et al., 2012; Carriel et al., 2014a).

In relation to the host response to our biomaterials, histological analyses confirmed that NFABNS, used alone or as intraluminal fillers, were progressively biodegraded by a local inflammatory host response mainly consisting of macrophages. These findings are in agreement with previous studies where FAH-based substitutes were reabsorbed by a comparable biodegradation process in some weeks (Carriel et al., 2012, 2013; Fernandez-Valades-Gamez et al., 2016; Martin-Piedra et al., 2017). Besides, histology also confirmed that NeuraGen R conduits protect the intraluminal NFABNS from host tissue biodegradation. This could reduce the intraluminal area needed for an optimal tissue regeneration and would reaffirm the hypothesis discussed above.

Finally, the use of cellular systems in PN repair demonstrated to be an efficient alternative to increase regeneration and functional recovery (Carriel et al., 2013, 2014a, 2017c; Kappos et al., 2015; Gonzalez-Perez et al., 2018; Zhang and Rosen, 2018). In this study, autologous undifferentiated ADMSCs were used to functionalize the NFABNS due to their well-demonstrated positive impact on PN repair (Kalbermatten et al., 2008; Lopatina et al., 2011; Gu et al., 2014; Faroni et al., 2016; Carriel et al., 2017b). In this study, acceptable PN regeneration and functional recovery profiles were obtained with the direct use of NFABNS in PN repair. However, due to technical reasons, it was not possible to identify the cells implanted within the NFABNS. Therefore, we cannot directly attribute to the cells the positive effect on PN regeneration obtained with our NFABNS. Therefore, the fate and potential role of these cells following in vivo implantation remains unknown and should be determined in future studies. In this regard, previous studies suggested that ADMSCs could contribute to the regeneration process through their differentiation to a SCs-like phenotype, releasing essential neurotrophic factors and collaborating with the synthesis of an essential ECM (Salgado et al., 2010; Tomita et al., 2012; Carriel et al., 2013; Kappos et al., 2015; Faroni et al., 2016; Zhang and Rosen, 2018).

#### CONCLUSION

In conclusion, the present study suggests that NFABNS support a closely comparable functional recovery and tissue regeneration than autograft technique. Overall results suggest that NFABNS may have potential clinical usefulness in future clinical trials in humans. However, further research should confirm their positive impact on the reconstruction of critical nerve defects and should improve the properties of these bioartificial tissue substitutes.

#### AUTHOR CONTRIBUTIONS

fncel-12-00501 December 21, 2018 Time: 15:45 # 17

VC, MA, and AC designed the experiments. VC, JC-A, and FC wrote the article. VC, JC-A, FC, OR, and DD-H performed the laboratory analyses. VC, JC-A, EM, JS-M, SG-G, and OR performed the functional and clinical analyses. VC, JC-A, FC, MA, and EM analyzed the results.

#### FUNDING

This study was supported by the Spanish "Plan Nacional de Investigación Científica, Desarrollo e Innovación Tecnológica," from the National Ministry of Economy and Competitiveness

#### REFERENCES

(Instituto de Salud Carlos III), Grants Nos FIS PI14-1343 and FIS PI17-0393, co-financed by "Fondo Europeo de Desarrollo Regional (FEDER)," European Union.

#### ACKNOWLEDGMENTS

The authors are grateful to Dr. Ariane Ruyffelaert (Faculty of Philosophy and Letters, University of Granada) for her proofreading service. The authors are grateful for the technical assistance of Amalia de la Rosa Romero, Concepción López Rodríguez, Dr. Víctor Domingo Roa (Experimental Unit of the University Hospital Virgen de las Nieves, Granada, Spain) and Leen Pieters (Ghent University, Belgium).

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

The handling Editor declared a past co-authorship with several of the authors VC, AC, and MA.

Copyright © 2018 Chato-Astrain, Campos, Roda, Miralles, Durand-Herrera, Sáez-Moreno, García-García, Alaminos, Campos and Carriel. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Long-Term Denervated Rat Schwann Cells Retain Their Capacity to Proliferate and to Myelinate Axons in vitro

Tessa Gordon1 \*, Patrick Wood2 and Olawale A. R. Sulaiman3

1Division of Neuroscience, Faculty of Medicine, University of Alberta, Edmonton, AB, Canada, 2The Miami Project to Cure Paralysis/Department of Neurological Surgery, University of Miami School of Medicine, Miami, FL, United States, 3Department of Neurosurgery, Ochsner Medical Center, New Orleans, LA, United States

Functional recovery is poor after peripheral nerve injury and delayed surgical repair or when nerves must regenerate over long distances to reinnervate distant targets. A reduced capacity of Schwann cells (SCs) in chronically denervated distal nerve stumps to support and interact with regenerating axons may account for the poor outcome. In an in vitro system, we examined the capacity of adult, long-term denervated rat SCs to proliferate and to myelinate neurites in co-cultures with fetal dorsal root ganglion (DRG) neurons. Non-neuronal cells were counted immediately after their isolation from the distal sciatic nerve stumps that were subjected to acute denervation of 7 days or chronic denervation of either 7 weeks or 17 months. Thereafter, equal numbers of the non-neural cells were co-cultured with purified dissociated DRG neurons for 5 days. The co-cultures were then treated with 3H-Thymidine for 24 h to quantitate SC proliferation with S100 immunostaining and autoradiography. After a 24-day period of co-culture, Sudan Black staining was used to visualize and count myelin segments that were elaborated around DRG neurites by the SCs. Isolated non-neural cells from 7-week chronically denervated nerve stumps increased 2.5-fold in number compared to ∼2 million in 7 day acutely denervated stumps. There were only <0.2 million cells in the 17-week chronically denervated stumps. Nonetheless, these chronically denervated SCs maintained their proliferative capacity although the capacity was reduced to 30% in the 17-month chronically denervated distal nerve stumps. Moreover, the chronically denervated SCs retained their capacity to myelinate DRG neurites: there was extensive myelination of the neurites by the acutely and chronically denervated SCs after 24 days co-culture. There were no significant differences in the extent of myelination. We conclude that the low numbers of surviving SCs in chronically denervated distal nerve stumps retain their ability to respond to axonal signals to divide and to elaborate myelin. However, their low numbers consequent to their poor survival and their reduced capacity to proliferate account, at least in part, for the poor functional recovery after delayed surgical repair of injured nerve and/or the repair of injured nerves far from their target organs.

#### Keywords: Schwann cells, nerve transection, myelination, SC proliferation, neurites, nerve injury

#### Edited by:

Giovanna Gambarotta, Università di Torino, Italy

#### Reviewed by:

Xavier Navarro, Autonomous University of Barcelona, Spain Valerio Magnaghi, University of Milan, Italy

#### \*Correspondence:

Tessa Gordon<EMAIL_ADDRESS>
Received: 18 September 2018 Accepted: 10 December 2018 Published: 07 January 2019

#### Citation:

Gordon T, Wood P and Sulaiman OAR (2019) Long-Term Denervated Rat Schwann Cells Retain Their Capacity to Proliferate and to Myelinate Axons in vitro. Front. Cell. Neurosci. 12:511. doi: 10.3389/fncel.2018.00511

### INTRODUCTION

The Schwann cells (SCs) in the peripheral nervous system support nerve regeneration after nerve injury in contrast to the oligodendrocytes in the central nervous system that do not (Cajal, 1928; Bunge, 1994; Fenrich and Gordon, 2004; Toy and Namgung, 2013). Yet, functional recovery is suboptimal after peripheral nerve injuries in patients who have sustained injury to large nerve trunks such as the brachial plexus (Sunderland, 1978; Kline and Hudson, 1995). The prevailing view is that this failure in recovery is due to the irreversible degeneration of chronically denervated muscles and their replacement with fat during nerve regeneration (Sunderland, 1978; Kline and Hudson, 1995). This is despite the early conclusions by Holmes and Young (1942) that there are ''various factors, in addition to atrophy of the end-organs, which are likely to reduce the effectiveness of recovery when suture is made after a long delay'' and the later conclusion that ''Most likely, multiple mechanisms contribute to this phenomenon'' (of diminished recovery of muscle mass and integrated motor function), ''which is sometimes referred to as 'irreversible denervation atrophy''' (Kobayashi et al., 1997). Peripheral nerves regenerate at 1–3 mm/day, resulting in delays of months and, in humans, even years before regenerating nerves reach their denervated targets (Fu and Gordon, 1997). Gutmann and Young (1944) reported that denervated rat neuromuscular junctions progressively deteriorate after delayed nerve surgery. The authors attributed poor functional recovery after such a delay to an irreversible inability of regenerating nerves to reinnervate chronically denervated neuromuscular junctions. However, more recent studies have demonstrated that the progressive failure of even freshly injured (axotomized) neurons to successfully regenerate their axons through chronically denervated distal nerve stumps is the major factor accounting for progressive decline in functional recovery (Fu and Gordon, 1995a; Vuorinen et al., 1995; Sulaiman and Gordon, 2000; Gordon et al., 2003, 2011) and the recovery of denervated muscle mass (Kobayashi et al., 1997). Chronically denervated muscles do accept reinnervation despite their declining satellite population and the consequent incomplete recovery of reinnervated muscle fiber size and muscle wet weight (Fu and Gordon, 1995a; Kobayashi et al., 1997; Sulaiman and Gordon, 2000; Jejurikar et al., 2002; Gordon et al., 2011). Hence, changes in the microenvironment of the chronically denervated nerve stumps provide, at least in part, the explanation for the progressive failure of axonal regeneration through the nerve stumps and to the denervated muscles.

The specific changes in the microenvironment of the denervated distal nerve stump accounting for poor axonal regeneration, are not understood. Regeneration is fostered in acutely injured peripheral nerves by the functional, structural, and molecular changes occurring in the denervated SCs in the distal stumps. Denervated SCs take an active role in the phagocytosis of myelin, a role that is taken over by macrophages that infiltrate the denervated nerve stump after 3 days (Gibson, 1979; Avellino et al., 1995; reviewed by Gordon, 2015). The SCs proliferate and extend long processes across the injury site within the first 10 days after injury (Salonen et al., 1988; Son and Thompson, 1995; Witzel et al., 2005). They also participate in the development of reversible and organized endoneurial structures that include the synthesis and positioning of collagen fibrils close to the laminin-containing basal lamina of the SCs and the formation of fascicle-like structures by the endoneurial cells (Salonen et al., 1987a,b, 1988).

The early changes in the denervated SCs include the switch in their phenotype from myelinating to a growth supportive one, the SCs downregulating genes that transcribe myelin associated proteins and glycoproteins and upregulating hundreds of growth-supportive genes reciprocally (De Leon et al., 1991; El Soury et al., 2018). The latter include genes that are responsible for ribonuclear RNA metabolic processes which contribute to the synthesis and production of new proteins and molecules required for Wallerian degeneration and nerve regeneration (El Soury et al., 2018). The many growth associated genes include those that transcribe transcription factors, c-jun and Notch, neurotrophic factors (reviewed by Jessen and Mirsky, 2016; Gordon and Borschel, 2017), soluble neuregulin 1 (NRG1) type I and type II isoforms, the NRG1 receptor ErbB2/3 (Carroll et al., 1997; Audisio et al., 2008; Gambarotta et al., 2013; Ronchi et al., 2016), and Erbin, an ErB2 interacting protein required for remyelination (Liang et al., 2012). The neurotrophic factors include nerve growth factor (NGF), brainand glial-derived neurotrophic factors, and pleiotrophin and the receptors include p75 and truncated trk receptors for the neurotrophic factors (Meyer et al., 1992; Funakoshi et al., 1993; Naveilhan et al., 1997; Hoke et al., 2002, 2006; Mi et al., 2007; Brushart et al., 2013). These early changes in the denervated SC phenotype are important in promoting axonal regeneration (Fu and Gordon, 1997; Boyd and Gordon, 2003; El Soury et al., 2018).

The decline in the capacity of chronically denervated SCs to support nerve regeneration is, at least in part, explained by the transient nature of neurotrophic factor and receptor upregulation (Boyd and Gordon, 2003; Gordon, 2015). Their upregulated gene expression declines within a month of denervation (Fu and Gordon, 1997; You et al., 1997; Hoke et al., 2002, 2006; Brushart et al., 2013), as does the upregulation of the erbB2/3 receptors (Li et al., 1997; Hall, 1999) required for the proliferation of denervated SCs (Carroll et al., 1997; Gambarotta et al., 2013). Importantly, the chronically denervated SCs atrophy and decline in numbers: progressive SC atrophy and loss with disruption of their basement membranes and shrinkage of their columns, have been reported in rat and rabbit hindlimb nerves over periods of 1 to 26 months (Weinberg and Spencer, 1978; Salonen et al., 1987a,b; Röyttä and Salonen, 1988; You et al., 1997; Bradley et al., 1998). SC numbers identified with S-100 immunoreactivity, declined from a peak of ∼3.5 times that of normal nerves at 4 weeks to the relatively low levels seen in intact nerves by 30 and 50 weeks after chronic sciatic nerve stump denervation in adult Wistar rats (Salonen et al., 1988).

In the present study, we evaluated the long-term survival of chronically denervated SCs and their capacities for proliferation and for myelination in response to contact with neurites. We provide the first quantitative evaluation of survival of very long-term (17 months) survival of chronically denervated SCs and demonstrate their sustained capacity to proliferate and to myelinate axons.

### MATERIALS AND METHODS

#### Ethical Approval

All surgical procedures and perioperative care measures were performed with strict accordance with the National Institutes of Health guidelines. All the procedures were reviewed and approved by the University of Alberta animal care committee following the Canadian Council of Animal Care (CCAC) guidelines. Rat embryos from pregnant females were removed on the 15th day of gestation for dissection of dorsal root ganglia (DRGs) for isolation of the DRG sensory neurons. All the procedures of removal and isolation of DRG neurons were carried out in Miami and were reviewed and approved by the Miami Animal Care and Use Committee.

### Animals

Adult female Sprague-Dawley rats (180–200 g body weight, n = 20) were used in the experiments in which hindlimb nerves were transected and ligated in vivo prior to removal of the denervated distal nerve stumps. Pregnant rats (n = 2) were used to remove 8–14 embryos each on the 15th day of gestation.

#### Materials

Cell culture reagents were purchased: Dulbecco's Modified Eagle Medium (DMEM) and Neurobasal medium and B27 supplement from Gibco, 0.05% collagenase from Worthington, 0.25% dispase from Boehringer-Mannheim, fetal bovine serum (FBS) from Hyclone, partially purified NGF from ProspectBio, trypsin from Worthington, TRL3, NTB2 emulsion from Kodak, mouse laminin from Collaborative Research, Inc., and rabbit anti-S100 antibody from Dako Corporation.

#### Anesthetic Protocols, Nerve Surgeries and Monitoring

All surgical procedures of sciatic nerve transection and ligation and the dissection of DRGs were made under anesthesia and using sterile surgical technique. Surgical anesthesia of the rats was induced with an intraperoneal injection of sodium pentobarbital (30 mg/kg).

#### In vivo Sciatic Nerve Transection of the Sciatic Nerves and Ligation of the Nerve Stumps

The left hindlimb was shaved and a lateral skin incision was made. The intermuscular septum between the biceps femoris and the vastus lateralis muscles was divided and the sciatic nerve identified. The sciatic nerve was freed using blunt dissection in order to cut the nerve sharply with scissors. The proximal and distal nerve stumps were ligated with 4.0 silk and sutured

to nearby innervated muscles in order to prevent reconnection and subsequent axonal regeneration (**Figure 1**), as previously described (Fu and Gordon, 1995a,b; Sulaiman and Gordon, 2000; Gordon et al., 2011). The overlying muscle was closed and the skin incision on the thigh was closed in layers. The rats were placed under heat lamps and their recovery monitored.

 Again, the nerve stumps were sutured to innervated muscle in order to prevent regeneration for 7 days (**Figure 1B**).

#### Monitoring for Evidence of Nerve Regeneration

In a third aseptic surgical procedure, the skin was shaved on both hindlimbs and then opened to expose, bilaterally, the sciatic nerve and it's tibial (TIB), common peroneal (CP) and sural nerve branches. An aseptic bipolar electrode attached to a Grass stimulator was used to deliver supramaximal stimuli at 10 Hz to the proximal sciatic nerve stump, followed by the distal stumps of the sciatic nerve and its branches to ensure that nerve regeneration had not occurred. No muscle contractions were observed in any of the rats confirming that the distal nerve stumps of the sciatic nerves were acutely denervated for 7 days and chronically denervated for 7 weeks and 17 months.

#### Surgical Removal of Sciatic, TIB, CP, and Sural Distal Nerve Stumps

In the same third surgical procedure, the chronically denervated (experimental) and the contralateral acutely denervated (control) distal stumps of the sciatic nerve and its branches in the left and right hindlimbs, respectively, were removed and placed in Belzer's solution at 4◦C for shipment to Miami for nerve dissociation, isolation of SCs, and preparation of co-cultures with DRG neurons.

#### Dissection of Dorsal Root Ganglia (DRGs) From Rat Embryos

Pregnant rats were deeply anesthetized with ether to allow surgery to remove embryos on the 15th day of gestation as previously described in detail (Kleitman et al., 1988). The DRGs from each of the 8–14 embryos were carefully dissected free and dissociated to establish DRG sensory neuron cultures (see below). The mothers were euthanized by exsanguination.

#### In vitro Cellular Dissociations and Co-culture

Non-neuronal Cells From Denervated Nerve Stumps Using sterile procedure, the epineurium was removed from the dissected distal nerve stumps of the sciatic nerve and the CP, TIB and sural nerve branches which had been chronically denervated for 7 weeks and 17 months (in the left experimental hindlimb or acutely denervated in the right control hindlimb for 7 days. Because the chronically denervated nerves were shrunken, especially those after the lengthy 17-month period of chronic denervation, the nerves for each of the three periods of nerve stump denervation were pooled for dissociation of their denervated non-neuronal cells, i.e., 10 nerves each were pooled after 7 weeks and 17 months of chronic distal nerve stump denervation and 20 nerves were pooled after 7 days of acute distal nerve stump denervation.

The nerves were incubated in culture medium containing 0.05% collagenase, 0.25% dispase solution in DMEM, and 10% FBS (Medium 1), for 18 h at 35–37◦C. The fascicles were rinsed free of collagenase/dispase with DMEM + 10% FBS. They were then dissociated into a single cell suspension by gentle trituration, using a glass micropipette with a tip diameter of 0.3–0.5 mm. The non-neuronal cells were harvested and re-suspended in Neurobasal medium with 2% B27 supplement and partially purified NGF at 50 units/ml (hereafter referred to as Medium 2). The cells were then counted and thereafter, their capacities to proliferate and to myelinate isolated DRG neurons were determined.

#### DRG Neurons From Rat Embryos

DRGs dissected from rat embryos on the 15th day of gestation were sequentially: (1) incubated with 0.25% trypsin in calcium and magnesium—free Hank's Balanced Salt Solution (Medium 3) for 1 h at 35–37◦C on a slowly rotating shaker; (2) rinsed free of trypsin with L-15 medium containing 10% heat-inactivated FBS; and (3) dissociated by gentle trituration using a glass pipette with a tip diameter of 0.3–0.5 mm. The dissociated DRG cells were then harvested and re-suspended in Medium 2. The DRG cell suspension was plated at a density of 20,000 cells in a volume of two drops onto collagen-coated clear coverslips, 22 mm in diameter (Kleitman et al., 1988) and placed into 25 mm clear mini-dishes that were incubated in a 5% CO2 atmosphere at 37◦C.

A day after plating, the DRG neuron cultures were flooded with 10 drops of the Medium 2 that was supplemented with fluorodeoxyuridine (FdU) and uridine, both at a concentration of 10 µM (antimitotic Medium 2). This antimitotic treatment was repeated on days 8 and 15 of culture. Thereafter, the antimitotic agents were excluded from the culture medium (Medium 2), the Medium 2 (without FdU and uridine) being used to wash out the antimitotic agents (on days 19, 21, 26, 28, and 33, and 42). This washing ensured the depletion of antimitotic agents from the culture environment of the DRG neurons prior to their co-culture with non-neural cells from the acutely and chronically denervated distal nerve stumps.

#### Co-culture of DRG Neurons and Non-neuronal Cells From Denervated Nerve Stumps

The DRG neurons in each mini-dish were plated with 100,000 non-neuronal cells on the same day as the dissociation of the non-neuronal cells from the chronically (7 weeks and 17 months) and acutely (7 days) denervated distal nerve stumps.

### Experiments

#### Cellular Counts

Small aliquots of non-neuronal cell suspensions, prepared in Medium 2 of each of the 7-week and 17-month chronically denervated nerve stumps and from the 7-day acutely denervated distal nerve stumps, were fixed in 4% paraformaldehyde in 0.05 M sodium phosphate buffer, pH 7.4 for 15 min at room temperature and stained with Hoechst 33342 dye (10 µM). The cells were visualized by fluorescence microscopy, counted in a hemocytometer, and the total cell numbers calculated. The resulting bright staining of cell nuclei allowed the unambiguous recognition of the cells even when the suspensions were heavily laden with myelin debris.

#### Cellular Proliferation Assay

The non-neuronal cell suspensions were diluted with Medium 2 to 100,000 cells/ml and plated onto cultures of isolated DRG neurons at 1.0 ml per culture. Control neuronal cultures received 1.0 ml of Medium 2 without suspended cells. For maintenance, the co-cultures were re-fed three times each week with Medium 2.

After 5 days of co-culture, the non-neuronal cells in each of four cultures from the 7-day acutely denervated control group and the 7-week and 17-month chronic denervation experimental groups were labeled for 24 h with 3H-thymidine (0.5 µCi/ ml) in Medium 2. The cultures were then rinsed and fixed with 4% paraformaldehyde in 0.1 M sodium phosphate buffer, pH 7.4 for 10 min at room temperature, permeabilized with 0.2% Triton X-100 in the fixative for another 10 min at room temperature, and prepared for immunostaining by blocking (30 s at room temperature) in a solution of 50% heat-inactivated goat serum in L-15 medium. The co-cultures were treated with rabbit anti-S100 antibody at 1:100 dilution in L-15/10% heat-inactivated goat serum for 30 min at room temperature, rinsed, and treated with fluorescein-conjugated goat anti-rabbit secondary antibody at 1:100 dilution in L-15/10% heat-inactivated goat serum for an additional 30 min at room temperature. The cultures were rinsed with buffer, distilled water and ethanol, dried and mounted on glass slides, cell side up, with DPX mountant. The slides were dipped in NTB2 emulsion and processed for autoradiography (Wood and Bunge, 1975). The counts of the S-100 positive cells were made directly under microscopic view and not made on photographic images with their possible distortions. A proliferation index was defined as the number of the S-100 positive SCs that were labeled with 3H-thymidine expressed as a percentage of the total number of S-100 positive SCs.

#### Myelination Assay

Six co-cultures per group of DRG neurons and non-neuronal cells isolated from the chronically denervated sciatic, TIB, CP, and sural nerve stumps, were maintained for 24 days in Medium 2 that was supplemented with 10 ng/ml of mouse laminin. Control cultures of DRG neurons without added non-neuronal cells were kept in parallel. All cultures were fixed overnight in FIX at 4◦C, osmicated for 1 h in 0.1% osmium tetroxide in 0.1 M sodium phosphate buffer, pH 7.4, dehydrated to 70% ethanol in water and stained for 1 h in 0.7% Sudan Black in 70% ethanol in water (Kleitman et al., 1988). The cultures were rehydrated and mounted in glycerin jelly on glass slides. A quantitative measurement of myelination in each culture was made by counting the number of myelin segments crossing the equator of the cultures, using an eyepiece containing a single line horizontally bisecting the field of view at 200× magnification.

#### Statistics

Statistical analysis was conducted using an ANOVA with the Tukey's post hoc test (confidence interval = 95%). Comparisons were made between the experimental chronically denervated (7 weeks and 17 months) distal nerve stump of the cut and ligated left sciatic nerve and the control acutely denervated (7 day) distal nerve stump of the cut and ligated right sciatic nerve (**Figure 1**).

#### RESULTS

#### The Number of Non-neuronal Cells in Peripheral Nerve Stumps Decline With Time After Chronic Denervation

The nuclei of the non-neuronal cells in suspension were unambiguously identified with Hoechst stain and their numbers counted in a hemocytometer to calculate their total cell numbers. The bright staining of cell nuclei allowed the unambiguous recognition of cells even when the suspensions were heavily laden with myelin debris.

The total numbers of non-neuronal cells from the experimental 7-week denervated distal nerve stump from the left hindlimb were >4 × 106 cells as compared to <2 × 106 cells in the acutely 7 day denervated distal stump from the right hindlimb in each rat (**Figure 2**). This increase was not sustained, the numbers after 17 months chronic nerve stump denervation falling to ∼10% of those numbers enumerated in the control 7-day acutely denervated distal nerve stumps. The increase of non-neuronal cells after chronic denervation at 7 weeks as compared to the numbers after acute denervation at 7 days, likely

FIGURE 2 | Decline in the number of non-neuronal cells and the proliferative capacity of S-100 positive SCs with chronic denervation. (A) Non-neuronal cells isolated from acutely (7 days) and chronically (7 weeks and 17 months) denervated distal nerve stumps of the sciatic nerve and its tibial (TIB), common peroneal (CP), and sural nerve branches, namely control acute and experimental chronically denervated distal nerve stump form the right and left hindlimbs, respectively. The cells were counted in small aliquots of single cell suspensions. (B) The mean (± SE) of the proliferation index. The index is the number of S-100 positive SCs that, in co-cultures with isolated dorsal root ganglion (DRG) neurons for 5 days, were labeled with 3H-thymidine and hence were dividing, expressed as a percentage of the total number of S-100 positive SCs. The proliferation indices at 7 days, 7 weeks and 17 months were significantly different from one another as indicated by \*'s (p < 0.025).

include fibroblast-like and mast cells that also accumulate in the denervated nerve stump (Salonen et al., 1988; Ronchi et al., 2017) in addition to proliferating SCs in the nerve stumps during the Wallerian degeneration period of ∼4 weeks (Salonen et al., 1988; You et al., 1997). The chronic denervation of the distal nerve stumps was controlled for the age and weight of the rats because the acutely and chronically denervated non-neuronal cells were dissociated from left and right hindlimbs, respectively, from the same rats.

#### Chronically Denervated Schwann Cells Retain Their Capacity to Proliferate, Albeit at a Reduced Rate

It is known that denervated SCs proliferate a second time when regenerating axons make contact with the SCs in the denervated distal nerve stump (Pellegrino and Spencer, 1985). Five days after co-culture of denervated SCs with isolated DRG neurons and a 24-h exposure to 3H-thymidine, most of the 7-day acutely denervated S-100 positive SCs were labeled with 3H-thymidine and hence, had entered into the cell cycle (**Figures 3A,D**). Some of these SCs are identified in the phase-contrast autoradiograph with an arrowhead (**Figure 3A**) whilst those that were not dividing are identified by an arrow (**Figure 3D**). The proportion of mitotic SCs in the co-cultures of DRG neurons and SCs from the acutely denervated nerves was high, the ratio of the numbers of 3H-thymidine labeled and unlabeled SCs, expressed as a percentage, the proliferative index in **Figure 2B**, being ∼75%.

The chronically denervated SCs from the left experimental hindlimb were also found to proliferate as shown in **Figures 3B–F** and quantitated in **Figure 2B**. However, the numbers of these 3H-thymidine positive SCs undergoing proliferation in the co-culture with DRG neurons declined significantly after chronic denervation (**Figures 3B–F**) with ∼50% and ∼30% of the SCs proliferating in the 7-week and 17-month chronically denervated nerves, respectively, the indices declining significantly with chronic denervation

FIGURE 3 | Many SCs in the nerve stump distal to the site of sciatic nerve transection and ligation are in an active proliferative state after acute as well as after chronic denervation, at least for up to 7 weeks. Phase-contrast autoradiographs of isolated acutely (A,D: 7 days) and chronically (B,E: 7 weeks and C,F: 17 months) denervated SCs from right and left hindlimbs, respectively. (A–C) Phase-contrast of the SCs that were exposed to 3H-thymidine for 24 h to identify dividing cells, 5 days after co-culture with DRG neurons. Many of the SCs incorporated 3H-thymidine (arrowhead) demonstrating SCs in an active proliferative state in the acute phase of denervation (A) and up to 7 weeks (B). There are visibly more 3H-Thymidine positive SCs than those that were not dividing (3H-Thymidine—negative cells shown by arrows) after 7 weeks (B) as compared to 7 days (A). (C,D) Immunostaining of SCs with S-100 identifies the dividing SCs (3H-thymidine positive SCs—arrows) and those that do not (3H-thymidine—negative cells shown by arrowheads). The proliferating S-100 positive SCs (arrows) are in close contact with the neurites of the DRG neurons. Scale bars = 30 µm.

(p < 0.05; **Figure 2B**). The differences in proliferative indices for the SCs from the control 1 week and experimental 7 weeks and 17 months were all significant (p < 0.05). Note that the S-100 immunostaining of the SCs in **Figure 3** revealed that the SCs were closely associated with and aligned along the neurites of the DRG neurons, irrespective of the duration of prior SC denervation in vivo.

### Chronically Denervated Schwann Cells Retain Their Capacity to Myelinate Axons

DRG neurons (without non-neuronal cells) were cultured for 24 days with and without non-neuronal cells that were freshly isolated from denervated nerve stumps of the sciatic nerve and its branches after acute (7 days) and chronic (7 weeks, 17 months) denervation in the right and left hindlimbs, respectively. In the absence of added non-neuronal cells, the DRG neurons exhibited extensive axon bundles that were not stained with Sudan Black (**Figures 4A,B**), a lipid stain widely used for detecting and distinguishing lipid-rich myelin sheaths in tissue culture (Eldridge et al., 1987). This result clearly indicates that myelin was not formed by any residual SCs that might not have been detected in the pure neuron cultures (**Figures 4A,B**). In contrast, in the cultures of DRG neurons to which non-neuronal cells (isolated from acute or chronically denervated sciatic nerve stumps) were added, many elongated Sudan black-positive

FIGURE 4 | Neurites extended from DRG sensory neurons in vitro become myelinated in co-cultures with chronically denervated SCs. Myelination of the axons is not evident in the absence of SCs but it proceeds normally in SC-DRG sensory neuronal co-cultures, even after chronic SC denervation. (A,B) DRG neurons (∗ ) maintained in culture for 24 days without SCs, were not myelinated as indicated by the arrows and seen by absence of Sudan Black staining in the unmyelinated axon bundles. Magnification was 20× (A) and 40× (B). (C–E) DRG axons were myelinated when DRG neurons were co-cultured with SCs whether or not the SCs were acutely denervated (C: 7 days) or chronically for 7 weeks (D,E) or 17 months by transecting and ligating the sciatic nerve in vivo in the right and left hindlimbs, respectively, prior to co-culture. Scale bar = 30 µm.

Gordon et al. Chronically Denervated Schwann Cells

profiles were observed (**Figures 4C–E**). These profiles exhibited the well characterized structure of myelin sheathes. These findings demonstrate that the SCs that remain in chronically denervated nerve stumps retain their capacity to form myelin sheaths around neurites.

Neurite myelination was quantitated by counting the myelin segments formed at 24 days after the co-culture of DRG neurons with the acutely and chronically denervated non-neuronal cells (see ''Materials and Methods'' section). In the assay, the non-neuronal cells that were isolated from chronically denervated sciatic nerve stumps from the left hindlimbs (i.e., the 7-week and 17-month groups) and co-cultured with DRG neurons, elaborated as many myelin segments as was elaborated by those cells isolated from the 7-day (acutely denervated) nerves from the right hindlimbs. The mean values (± standard deviations) were not significantly different (p > 0.05; **Figure 5**).

### DISCUSSION

This study demonstrated that first, SCs do survive in hindlimb nerve stumps that were chronically denervated for periods of up to 17 months. However, their numbers diminish dramatically. Second, the chronically denervated and atrophic SCs retain their abilities to associate with and to proliferate in response to contact with the neurites that extend from co-cultured DRG neurons. However, the rate at which the SCs re-enter the cell cycle is lower after a long period of denervation. Third, the chronically denervated SCs also sustain their capacity to myelinate the neurites extended in co-cultures of the cells with DRG neurons.

### Chronically Denervated SCs Atrophy and Many Die

Regeneration of injured peripheral nerves requires the presence of viable SCs in the denervated nerve stump (Hall, 1986).

Orphaned SCs normally proliferate in response to the mitogenic factor(s) provided by ingrowing axons (Wood and Bunge, 1975; Salzer and Bunge, 1980; Salzer et al., 1980a,b; Sabue et al., 1984; Pellegrino and Spencer, 1985; Stewart et al., 1991; Morrissey et al., 1995). When SCs are chronically denervated for extended periods of time, the SCs undergo atrophy and decline in number (Weinberg and Spencer, 1978; Röyttä and Salonen, 1988; Salonen et al., 1988; You et al., 1997; Bradley et al., 1998; Dedkov et al., 2002; Jonsson et al., 2013; Kumar et al., 2016). There is a concomitant decline in the capacity of injured neurons, both motor and sensory, to regenerate their axons (Fu and Gordon, 1995a; Gordon et al., 2011; Jonsson et al., 2013), and in the numbers of regenerated nerve fibers (Jonsson et al., 2013). In our present study, the numbers of non-neuronal cells doubled 7 weeks after chronic distal nerve stump denervation in the experimental left hindlimb as compared to the number in the control right hindlimb (**Figure 2A**) but the cells likely included mast cells and fibroblasts that accumulate in the denervated stump (Salonen et al., 1988; Ronchi et al., 2017) which, in turn, complete their phagocytic activities within 6–8 weeks prior to withdrawing their processes (Röyttä and Salonen, 1988). SC nuclei in the distal nerve stump increase ∼8-fold 25 days after a crush injury (Abercrombie and Johnson, 1946; Abercrombie and Santler, 1957) and to decline thereafter (Bradley and Asbury, 1970). Four weeks after chronic denervation, Salonen et al. (1988) reported an increase of 3.5-fold in SC number, suggesting that the number of proliferating SCs had already declined by 7 weeks. The macrophages that enter into distal nerve stumps within 3 days of denervation decline to low levels within a month (Gibson, 1979; Avellino et al., 1995) and therefore, are not likely to contribute to the non-neuronal cells that were enumerated after chronic denervation in the experimental nerves in the left hindlimbs (**Figure 2A**) nerve stumps in this study. That several non-neuronal cells enumerated after chronic denervation were likely to be SCs is supported by their S100 immunostaining (**Figures 3D–F**) after their co-culture with DRG neurons and hence, their exposure to endogenous mitogens on the surface of their neurites (Wood and Bunge, 1975; Salzer and Bunge, 1980; Salzer et al., 1980a,b; Sabue et al., 1984). These SCs demonstrated processes that were closely associated with and aligned along the neurites of the DRG neurons, irrespective of the duration of prior SC denervation in vivo.

The apparent discrepancy between the elevated non-neuronal cell numbers, of which most are likely to be SCs, and the reduced neuronal regenerative capacity at 7 weeks (Fu and Gordon, 1995a; Gordon et al., 2011) is likely due to the regression of the SCs to a non-supportive phenotype in which upregulated neurotrophic factors, including brain derived neurotrophic and glial derived neurotrophic factors, their p75 and GFRα-1 and GFRα-2 receptors, respectively, and soluble neuregulin 1 (Type I/II) and its Erb3/4 receptors, have declined back to low baseline values (Hoke et al., 2002, 2006; Boyd and Gordon, 2003; Gordon, 2015; Ronchi et al., 2017).

The non-neural cell numbers of >5 million at 7 days of acute nerve stump denervation in the right control hindlimb is dramatically lower, <2 million, in the right experimental hindlimb after 17 months of chronic stump denervation (**Figure 2A**), consistent with the progressive decline in numbers of viable SCs that were counted in rat hindlimb nerves after 3 and 6 months chronic denervation followed by 13 weeks nerve regeneration (Jonsson et al., 2013) and with the observed decline with light and electron microscopy, in 1–26-month chronically denervated rabbit nerves (Bradley et al., 1998), and 3–58-week chronically denervated rat nerves (Weinberg and Spencer, 1978). Microscopic observations demonstrated progressive Wallerian degeneration over the course of 7 weeks of chronic nerve stump denervation (You et al., 1997) with progressive disappearance of lipid-laden cells and residual myelin debris with cells of perineurial type (not fibroblasts) encircling small groups of denervated SC columns and progressive increase in the mass of collagen and elastin (Weinberg and Spencer, 1978; Bradley et al., 1998; see also Holmes and Young, 1942; Salonen et al., 1987a,b). The SC basement membranes progressively fragment, the shorter fragments dispersing throughout the endoneurium (Giannini and Dyck, 1990). It is likely that death of non-neuronal cells within chronically denervated nerve stumps explains the dramatic decline in their numbers. The nature of the death pathway is currently unknown and remains to be addressed.

#### Chronically Denervated SCs Proliferate but Less so Than Acutely Denervated SCs

Despite high number of non-neuronal cells in the 7-week chronically denervated distal nerve stumps in the experimental left hindlimb, the S-100 positive SCs were less able than acutely denervated SCs in the control right hindlimb to proliferate in response to endogenous mitogens released by 5-day co-cultured DRG neurons (**Figure 2B**). Reduced capacity of p75-positive SCs to proliferate was reported by Kumar et al. (2016). At 1 week, ∼75% of the acutely denervated SCs proliferated (**Figure 2B**), after which the proliferative capacity fell to ∼50% and ∼30% 7 weeks and 17 months after chronic denervation, respectively.

These findings of reduced proliferative capacity of chronically denervated SCs in the left hindlimbs contrast with earlier findings that SC proliferation was not altered by prior chronic denervation when the proliferation was evaluated in reinnervated nerve stumps, 13 weeks after surgical repair of 1–6-month chronically injured sciatic nerve stumps via an autograft (Jonsson et al., 2013). It must be recognized though, that, once the epineurium was removed from the excised distal nerve stump in that study, the ∼1 mm cut nerve pieces were incubated in a SC growth medium for 2 weeks and that the medium was supplemented by the exogenous mitogens of forskolin and neuregulin NRG1 prior to the isolation and incubation of the SCs for 7 days (see also Kumar et al., 2016). That the proliferative capacity of the SCs within the 13-week reinnervated nerve stumps was the same after this procedure, irrespective of the period of acute or chronic SC denervation that preceded the 13 weeks of nerve regeneration (Jonsson et al., 2013), likely reflected a renewed capacity of the SCs contained within the distal stump to proliferate normally in response to regenerating axons. The demonstration by Casella et al. (1996) of significantly enhanced proliferation of isolated human SCs after their exposure to the same mitogens immediately after their isolation, as compared to that of the SCs isolated from untreated control nerves, provides further evidence that the proliferative properties of isolated SCs are altered after isolation and culture in media containing exogenous mitogens. In our experiments, proliferation of the SCs was analyzed only after the cells were exposed to the specific SC mitogens that are present on the surface of DRG neurites in the 5-day SC-DRG co-cultures (Wood and Bunge, 1975; Salzer and Bunge, 1980; Salzer et al., 1980a,b).

In the proliferation assay that we used, the isolated non-neuronal cells were labeled for 24 h with 3H-thymidine after SC-DRG co-culture for 5 days. The cells were then fixed and the dividing SCs identified as p75 immunoreactive (**Figure 3**). Therefore, the proliferation index measures the occurrence of new DNA synthesis and thereby, determines the capacity of the denervated SCs to enter into the cell cycle in response to the endogenous mitogens on the DRG neurons. SCs, whether immediately reinnervated or after a delay of 1–6 months, require time to re-enter the G1 phase of proliferation because significantly more reinnervated SCs proliferated at 5 as compared to 3 days after a 2-week period of incubation (Jonsson et al., 2013). Nonetheless, the reduced proliferation of chronically denervated SCs during the early phase of proliferation argue that the slower rate is unlikely to account for the findings.

The vigorous proliferation that we observed in acutely denervated SCs from the control right hindlimbs (**Figure 2B**) is consistent with the spontaneous upregulation of their erbB receptors following nerve transection (Carroll et al., 1997; Fricker and Bennett, 2011; Fricker et al., 2011; Chang et al., 2013) whilst their reduced proliferation with chronic denervation is likely linked to their downregulation of erbB2/erbB3 receptors (Li et al., 1997; Hall, 1999; Jonsson et al., 2013; Ronchi et al., 2017). The retained, albeit reduced, capacity of chronically denervated SCs to proliferate in vitro (**Figure 2B**) and for 5%–10% of freshly injured neurons to regenerate their axons through chronically denervated nerve stumps (Fu and Gordon, 1995a; Sulaiman and Gordon, 2000; Gordon et al., 2011), elaborate normal myelin sheaths, and to recover their normal size once target contacts are remade (Sulaiman and Gordon, 2000), demonstrate that the proliferative quiescence of the chronically denervated SCs from the experimental left hindlimbs is reversible: the dormant SCs re-enter the cell cycle and proliferate (**Figures 2B**, **3**), albeit at a more sluggish initial rate.

#### Chronically Denervated SCs Remyelinate Neurites

Sudan Black staining of the SCs isolated from chronically denervated distal nerve stumps of the experimental left hindlimbs and co-cultured with DRG neurons for 24 days, demonstrated normal capacity to remyelinate the neurites elaborated by the neurons (**Figures 4**, **5**). Sudan Black staining of the lipid-rich membranes of compacted myelin sheaths was chosen to demonstrate remyelination of neurites rather than immunostaining because SC immunostaining without Sudan Black staining reveals SCs that are making myelin proteins but are not necessarily making myelin sheathes. SC immunostaining thereby, could give the erroneous impression that the SCs were making myelin sheaths when they were not. The in vitro findings of neurite myelination reported here, corroborate our in vivo findings of the remyelination of the low numbers of axons that injured neurons regenerate into chronically denervated nerve stumps (Sulaiman and Gordon, 2000). There were no significant differences in the extent of myelination by acutely or chronically denervated SCs derived from the right and left hindlimbs, respectively, whether the duration of chronic denervation was 7 weeks or 17 months (**Figure 5**). 1% to 5% of the DRG neurites were well myelinated, irrespective of the duration of the SC denervation in vivo prior to their co-culture, the remaining neurites being <1 µm in diameter and hence, unmyelinated. This ''normal'' myelinating capacity of chronically denervated SCs contrasts with the ''severe impairment'' of the myelination of the neurites of DRG neurons by 56-day denervated SCs after the non-neural passaging in the presence of mitogens and subsequent cell sorting (Kumar et al., 2016). This reported ''severe impairment'' however, does not correspond with the capacity of chronically denervated SCs to myelinate regenerated axons with restoration of the normal proportional relationship between myelin thickness and the size of the myelinated nerves (Sulaiman and Gordon, 2000).

#### Summary

The survival and sustained capacity of chronically denervated SCs to re-enter the cell cycle together with the sustained capacity for these SCs to form myelin around neurites in vitro and regenerating axons in vivo indicates that surviving SCs may be stimulated to divide and support axon regeneration and re-myelination after chronic nerve injuries. Overall our results

#### REFERENCES

suggest that failure of axons to regenerate into long denervated nerve trunks is not due to lack of ability of the SCs to interact or respond to the axons. Rather, it is more likely to result from the reduced numbers of chronically injured neurons to regenerate their axons (Fu and Gordon, 1995b) and the extremely small number of SCs available to support and remyelinate the few regenerating axons in chronically denervated nerve stumps that and support nerve regeneration (Fu and Gordon, 1995a; Sulaiman and Gordon, 2000; Gordon et al., 2011). As such, cellular therapy may be a clinically viable option for patients who have sustained injuries to large nerve trunks.

### AUTHOR CONTRIBUTIONS

All authors made equal contributions to the data interpretation and writing of the manuscript. PW and OS made the major contributions to the experimental execution.

### FUNDING

We are grateful for funding from the Canadian Institutes of Health Research (CIHR).

#### ACKNOWLEDGMENTS

Our thanks to our colleague Ernesto Felipe-Cuervo for his help with the experiments in Dr. Patrick Wood's laboratory. We are also very grateful to the late Dr. Richard Bunge for his contribution to this manuscript as well as to his many articles during his imminent scientific career. Our thanks to Dr. Doan Nguyen for his assistance with the figures for the manuscript.

schwann cells. Front. Mol. Neurosci. 11:157. doi: 10.3389/fnmol.2018. 00157

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Gordon, Wood and Sulaiman. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# The Role of BDNF in Peripheral Nerve Regeneration: Activity-Dependent Treatments and Val66Met

Claire Emma McGregor\* and Arthur W. English

Department of Cell Biology, Emory University School of Medicine, Atlanta, GA, United States

Despite the ability of peripheral nerves to spontaneously regenerate after injury, recovery is generally very poor. The neurotrophins have emerged as an important modulator of axon regeneration, particularly brain derived neurotrophic factor (BDNF). BDNF regulation and signaling, as well as its role in activity-dependent treatments including electrical stimulation, exercise, and optogenetic stimulation are discussed here. The importance of a single nucleotide polymorphism in the BDNF gene, Val66Met, which is present in 30% of the human population and may hinder the efficacy of these treatments in enhancing regeneration after injury is considered. Preliminary data are presented on the effectiveness of one such activity-dependent treatment, electrical stimulation, in enhancing axon regeneration in mice expressing the met allele of the Val66Met polymorphism.

#### Edited by:

Esther Udina, Autonomous University of Barcelona, Spain

#### Reviewed by:

Doychin N. Angelov, Medizinische Fakultät, Universität zu Köln, Germany Guilherme Lucas, University of São Paulo, Brazil

#### \*Correspondence:

Claire Emma McGregor<EMAIL_ADDRESS>
Received: 26 September 2018 Accepted: 14 December 2018 Published: 11 January 2019

#### Citation:

McGregor CE and English AW (2019) The Role of BDNF in Peripheral Nerve Regeneration: Activity-Dependent Treatments and Val66Met. Front. Cell. Neurosci. 12:522. doi: 10.3389/fncel.2018.00522 Keywords: peripheral nerve injury, BDNF, Val66Met, trkB, electrical stimulation, exercise, optogenetics

#### PERIPHERAL NERVE INJURY

Despite the ability of axons in peripheral nerves to regenerate, recovery is generally very poor (Portincasa et al., 2007; Scholz et al., 2009). The cellular changes that occur after an injury often cannot sustain axon regeneration for the duration required to reinnervate target organs (Fu and Gordon, 1995a,b). The neurotrophins have emerged as an important modulator of axon regeneration, particularly brain derived neurotrophic factor (BDNF). Here, we will review BDNF and its role in activity-dependent treatments to enhance regeneration. Then we will discuss a single nucleotide polymorphism in the bdnf gene, Val66Met, which is present in 30% of the human population and may hinder the efficacy of these treatments (Egan et al., 2003; Shimizu et al., 2004). Finally, we will present preliminary data on the effectiveness of one such activity-dependent treatment, electrical stimulation (ES), in enhancing axon regeneration in mice expressing the met allele of the Val66Met polymorphism.

### BRAIN DERIVED NEUROTROPHIC FACTOR

BDNF is a member of the neurotrophin family, which also includes nerve growth factor (NGF), neurotrophin 3 (NT3), and neurotrophin 4/5 (NT4/5). BDNF is required for normal development—BDNF knockout (KO) is embryonic lethal (Jones et al., 1994; Schwartz et al., 1997). In adulthood, BDNF is involved in synaptic plasticity, long term potentiation (LTP), learning and memory as well as hippocampal neurogenesis and regeneration after injury (Lindsay, 1988; Lewin and Barde, 1996; Lu et al., 2014; Richner et al., 2014). In the subsequent paragraphs, we review how BDNF is regulated at the level of mRNA transcripts, protein trafficking, and receptor binding, following with its role in peripheral nerve regeneration.

#### Regulation of BDNF Transcripts

The human BDNF gene resides on the short arm of the 11th chromosome (Maisonpierre et al., 1991). It consists of 9 exons—eight 5′ untranslated exons and one protein coding 3′ exon (**Figure 1**) (Liu et al., 2006; Aid et al., 2007; Pruunsild et al., 2007). Through alternative splicing, 17 distinct mRNA transcripts for BDNF have been identified in humans and 11 in rodents (Pruunsild et al., 2007). Additionally, the 3′UTR of the gene contains two polyadenylation sites, resulting in both a long 3′UTR and a short 3′UTR, doubling the possible splice variants. The entire protein-coding region resides on exon IX, so the mature BDNF protein synthesized is identical regardless of mRNA splicing. Splice variants allow for spatial and temporal control of the BDNF transcript.

Spatial control of the 5′UTRs can be seen in exon expression throughout the body. BDNF transcripts containing exons I, II, and III are found exclusively in the brain, and transcripts containing exon IV are predominantly found peripherally in the lung and heart, but can also be found in brain tissue (Timmusk et al., 1993). Even within brain tissue, different promoters can be found in different cell types. For example, exon IV transcripts are required for proper GABAergic interneuron function in the prefrontal cortex (Sakata et al., 2009).

Many different stimuli exert temporal control over BDNF transcription. In cultured cortical neurons, Ca2+ influx results predominantly in transcription of exon IV-containing mRNA (Tao et al., 1998). This promoter contains a cAMP/Ca2+ response element-like element (CaRE3/CRE) that is required for activity-dependent transcription (Tao et al., 1998; Hong et al., 2008). The transcription factor CREB binds this element, is phosphorylated by calcium-regulated kinase cascades, and recruits transcriptional machinery resulting in Ca2+ dependent transcription of exon IV-containing BDNF mRNA (West et al., 2001; Lonze and Ginty, 2002). Other stimuli have been identified in modulating BDNF expression. In motoneurons, exon VI transcripts are androgen sensitive, despite no known androgen response element on the bdnf gene (Ottem et al., 2010; Sabatier and English, 2015). There is, however, an estrogen response element (Sohrabji et al., 1995). SRY-box containing gene 11 (Sox11), a transcription factor involved in neuronal survival, axon growth, and regeneration after injury, increases exon I containing BDNF mRNA transcripts specifically in peripheral DRG neurons, but not in CNS neurons (Jankowski et al., 2006; Salerno et al., 2012; Struebing et al., 2017). Exons II and VI are sensitive to tricyclic and atypical antidepressants (Vaghi et al., 2014).

A further role for 5′ promoter exons regulating BDNF mRNA may lie in mRNA trafficking. In both cortical and hippocampal neurons, BDNF mRNA is found in dendrites and activity induces trafficking of BDNF mRNA to distal dendrites (Tongiorgi et al., 1997; Capsoni et al., 1999; Chiaruttini et al., 2008, 2009). Interestingly, only certain splice variants are found in dendrites those containing exons IIB, IIC, and VI (Pattabiraman et al., 2005; Chiaruttini et al., 2008). Transcripts containing exons I, III, and IV are restricted to the cell body.

Further spatial and temporal translational control of BDNF mRNA may come via the 3′UTR. The bdnf 3 ′ UTR contains two polyadenylation sites. This allows for both a long 3′UTR and a short 3′UTR to be transcribed (Timmusk et al., 1993; Aid et al., 2007; Pruunsild et al., 2007). These different 3′UTRs are thought to determine mRNA trafficking within the cell. Both the long and short 3′UTR transcripts can be found in the dendrites under different conditions (Vicario et al., 2015). However, in general, the long 3 ′UTR transcripts are trafficked to the dendrites, where local BDNF synthesis can regulate pruning and enlargement of synapses, whereas the short 3′UTR transcripts stay in the cell body (An et al., 2008). Both depolarization of the neuron as well as BDNF itself increase the number of BDNF mRNA transcripts targeted to the dendrites (Tongiorgi et al., 1997; Righi et al., 2000). Remarkably, the short 3′UTR transcripts account for the majority of BDNF translation, whereas the long 3 ′UTR transcripts are translationally repressed by RNA binding proteins which stabilize the mRNA until neural activity elicits rapid local translation (Lau et al., 2010; Allen et al., 2013; Vaghi et al., 2014). Calcium influx associated with neuronal activity

also results in increased stabilization of BDNF mRNA (Fukuchi and Tsuda, 2010). Conversely, the 3′UTR also contains regions which interact with microRNAs, which are short, non-coding RNA strands that complement mRNA transcripts and result in transcript degradation (Bartel, 2004). Numerous microRNAs have been identified as regulators of BDNF, and microRNA access to binding sites depends on the presence of the long or short 3 ′UTR (Mellios et al., 2008; Varendi et al., 2014).

### BDNF Trafficking and Secretion

BDNF, like the other neurotrophins, is synthesized as a preproprotein. The pre-domain functions as a signaling peptide that directs synthesis to the endoplasmic reticulum (ER) for future packaging as a secretory protein. It is immediately cleaved to form proBDNF upon sequestration in the ER (Lessmann et al., 2003). Within the ER, proBDNF forms homodimers (Kolbeck et al., 1994). Both proBDNF and a further cleaved form, mature BDNF, can be packaged into vesicles and secreted. ProBDNF is approximately 29 kDa, and once cleaved, mature BDNF is approximately 14 kDa (Seidah et al., 1996a). Intracellularly, cleavage can occur within the trans-Golgi network or secretory vesicles by furin, a protease, and proprotein convertases PCSK6 and PC5-6b. Extracellularly, cleavage is executed by tissue plasminogen activator or matrix metalloproteinases, which are secreted in an activity-dependent manner (Krystosek and Seeds, 1981; Gualandris et al., 1996; Seidah et al., 1996a,b; Lee et al., 2001; Mowla et al., 2001; Hwang et al., 2005; Keifer et al., 2009; Nagappan et al., 2009; Yang et al., 2009). Once cleaved, the prodomain is not immediately degraded, and can be secreted with mature BDNF (Anastasia et al., 2013).

From the trans-Golgi network, BDNF is directed to two different secretory pathways: a constitutive pathway and a regulated, Ca2+ dependent pathway (Mowla et al., 1999; Lessmann et al., 2003; Kuczewski et al., 2009). The constitutive pathway consists of small granules (50–100 nm diameter) that fuse with the cell membrane near the neuronal somata and proximal processes. The regulated secretory pathway consists of larger granules (300 nm diameter) that fuse in distal processes and axon terminals (Conner et al., 1997; Kohara et al., 2001; Brigadski et al., 2005; Dieni et al., 2012). The dual pathway for release is distinctive of BDNF. The other neurotrophins are preferentially secreted through a constitutive pathway. Under normal conditions, most neuronal BDNF is packaged into the regulated pathway (Lu et al., 2014).

Two important interactions have been identified in the sorting of BDNF into the regulatory pathway. The first to be discovered was the interaction between the sorting receptor carboxipeptidase E (CPE) and a three-dimensional motif on the mature domain of BDNF (Lou et al., 2005). Knocking out CPE in cortical neurons blocks activity-dependent release of BDNF and increases constitutive release. Similarly, adding this motif to NGF redirected its release to the regulated secretory pathway (Lou et al., 2005). The second interaction is between the prodomain and the protein sortilin (Chen et al., 2005). Sortilin is localized predominantly to the Golgi apparatus and interacts with the BDNF prodomain to direct it to the regulated secretory pathway (Nielsen et al., 2001; Chen et al., 2005). When sortilin is unable to interact with the prodomain of BDNF, regulated release is decreased, but there is no compensatory increase in constitutive release (Chen et al., 2005; Lu et al., 2005). This has led to the hypothesis that the interaction between sortilin and the prodomain of BDNF is necessary for proper protein folding, which allows CPE to interact with the mature domain and sort BDNF into one of the two pathways (Lu et al., 2005). When NT-4/5, which is secreted constitutively, is modified to contain the BDNF prodomain, it is trafficked to the regulated secretory pathway (Brigadski et al., 2005). Similarly, blocking the cleavage of the prodomain of NGF, which is also released constitutively, results in its sorting into regulated secretory pathways (Mowla et al., 1999).

#### BDNF Receptors

Once secreted, BDNF can bind to one of two receptors tropomyosin receptor kinase B (trkB) or the common neurotrophin receptor, p75NTR. Mature BDNF preferentially binds trkB, resulting in pro-growth signaling, whereas proBDNF (as well as the other proneurotrophins) preferentially binds p75NTR, resulting in antigrowth signaling (Lee et al., 2001). BDNF is primarily secreted as proBDNF (Mowla et al., 1999, 2001; Chen et al., 2004). Thus the availability of proteins that cleave the prodomain may regulate which receptor is activated by BDNF release, providing another mechanism for control of BDNF signaling.

The trkB receptor is a typical tyrosine kinase. When ligand is bound, it dimerizes and autophosphorylates. In addition to BDNF, trkB can also bind NT-4/5. Several isoforms of trkB have been discovered, including isoforms that change its sensitivity to NT-4/5, as well as a truncated form that lacks an intracellular kinase domain (Eide et al., 1996). The truncated form acts as a dominant negative receptor, forming heterodimers with full length trkB receptors and blocking neurotrophin signaling (Eide et al., 1996; Fryer et al., 1997). Another possible role for truncated trkB on astrocytes and Schwann cells may be to act to control the pool of available neurotrophins, preventing them from degrading or signaling until released into the extracellular space (Alderson et al., 2000). In its fulllength form, trkB has several intracellular tyrosine residues that can be phosphorylated (Huang and Reichardt, 2003). Three possible signaling cascades are then activated: phospholipase C gamma (PLCG); phosphotidyl-inositol-3 kinase (PI3K); and mitogen activated protein kinase/extracellular receptor kinase (MAPK/ERK) (Reichardt, 2006).

The phosphorylation of residue Y490 creates a binding site for adaptor protein Shc (Patapoutian and Reichardt, 2001). Shc binding trkB allows for activation of Ras and further activation of the MAPK/ERK pathway. Downstream of this pathway is mechanistic target of rapamycin (mTOR). Shc binding residue Y490 also results in the recruitment of PI3K and activation of protein kinase B (Akt) (Reichardt, 2006). The phosphorylation of residue Y785 creates a binding site for PLCG, which is then phosphorylated by trkB (Patapoutian and Reichardt, 2001). This phosphorylation activates PLCG, which then hydrolizes phosphotidylinositides to generate diacylglyerol (DAG), which activates protein kinase C (PKC), and inositol 1,4,5 triphosphate (IP3), which results in an influx of intracellular Ca2+ stores from the ER. These signaling cascades all converge at the level of the nucleus, where transcription is affected through CREB and other transcription factors (Minichiello, 2009).

Once bound to ligand, trkB is endocytosed to form a signaling endosome (Delcroix et al., 2003; Reichardt, 2006). Both ligand and receptor are contained within the endosome, allowing trkB to continue signaling as it is trafficked through the cell. In this way, trkB can be moved closer to the nucleus, where it can affect gene transcription, as well as brought into closer proximity to signaling effectors (Delcroix et al., 2003). However, not all actions of trkB happen at the level of the soma—BDNF-trkB activation has been shown to affect local protein synthesis in the growth cone as well (Yao et al., 2006).

The second receptor for BDNF is the pan-neurotrophin receptor, p75NTR (Rodríguez-Tébar et al., 1992). Generally thought of as a pro-death receptor, p75NTR is a member of the tumor necrosis factor receptor super family and contains a cytosolic death domain (Liepinsh et al., 1997; Locksley et al., 2001). It is expressed primarily during development, but sensory neurons and spinal motoneurons maintain low expression through adulthood (Ernfors et al., 1989; Heuer et al., 1990; Wyatt et al., 1990; Ibáñez and Simi, 2012). Its cytosolic domain is nonenzymatic, so its actions depend entirely on associations with cytoplasmic proteins (Nagata, 1997). Despite its canonical role, p75NTR can mediate both pro-death and pro-survival signals depending on its cytosolic partners. For example, p75NTR is required during development for normal neuron growth and ramification (Yamashita et al., 1999). Multiple adaptor complexes interact with its cytosolic domain to mediate downstream effects (Dechant and Barde, 2002).

Additionally, p75NTR has multiple membrane-bound and extracellular binding partners which can alter whether its signaling is pro-survival or pro-death. Through extracellular pairing with sortilin, p75NTR is able to bind the proneurotrophins (Nykjaer et al., 2004; Teng et al., 2005), resulting in pro-death or anti-growth signaling through downstream JNK activation or caspase activation (Reichardt, 2006). However, neurotrophin binding to p75NTR can also result in NFκB activity, which is a pro-survival signal (Carter et al., 1996; Hamanoue et al., 1999; Middleton et al., 2000). One key protein regulated by p75NTR is RhoA, a small GTPase that regulates the actin cytoskeleton and inhibits axon elongation (Walsh et al., 1999; Schmidt and Hall, 2002). Through such interactions with the socalled death domain of p75NTR, neurotrophin binding inhibits Rho (Yamashita et al., 1999; Roux and Barker, 2002). Through forming a receptor complex with the Nogo receptor, NgR1, p75NTR can act as a receptor for myelin-associated glycoprotein (MAG) (Wang et al., 2002), which enhances Rho activation and results in neurite collapse (Mi et al., 2004). Curiously, p75NTR can act as a binding partner for the trks, including trkB, and increases affinity and selectivity of binding and thus enhancing trk signaling (Bibel et al., 1999).

Like trkB, truncated forms of p75NTR have been identified. One short p75NTR isoform lacks an extracellular ligand binding domain, but contains its intracellular machinery (Roux and Barker, 2002). This form is unable to bind the neurotrophins (Dechant and Barde, 1997). The extracellular domain of p75NTR can also be cleaved by extracellular metalloproteinases (Roux and Barker, 2002). These isoforms could act as modulators of neurotrophin signaling.

The two receptors for BDNF are generally thought to have opposing roles and may mediate a balance between growth and death. trkB has a higher affinity for mBDNF, but as levels of neurotrophin increase, p75NTR will also bind mBDNF and activate signals in direct opposition to trkB. Because of the different affinities for pro- and mature BDNF, cleavage of BDNF becomes another mechanism to control its downstream signaling effects (Lee et al., 2001). Depolarization of a neuron, which results in secretion of BDNF, also results in secretion of tissue plasminogen activator which cleaves proBDNF to create mature BDNF (Gualandris et al., 1996). BDNF-trkB signaling increases expression of matrix metalloproteinase 9, which also cleaves BDNF (Kuzniewska et al., 2013).

### ROLE OF BDNF IN PERIPHERAL NERVE INJURY

In peripheral nerves, BDNF is synthesized by motoneurons, a subset of DRG neurons, and Schwann cells (Apfel et al., 1996; Cho et al., 1997; Michael et al., 1997). After nerve crush or complete transection, BDNF mRNA increases in all three cell types, including in trkB- and trkC-expressing DRG neurons not found previously to express BDNF (Meyer et al., 1992; Funakoshi et al., 1993; Kobayashi et al., 1996; Michael et al., 1999; Al Majed et al., 2000a; English et al., 2007). BDNF mRNA can be found in low levels in the sciatic nerve, and after injury, that expression is upregulated. This upregulation is sustained over the course of weeks and can be attributed to both neuronal and non-neuronal sources (Meyer et al., 1992; Funakoshi et al., 1993). In facial nerve injury, upregulation of BDNF is correlated with enhanced functional outcome (Grosheva et al., 2016).

Following sciatic nerve injury, a transient increase in both BDNF and full length trkB mRNA is found in motoneurons (Kobayashi et al., 1996; Al Majed et al., 2000a). Unlike sensory neurons, NGF and trkA are not expressed by motoneurons, nor are they upregulated after injury (Funakoshi et al., 1993; Escandon et al., 1994). There is a small and short-lived upregulation of NT3 and NT4/5 in motoneurons (Funakoshi et al., 1993). TrkC is expressed by adult motoneurons, but it is not upregulated after injury (Johnson et al., 1999). Thus, the rapid upregulation of BDNF and trkB make it likely that BDNF is the main neurotrophin mediating early motoneuron response to nerve injury (Boyd and Gordon, 2003).

Schwann cells express only the truncated form of trkB, which has the potential to act as a dominant negative receptor for BDNF and NT4/5. Schwann cell truncated trkB mRNA levels decrease significantly after sciatic nerve injury (Frisén et al., 1993). This could be viewed as pro-regenerative, enabling available BDNF to bind to trkB receptors on regenerating neurites and enhance their growth. Conversely, after injury, Schwann cells upregulate p75NTR, which has been suggested to result in sequestration of neurotrophins and inhibit regeneration (Taniuchi et al., 1986; Bibel et al., 1999; Scott and Ramer, 2010).

#### trkB in Peripheral Nerve Injury

Axons regenerate through the formation of growth cones, which need cytoskeletal proteins, such as actin and tubulin, to extend and stabilize the new growth. Beta actin mRNA is localized to peripheral axons, and peripheral nerve injury triggers actin mRNA to be transported down the axon for local protein synthesis (Koenig et al., 2000; Sotelo-Silveira et al., 2008; Willis et al., 2011). BDNF/trkB signaling triggers local translation of transported mRNAs through a Ca2+-dependent mechanism, and this is required for bidirectional turning toward BDNF (Yao et al., 2006). BDNF application to injured axons increases the number of actin waves (transport of actin filaments and associated proteins toward the growth cone) per hour (Difato et al., 2011; Inagaki and Katsuno, 2017). Neurotrophins also stimulate growth cone sprouting and actin accumulation in the sprouts (Gallo and Letourneau, 1998). Both of these processes are mediated through the PIP3/PI3K signaling pathway described above (Asano et al., 2008). When an actin wave reaches the growth cone, the growth cone enlarges, branches, and undergoes forward expansion (Flynn et al., 2009). Application of BDNF to growth cones results in microtubule reorganization to form lamellipodial as well as filopodial elongation (Gibney and Zheng, 2003).

In addition to local protein synthesis, trkB signaling has effects on cyclic AMP (cAMP) production, which may be important for the initial extension of growth cones across the site of injury. The MAPK/ERK pathway of BDNF/trkB signaling results in inhibition of phosphodiesterases (PDE) which normally degrade cAMP (Gao et al., 2003). As such, BDNF/trkB signaling results in increased levels of cAMP (Souness et al., 2000; Gao et al., 2003). This pathway has been shown to be necessary to overcome inhibition by MAG, and therefore PDE inhibition has been most thoroughly studied in models of spinal cord injury, where MAG inhibition of axon growth creates a substantial barrier to regeneration (Cai et al., 1999; Gao et al., 2003; Batty et al., 2017). Although injured peripheral nerves do not suffer inhibition by MAG to the same extent as that seen in the central nervous system, early in the regeneration process, inhibitory proteoglycans and myelin debris form an impermissible environment for axon regeneration (shen et al., 1998). Increasing cAMP through PDE inhibition enhances peripheral regeneration after injury, and it is likely that trkB activation contributes to this cAMP-mediated effect on regeneration (Gordon et al., 2009; Udina et al., 2010).

The different neurotrophin signaling pathways activated through trk receptors converge at the level of transcription in the nucleus. CREB, resulting from trkB-generated PI3K-Akt activation, increases sensory neurite outgrowth (White et al., 2000). Inhibiting phosphatase and tensin homolog (PTEN), an endogenous inhibitor of the PI3K pathway, through genetic knock out or pharmacology, enhances peripheral nerve regeneration in vivo and neurite outgrowth in vitro (Park et al., 2008; Christie et al., 2010). Numerous other transcription factors downstream of MAPK/ERK signaling, such as c-jun, STAT3, and ATF-3, have all been associated with changes in gene expression after injury that enhance survival and regeneration (Makwana and Raivich, 2005). mTOR, also downstream of MAPK/ERK signaling and repressed by PTEN, regulates protein synthesis and is also beneficial for DRG regeneration after injury (Park et al., 2008; Abe et al., 2010).

### p75NTR in Peripheral Nerve Injury

Because of its roles in both pro-death and pro-survival signaling, it is not surprising that the role of p75NTR in regeneration after injury has been controversial. Although generally considered an anti-growth signal, its role is far more complex as evidenced by conflicting results using p75NTR knock-out mice.

Although expression is high during development, mature Schwann cells do not express p75NTR. Schwann cell expression of p75NTR increases after injury (Taniuchi et al., 1986; Heumann et al., 1987a,b). Deletion of p75NTR in Schwann cells mediates improved regeneration in DRG neurons (Scott and Ramer, 2010). Conversely, for motoneurons, Schwann cell p75NTR deletion results in diminished functional recovery and axonal growth (Tomita et al., 2007). Expression of p75NTR is thought to mediate remyelination through a BDNF-dependent mechanism.

In motoneurons, p75NTR levels rise dramatically after injury, returning to baseline levels by 30 days (Raivich and Kreutzberg, 1987; Yan and Johnson, 1988; Ernfors et al., 1989; Koliatsos et al., 1991; Saika et al., 1991; Rende et al., 1995; Gschwendtner et al., 2003). This upregulation in p75NTR does not result in motoneuron cell death, however (Bueker and Meyers, 1951; Kuzis et al., 1999). Treating injured motoneurons with lowlevels of recombinant human BDNF enhances their regeneration. Higher doses, however, result in failure to regenerate, which can be reversed by p75NTR blockade (Boyd and Gordon, 2002). There is currently no motoneuron-specific p75NTR knockout model, but conflicting results have been found with regard to motoneuron regeneration in p75NTR pan-knockout mice. Boyd and Gordon found improved motor axon regeneration in knockout mice after peroneal nerve transection (Boyd and Gordon, 2001). Gschwendtner et al. found no effect of knocking out p75NTR on facial nerve axon regeneration (Gschwendtner et al., 2003). Ferri et al. found worse axon regeneration but improved functional recovery after facial nerve crush (Ferri et al., 1998). Song et al. found fewer regenerating axons in p75NTR knockout mice using both sciatic nerve and facial nerve crush injuries (Song et al., 2009). Most recently, Zhang et al. found worse axonal regeneration among p75NTR knockout mice using a facial nerve crush model (Zhang et al., 2010). Using cell-type specific knockout of p75NTR or targeting the binding partners of p75NTR that result in different signaling could provide clarity to these conflicting results in how p75NTR is affecting motoneuron regeneration.

Sensory neurons decrease expression of p75NTR after injury (Zhou et al., 1996). Unlike adult motoneurons, there is significant cell death of DRG neurons after an injury, but this is restricted to small-diameter, mainly cutaneous afferent neurons (Welin et al., 2008; Wiberg et al., 2018). Cell death in these smaller DRG neurons can be blocked by application of NGF (Rich et al., 1987; Ljungberg et al., 1999). Likewise, in injured DRG neurons, BDNF acts in an autocrine loop to prevent cell death, and disruption in BDNF expression increases cell death (Acheson et al., 1995). A decrease in expression of p75NTR could act as a survival signal, such that the endogenous increased neurotrophin secretion would be more likely to bind trk receptors and result in pro-survival signaling. In support of this hypothesis, in cultures of DRG neurons in which p75NTR has been rendered inactive, cell survival is higher, and increased neurotrophin concentration does not result in increased cell death (Zhou et al., 2005). Similarly, disrupting the NT binding domain of p75NTR results in increased sprouting after injury in DRG neurons (Scott et al., 2005).

Despite the upregulation of BDNF and its receptors after injury, neither BDNF nor the other neurotrophins (NGF, NT3, and NT4/5), is required for spontaneous regeneration of peripheral neurons (Diamond et al., 1987, 1992; Streppel et al., 2002; Wilhelm et al., 2012). However, application of exogenous BDNF enhances axonal regeneration, functional recovery and decreases synaptic stripping (Lewin et al., 1997; Boyd and Gordon, 2002, 2003; Davis-Lopez de Carrizosa et al., 2009). Recently, small molecule trkB agonists have been developed, and these also enhance regeneration after injury (English et al., 2013).

### ACTIVITY DEPENDENT TREATMENTS ENHANCE REGENERATION

The first published report using activity-dependent treatments to enhance peripheral nerve regeneration was from Hines in 1942. He tested both electrical stimulation (ES) and different exercise paradigms in enhancing functional outcome in rats with tibial nerve transections (Hines, 1942). Since then, there has been great interest in treatments which activate injured neurons, collectively known as activity-dependent treatments, to enhance nerve regeneration (Udina et al., 2011a). These treatments include ES (**Table 1**), exercise (**Tables 2**–**4**), and more recently, optogenetic stimulation (**Table 5**). One benefit of activity-dependent treatments is the potential for easy translation from bench to bedside—using ES and exercise in human patients would require meeting far fewer regulatory requirements than the use of an experimental drug. Moreover, for nerve injuries that require surgical intervention, ES could be performed easily at the time of surgical repair of the nerve, as has already begun with clinical trials for patients undergoing surgery for carpal tunnel release and complete digital nerve transection (Gordon et al., 2010; Wong et al., 2015). Exercise has the advantage of being low cost and allowing patients to take control of their recovery. However, in the case of extensive injuries, exercise may not be an option for a recovering patient. For this reason, finding treatments that mimic the effects of exercise, such as optogenetic stimulation, may be advantageous in treating patients. To accomplish such a goal, an understanding of the biological basis for the enhancement seen with these treatments is necessary.

#### Electrical Stimulation

Immediately after a peripheral nerve injury, a calcium wave propagates along the cut axons toward their cell bodies. Blocking this calcium wave through inhibition of voltage gated calcium channels or inhibition of calcium release from the neuronal endoplasmic reticulum blocks regeneration (Ghosh-Roy et al., 2010). It has been proposed that ES mimics the retrograde calcium wave that propagates at the time of injury in order to elicit cell-autonomous mechanisms that initiate regeneration (Mar et al., 2014). This hypothesis is supported by evidence that ES enhances early regeneration by accelerating the process of axons crossing the site of injury to enter endoneurial tubes in the segment of the nerve distal to the injury (Brushart et al., 2002). ES results in a doubling of the number of motoneurons crossing the site of injury into the distal nerve at 1 week after nerve injury (Brushart et al., 2002). Without treatment, axons can take as long as 4 weeks to cross the site of injury, but by 3 weeks after injury, Al Majed et al. found all electrically stimulated motoneurons had already regenerated to their target muscle compared to 8 weeks for untreated controls (Al Majed et al., 2000b; Brushart et al., 2002).

The first applications of ES focused on the functional recovery of the affected muscles. In 1983; Nix and Hopf described that as early as 2 weeks after injury, treatment with 4 Hz stimulation 24 h daily increased twitch force, tetanic tension, and muscle action potentials (Nix and Hopf, 1983). In 1985; Pockett and Gavin found earlier return of the plantar extensor reflex with just 15–60 min of 20 Hz stimulation (Pockett and Gavin, 1985). Al Majed et al. chose their 20Hz regimen based on the mean physiological frequency of motoneuron discharge and tried numerous stimulation regimens, stimulating continuously for 1 h, 1 day, 1 week, and 2 weeks. They were the first to examine the effect of ES on the regenerating axons (Loeb et al., 1987; Al Majed et al., 2000b). Just 1 h of 20 Hz stimulation resulted in longlasting enhancement of peripheral nerve regeneration. Following publication of this paper, 20 Hz stimulation became the standard for studying ES (**Table 1**).

Without treatment, axon regeneration into motor or sensory pathways in the distal segment of a cut nerve is random for the first 2 weeks following injury (Brushart, 1993). Motoneurons whose axons have entered only sensory pathways (endoneurial tubes previously occupied by cutaneous axons) remain permanently mistargeted (Brushart, 1993). Enhancing the speed of regeneration but increasing mistargeting could result in poorer functional recovery. However, ES has been shown to increase the sensorimotor specificity of regenerating axons after peripheral nerve injury. More motoneurons regenerate exclusively into motor pathways in rats treated with ES (Al Majed et al., 2000b). Fewer than 40% of injured DRG neurons reinnervated sensory pathways in controls compared to 75% in ES-treated animals (Brushart et al., 2005). However, innervating a motor endoneurial tube does not necessitate reaching the appropriate muscle target. Indeed, topographic analysis of motoneuron regeneration after ES revealed increased

Regimen specifies stimulation paradigm. Model specifies which animal and injury model was used. Result specifies what outcome measure was analyzed. Denotes improvement in

outcome measured, denotes worse outcome.

misdirection of regenerating motor axons to functionally inappropriate targets by over 500% (English, 2005). This misdirection resulted in motoneurons previously innervating extensor muscles reinnervating antagonistic flexor muscles.

Appropriate pathway innervation may rely on Schwann cells secreting specific growth factors for motor and sensory tubes. Schwann cells in the cutaneous nerves express high levels of NGF after injury, whereas Schwann cells in ventral roots express high levels of glial cell line-derived neurotrophic factor (GDNF) (Hoke et al., 2006; Brushart et al., 2013). This difference in growth factor expression has been proposed to be the mechanism through which preferential motor reinnervation occurs, with DRG neurons choosing paths with high NGF, and motoneurons entering paths with high GDNF. With no treatment, in rodents, neurotrophin expression peaks 15 days after injury and declines back to baseline by day 30. The day 15 peak coincides with the

#### TABLE 2 | Effects of treadmill training on peripheral nerve regeneration.

Regimen is the speed of running and for how long each day. Duration is how many days TT was performed. Delay refers to how long after injury before exercise was performed. Model specifies what type of injury and in what animal. Sex of animals is specified by ♀ or ♂. If this is not listed, it was not specified. Result specifies what outcome measure was analyzed.

Denotes improvement in outcome measured, denotes worse outcome. No arrow denotes no effect.

onset of pathway preference for regenerating axons (Gordon, 2015). ES, however, dramatically increases NGF secretion from Schwann cells for 3 days following stimulation (Koppes et al., 2014), possibly providing an earlier signal to regenerating sensory axons as to which pathways to take, and thus improving pathway targeting.

Regimen specifies how long swimming exercise lasted each day and how many days swimming was performed. Delay refers to how long after injury before exercise was performed. Model specifies what type of injury and in what animal. Sex of animals is specified by ♀ or ♂. If this is not listed, it was not specified. Result specifies what outcome measure was

analyzed. Denotes improvement in outcome measured, denotes worse outcome. No arrow denotes no effect.

#### Exercise Treatment

For years, the evidence for exercise enhancing regeneration was not as clear as the evidence for ES. Many different types of exercise with varying intensities applied at different times prior to or after injury have resulted in conflicting results. It was hypothesized that increased neuronal activity through exercise would enhance regeneration as early as 1979, but early studies utilizing treadmill training, voluntary wheel running, and swimming found unfavorable results (Hoffer et al., 1979; Herbison et al., 1980a; Gardiner et al., 1984; Badke et al., 1989; van Meeteren et al., 1998; Tam et al., 2001). These experiments largely focused on the effect of exercise on muscle fiber alterations and muscle function, and did not probe the effect of exercise on axon regeneration.

The change in emphasis from the effect of exercise on denervated muscle to the effect of exercise on injured spinal motoneurons and DRG neurons encouraged scientists to continue researching exercise, despite previous underwhelming results. In 2008, English and colleagues tested the efficacy of interval training (short high-speed sprints followed by periods of rest) in enhancing regeneration as a model that resembles how mice voluntarily run (De Bono et al., 2006; Sabatier et al., 2008). They found the surprising result that this regimen was effective only in female mice, and in fact the more commonly used training regimen of slow continuous treadmill walking was effective only in males (Wood et al., 2012). This previously unknown sex difference could have affected outcomes in numerous exercise experiments. For example, Seo et al. treated intact male rats with either high or low intensity treadmill training before culturing their DRGs and found only low intensity treadmill training increased neurite outgrowth (Seo et al., 2009). Their treadmill training regimen was very similar to the one used by Wood et al. that proved effective only in male mice, and the results of this experiment could have been different had females been included. Many of the prior experiments mentioned used animals of only one sex, and this could explain some of the variability in the effects of exercise (**Tables 2**–**4**).

There are a few advantages to exercise over ES. For example, while ES may increase misdirection of motoneurons reaching target muscles, treadmill training enhances motoneuron regeneration without decreasing topographic specificity (English et al., 2009). The mechanism of ES is to accelerate crossing the site of injury by regenerative sprouts; exercise does the same but also sustains pro-growth signaling throughout the process of regeneration (Gordon and English, 2016). There is also evidence that the enhancing effects of 2 weeks of exercise are more robust than that of a single bout of ES (Sabatier et al., 2008; Wood et al., 2012; Gordon and English, 2016). In 2009; Asensio-Pinilla et al. combined treadmill training with a single bout of ES given at the time of injury, and found greater enhancement of muscle reinnervation in the initial phase of recovery compared to either treatment alone (Asensio-Pinilla et al., 2009). Thus, after inauspicious beginnings, exercise has shown great promise as a treatment in the field of peripheral nerve regeneration.

#### Optogenetic Stimulation

The advent of optogenetics enabled cell-specific neuronal activation with the use of the light-sensitive cation channel, Channelrhodopsin (ChR2) (Krook-Magnuson et al., 2014). Whereas ES stimulates all cells within the nerve (including Schwann cells and various immune cells) and exercise likely affects cells throughout the entire body, specific neuronal activation can be achieved using optogenetics by expressing ChR2 only in neurons. Park et al. were the first to demonstrate the efficacy of light stimulation in enhancing regeneration by replicating the common ES protocol using light stimulation of 20 Hz for 1 h on explanted neonatal DRGs (Park et al., 2015). Although they tested a number of different stimulation regimens, the 1 h of 20 Hz stimulation provided the largest effect on neurite outgrowth. Ward et al. recapitulated this in vivo, finding that 1 h of 20 Hz light stimulation of light-sensitive neurons enhanced axon regeneration only in the light-sensitive cells (Ward et al., 2016, 2018).

#### MECHANISMS

#### Neurotrophins

Activity-dependent treatments require neuronal neurotrophin production.  TABLE 4 | Effect of other exercise paradigms on peripheral nerve regeneration.

Exercise refers to what type of paradigm was used. Regimen specifies how long exercise lasted each day and how many days exercise was performed. Delay refers to how long after injury before exercise was performed. Model specifies what type of injury and in what animal. Sex of animals is specified by ♀ or ♂. If this is not listed, it was not specified. Result specifies

what outcome measure was analyzed. Denotes improvement in outcome measured, denotes worse outcome. No arrow denotes no effect.

Electrically stimulating Schwann cells increases their secretion of NGF specifically, and not BDNF (Koppes et al., 2014). While Schwann cell NGF is sufficient to promote axon growth, the study of axon regeneration through nerve grafts made acellular by repeated freezing and thawing has demonstrated that stimulation of Schwann cells (and other cell types) is not required for the efficacy of ES to enhance axon regeneration (English et al., 2007; Koppes et al., 2014). Moreover, the use of optogenetics to stimulate neurons selectively has shown that specific neuronal activation is sufficient to enhance regeneration (Ward et al., 2016).  Thus, while activity-dependent treatments may increase Schwann cell neurotrophin secretion, this is not required for their enhancing effects.

Unlike non-neuronal cells, neuronal neurotrophin secretion is required for the efficacy of activity-dependent treatments. Genetically deleting NT4/5 or BDNF from Schwann cells does not alter the efficacy of ES or treadmill training in enhancing axon growth, but deleting these neurotrophins from neurons abolishes the effectiveness of these activity-dependent treatments (English et al., 2007; Wilhelm et al., 2012). Both exercise and ES have been shown to increase neuronal BDNF and its receptor,

Regimen specifies stimulation paradigm. Mouse specifies what transgenic mouse model was used. Result specifies what outcome measure was analyzed. Denotes improvement in outcome measured, denotes worse outcome.

trkB (Al Majed et al., 2000a; Gomez-Pinilla et al., 2002; English et al., 2007; Park and Höke, 2014; Park et al., 2015). Through coculturing light-sensitive DRG explants with wild type DRGs, Park et al. demonstrated that the BDNF secreted in response to light stimulation was sufficient to increase neurite outgrowth not only from cells in a light-sensitive (ChR2-expressing) DRG, but also in neighboring ganglia derived from wild type mice. Protein analysis of the media revealed increased BDNF and NGF secretion in response to optical stimulation from the light-sensitive DRGs

only (Park et al., 2015). There is a dose-dependence in activity-dependent treatments for enhancing nerve regeneration. Whereas 1 h of 20 Hz stimulation has been shown to enhance DRG regeneration after injury, an increase to just 3 h of ES decreased sensory neuron regeneration, and was associated with a downregulation in expression of the regeneration associated gene, GAP-43 (Geremia et al., 2007). In vitro, neurites from DRG explants containing ChR2 had higher rates of growth with stimulation paradigms resulting in 72k pulses of light (1 h 20 Hz, 2 h 10 Hz, 4 h 5 Hz) than stimulation paradigms that resulted in a higher number of pulses (20 Hz for 1–3 days) or much lower number of pulses (20 Hz for 15 min) (Park et al., 2015). Three days of continuous depolarization through ES or high concentrations of KCl results in complete failure of dissociated DRG neurons to grow neurites in culture (Enes et al., 2010). For motoneurons, high intensity exercise or repeated bouts of ES result in decreased sprouting and fewer synaptic contacts at neuromuscular junctions (Tam et al., 2001). Application of 1 h of 20 Hz ES every third day for 2 weeks after sciatic nerve transection and repair did not enhance the regeneration of motor axons in mice (Park et al., under review). Interestingly, exogenous application of BDNF resulted in a dosedependent enhancement of axon regeneration as well (Boyd and Gordon, 2002). Low to modest doses produced enhanced axon regeneration, but higher doses inhibited regeneration. Treatments with high doses of BDNF caused p75NTR activation, which prevented DRG neurite outgrowth (Boyd and Gordon, 2002).

#### Neuronal Activity

The success of activity-dependent treatments in promoting axon regeneration requires activation of the injured neurons. Treating the neurons proximal to the stimulation site with tetrodotoxin (TTX) to block their ability to conduct antidromic action potentials abolishes the effect of ES, despite the continued orthodromic firing of distal axons and muscle fibers (Al Majed et al., 2000b). Similarly, inhibition of motoneuron activity during treadmill training, using bioluminescent optogenetics (BL-OG), abolishes the enhancing effect of exercise on motoneuron regeneration (Jaiswal et al., 2017). Whether the increased activation needed to promote axon regeneration requires action potential generation is not entirely clear. Enhancement of regeneration of axons of many more motoneurons than are likely to be brought into full activity is found after treatments with exercise at a slow treadmill speed (Gordon and English, 2016). Simply increasing the excitability of injured neurons using chemogenetics could be sufficient to enhance regeneration (Jaiswal et al., 2018).

Although Park et al. found that BDNF secretion from neighbors can stimulate regeneration in neurons that were not activated in vitro, optogenetic stimulation in vivo of only motoneuron axons did not enhance DRG axon regeneration, nor vice versa (Ward et al., 2018). When BDNF is knocked out in only a subset of neurons, those specific neurons do not benefit from exercise treatment (Wilhelm et al., 2012). Thus, it appears neuronal BDNF is acting as an autocrine signal facilitating enhanced regeneration (English et al., 2014; Gordon, 2016).

#### Androgens

The sex difference found in response to different exercise regimens led to the hypothesis that androgen receptor signaling was involved in activity-dependent treatments. The effect of androgens in enhancing peripheral nerve regeneration had already been thoroughly explored several years prior (Fargo et al., 2009). All motoneurons contain androgen receptors, though testosterone is not required for spontaneous regeneration treating animals of both sexes with the androgen receptor blocker flutamide does not inhibit regeneration, nor does castration of males (Freeman et al., 1995; Thompson et al., 2013). Application of exogenous androgens in males and females, however, enhances axon regeneration in both cranial and spinal nerve injuries (Kujawa et al., 1989, 1991; Jones, 1993; Freeman et al., 1995; Tanzer and Jones, 1997; Brown et al., 1999). This effect is androgen receptor-dependent, and blocking the androgen receptor with flutamide prevents testosterone-induced enhanced regeneration (Kujawa et al., 1995). In females, treating mice with anastrazole, an aromatase inhibitor which blocks the conversion of testosterone into estradiol, also dramatically enhanced axon regeneration, without increasing serum androgen levels (Thompson et al., 2013).

The sex difference in response to exercise regimens was evidence that androgen receptor signaling and activitydependent treatments were linked. Slow, continuous treadmill training resulted in an increase in serum testosterone levels in males, though no similar increase was found for females with interval training (Wood et al., 2012). Castrating males prior to treadmill training abolishes its enhancing effect, which cannot be rescued with interval training (Wood et al., 2012). Treating both sexes with flutamide before appropriate exercise regimens abolishes the effectiveness of this treatment in enhancing peripheral nerve regeneration (Thompson et al., 2013). Testosterone is also necessary for the beneficial effects of ES—castrated rats treated with ES have poorer regeneration compared to littermates who are treated with exogenous testosterone (Hetzler et al., 2008; Sharma et al., 2009). As with exercise, flutamide blocks the enhancing effects of ES in both males and females (Thompson et al., 2013). Conversely, combined exogenous androgen treatment with ES enhances facial nerve regeneration in gonadally intact rats (Sharma et al., 2010b).

Androgens regulate BDNF and its receptor, trkB, in motoneurons (Osborne et al., 2007; Ottem et al., 2010; Sharma et al., 2010a; Verhovshek et al., 2010). Exercise elicits an upregulation of testosterone that is sustained and could result in an increased duration of BDNF and trkB expression (Thompson et al., 2013; English et al., 2014). ES elicits an early increase in BDNF expression, whereas exogenous androgen application results in a later and longer-duration increase in expression. Combining the two treatments results in an additive upregulation of BDNF and could explain the improved recovery over either treatment alone (Sharma et al., 2010a).

#### SYNAPTIC REARRANGEMENTS

After peripheral nerve injury both excitatory and inhibitory synaptic inputs onto injured motoneurons are withdrawn (Blinzinger and Kreutzberg, 1968; Brannstrom and Kellerth, 1998; Linda et al., 2000; Oliveira et al., 2004). If motor axons regenerate and reinnervate muscle targets, many of these inputs are restored but, for those expressing the vesicular glutamate transporter 1 (VGLUT1) and arising from muscle spindles, a gradual withdrawal of their central axonal processes from the ventral horn follows, resulting in a permanent loss of these synaptic inputs (Alvarez et al., 2011; Rotterman et al., 2014). In animals treated with exercise during the first few days following sciatic nerve transection and repair, the extent of synaptic contacts between these important sources of proprioceptive feedback and motoneurons is not reduced (English et al., 2011; Liu et al., 2014; Krakowiak et al., 2015). This robust connectivity by VGLUT1+ inputs is retained at least 12 weeks later. No similar effect is found if the onset of the exercise treatment is delayed (Brandt et al., 2015). Application of 1 h of 20 Hz ES had no effect on synaptic coverage after nerve injury, but repeated applications every third day for 2 weeks resulted in an effect similar to that observed using exercise (Park et al., under review). It is not clear whether this effect of these activity-dependent therapies is the prevention of the original synaptic withdrawal, a stimulation of new synapse formation to replace the withdrawn inputs, or some combination of both. More studies are needed.

It is clear that BDNF plays a role in maintaining and preserving synaptic inputs on motoneurons. Without exercise, axotomized motoneurons lose approximately 35% of their overall synaptic coverage (Krakowiak et al., 2015). This effect is BDNFdependent—knocking out BDNF in a subset of motoneurons reduces synaptic coverage in those specific cells in intact animals, and this synapse loss cannot be rescued with exercise (Krakowiak et al., 2015). Wild-type motoneurons within an animal maintain their synaptic contacts after nerve injury with exercise, but those in which BDNF has been knocked out do not (Krakowiak et al., 2015).

### BDNF VAL66MET POLYMORPHISM

Given the relationship between activity-dependent treatments and BDNF, any genetic mutations altering BDNF signaling among the human population could affect the success of these treatments. Such a mutation exists—a common single nucleotide polymorphism in the BDNF gene. The G to A mutation at site 196 results in a Valine to Methionine substitution in the 66th codon (**Figure 1**). This polymorphism was first described by Egan et al. in 2003 and was quickly identified as incredibly common—the met allele of the BDNF gene is present in 25% of the American population and up to 50% of East Asian populations (Egan et al., 2003; Shimizu et al., 2004). Carrying the met allele was originally described as a risk factor for schizophrenia (Egan et al., 2003). It has since been linked to numerous other disorders and diseases, including Alzheimer's disease, obsessive compulsive disorder, anorexia nervosa, and bipolar disorder (Neves-Pereira et al., 2002; Sklar et al., 2002; Egan et al., 2003; Hall et al., 2003; Ribases et al., 2003; Notaras et al., 2015). Physiologically, Met-carriers have been found to have decreased hippocampal volume, and cells transfected with the Met allele have altered activity-dependent secretion of BDNF (Egan et al., 2003).

Testing for deficient activity-dependent secretion of BDNF in humans can be tricky. Generally, BDNF secretion is measured through serum as an indirect measure of neuronal BDNF, and exercise is a reliable method to increase serum BDNF levels (Berchtold et al., 2005; Elfving et al., 2010; Klein et al., 2011; Szuhany et al., 2015). Although one study has found that healthy adult Met-carriers did have increased serum BDNF after exercise (Helm et al., 2017), others have found serum levels of BDNF did not increase after high intensity exercise in elderly (Nascimento et al., 2015), spinal cord injured (Leech and Hornby, 2017), or healthy Met-carriers (Lemos et al., 2015). In mice expressing the met allele, exercise results in deficient mRNA production as well as decreased protein expression of BDNF (Ieraci et al., 2016). These deficiencies in exercise-induced BDNF secretion mirror the findings in cultured neurons expressing the Met allele, and the use of cells transfected with the met allele in vitro as well as the development of a transgenic mouse have allowed researchers to elucidate the mechanism behind this deficient secretion (Egan et al., 2003; Chen et al., 2004, 2006).

The valine to methionine substitution in this SNP occurs in the prodomain of the BDNF protein (Egan et al., 2003). Although it does not affect the ability of mBDNF to bind its receptor, this substitution results in disorganized folding of the prodomain, resulting in abnormal interactions with sortilin (see above) (Chen et al., 2004; Anastasia et al., 2013). BDNFMet is thus packaged inefficiently into calcium-sensitive secretory vesicles, accounting for the deficient activity-dependent secretion that has been reported (Chen et al., 2004). Being heterozygous for the met allele does not protect from this deficient BDNF secretion— BDNF forms homodimers, and in cells heterozygous for the met allele, BDNFMet dimerizes with BDNFVal and prevents its packaging into Ca2+-regulated secretory vesicles (Kolbeck et al., 1994; Chen et al., 2004). Analysis of activity-induced BDNF secretion from cultured hippocampal cells bears this out—those cells heterozygous for the Met allele have deficient activitydependent secretion despite the presence of one copy of the BDNFVal allele (Chen et al., 2006). Furthermore, once secreted, BDNF availability may be affected by binding with the cleaved prodomain. The prodomain binds BDNF with high affinity, and the met allele results in enhanced BDNF binding and slower dissociation once bound (Uegaki et al., 2017). This could limit the availability of BDNF to bind its receptors.

Activity-dependent secretion of BDNF relies not only on packaging into calcium-sensitive vesicles, but also on the spatial targeting of mRNA into distal processes where BDNF can be locally translated (Chiaruttini et al., 2008). This targeting is achieved through binding of BDNF mRNA with translin, a DNA/RNA binding protein involved in dendritic trafficking of mRNAs (Li et al., 2008). The G to A mutation at site 196 disrupts translin binding of BDNF mRNA, and thus Metcarriers have deficient trafficking of BDNF mRNA to distal processes (Chiaruttini et al., 2009). Moreover, the transcripts containing exon VI, which is upregulated by exercise, and exon IV, which is calcium-sensitive, are found in reduced levels in the hippocampus of mice homozygous for the met allele (Tao et al., 1998; Baj et al., 2012; Mallei et al., 2015). These transcripts, along with those containing exon II, are generally trafficked to distal processes (Baj et al., 2011).

In addition to deficient activity-dependent BDNF secretion, the met allele may result in increased p75NTR activation. Unlike BDNFVal, when the prodomain is cleaved from BDNFMet, it is bioactive and able to activate p75NTR with the help of SorCS2, a member of the sortilin family of receptors (Deinhardt et al., 2011; Anastasia et al., 2013). In vitro, application of exogenous prodomain protein results in growth cone collapse and dendritic spine disassembly (Anastasia et al., 2013; Giza et al., 2018). Stimulating cells with high KCl concentration results in activitydependent secretion of both Val and Met prodomains, though secretion is deficient in Met-carriers (Anastasia et al., 2013). Although endogenous secretion of the Met prodomain has yet to be linked to alterations in dendrites, decreased arborization has been found in hippocampal and cortical neurons (Chen et al., 2006; Liu et al., 2012).

The deficit in activity-dependent release of BDNF led to the hypothesis that activity-dependent treatments to enhance axon regeneration after peripheral nerve injury would be ineffective in this population. Using a mouse model of this polymorphism which recapitulates certain phenotypic aspects of the human population such as decreased hippocampal volume and increased anxiety-like behavior (Chen et al., 2006), we tested the efficacy of treadmill training on motor axon regeneration 4 weeks after complete sciatic nerve transection and repair in mice both heterozygous and homozygous for the met allele of the Bdnf gene (McGregor et al., under review). Exercise was completely ineffective in enhancing axon regeneration in the Met-carriers. However, peripheral axon regeneration in Met-carriers was surprisingly enhanced without any treatment (McGregor et al., under review). One possibility for the failure of exercise to enhance regeneration in Met-carriers is a ceiling effect exercise was not able to further enhance an already accelerated regeneration. The question of why regeneration is accelerated remains to be explored. The enhanced regeneration was found both in vivo and in cultured DRG neurons, indicating that enhanced axon outgrowth is a neuronal trait in Met-carriers. This unanticipated result is some of the first good news regarding what is a maligned SNP, although others have reported the met allele may also be protective in stroke and traumatic brain injury (Krueger et al., 2011; Rostami et al., 2011; Qin et al., 2014; Failla et al., 2015). Thus, in human populations there may be striking differences in response to peripheral nerve injury dependent on individual gene expression. Enhanced regeneration associated with the Val66Met polymorphism may explain the persistence of the mutation within the human population.

### CONCLUSION

The suboptimal regeneration of peripheral nerves presents a challenge in medical care. Neurotrophins, particularly BDNF, have been studied for their pro-growth properties, and treatments that stimulate endogenous release of neurotrophins have been successful in enhancing regeneration in animal models. These treatments are currently being tested to enhance peripheral nerve regeneration in patients with some success (Gordon et al., 2010; Wong et al., 2015). The existence of genetic polymorphisms in the bdnf gene, however, will affect the outcome of these experiments, and preliminary investigations as to the efficacy of activity-dependent treatments in individuals with the met allele will hopefully spur the field toward personalized medicine. Activity-dependent treatments can be a powerful tool for those responsive to them, and for the rest, new therapies that do not rely on endogenous BDNF-signaling must be developed.

#### AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

#### REFERENCES

#### ACKNOWLEDGMENTS

Support provided by grant NS057190 to AE from the USPHS. Many thanks to Dr. Francis Lee for the gift of the Val66Met mouse. The manuscript is based in part on a thesis for the Ph.D. degree in Neuroscience from The Laney Graduate School of Emory University (McGregor, 2018).

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 McGregor and English. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# The Success and Failure of the Schwann Cell Response to Nerve Injury

Kristjan R. Jessen\* and Rhona Mirsky

Department of Cell and Developmental Biology, University College London, London, United Kingdom

The remarkable plasticity of Schwann cells allows them to adopt the Remak (nonmyelin) and myelin phenotypes, which are specialized to meet the needs of small and large diameter axons, and differ markedly from each other. It also enables Schwann cells initially to mount a strikingly adaptive response to nerve injury and to promote regeneration by converting to a repair-promoting phenotype. These repair cells activate a sequence of supportive functions that engineer myelin clearance, prevent neuronal death, and help axon growth and guidance. Eventually, this response runs out of steam, however, because in the long run the phenotype of repair cells is unstable and their survival is compromised. The re-programming of Remak and myelin cells to repair cells, together with the injury-induced switch of peripheral neurons to a growth mode, gives peripheral nerves their strong regenerative potential. But it remains a challenge to harness this potential and devise effective treatments that maintain the initial repair capacity of peripheral nerves for the extended periods typically required for nerve repair in humans.

#### Edited by:

Esther Udina, Autonomous University of Barcelona, Spain

#### Reviewed by:

Valerio Magnaghi, University of Milan, Italy Edgar Richard Kramer, Plymouth University, United Kingdom

#### \*Correspondence:

Kristjan R. Jessen<EMAIL_ADDRESS>
Received: 03 October 2018 Accepted: 22 January 2019 Published: 11 February 2019

#### Citation:

Jessen KR and Mirsky R (2019) The Success and Failure of the Schwann Cell Response to Nerve Injury. Front. Cell. Neurosci. 13:33. doi: 10.3389/fncel.2019.00033 Keywords: PNS, repair cell, nerve injury, regeneration, c-Jun, re-programming, Schwann cell

#### INTRODUCTION: OVERVIEW OF SCHWANN CELLS AND NERVE INJURY

Examination of the Schwann cells in uninjured nerves shows two surprisingly different cell types. One of them is the rather inconspicuous Remak cell, or non-myelin Schwann cell. These cells envelop all the small diameter axons, including many sensory axons and the all-important axons of the autonomic nervous system. The axons lie in troughs along the cell surface, and in rodents each cell generally ensheaths several axons, although in human nerves, one axon per Remak cell is common. The larger axons, including some sensory axons and the axons of motor neurons, are wrapped by myelin Schwann cells. They are 2–3 times longer that Remak cells and much bulkier, containing the myelin sheath, which is formed by the Schwann cell membrane wrapping multiple times around the axon and condensing to form a compact myelin cuff around the axon.

Both Remak and myelin cells are coated by a basal lamina, outside of which lies the connective tissue, the endoneurium, which contains fibroblasts, blood vessels and a few macrophages and is ultimately surrounded by a multi-layered cellular tube, the perineurium (**Figure 1**). This assembly is termed a fascicle. Small nerves are uni-fascicular, while large nerves contain many fascicles bound together by connective tissue, the epineurium.

While both Schwann cells are thought to provide axons with metabolic and trophic support, only the myelin cells have the key role of accelerating nerve impulse conduction. Remak and myelin cells also express some common proteins, such as S100, a classical Schwann cell marker, while at the same time each cell possesses a characteristic molecular profile (reviewed in Jessen and Mirsky, 2005, 2016; Glenn and Talbot, 2013; Brosius Lutz and Barres, 2014; Monk et al., 2015). Thus, Remak cells express several markers also found on developing Schwann cells, such as neural cell adhesion molecule (NCAM), p75 neurotrophin receptor (p75NTR) and glial fibrillary acidic protein (GFAP), and L1 NCAM. The myelin cells on the other the hand express a large range of molecules that relate to the synthesis, maintenance or structure of the myelin sheath. This includes the major pro-myelin transcription factor Egr2 (Krox20), high levels of the enzymes that control cholesterol synthesis, structural proteins such as myelin protein zero (MPZ) and myelin basic protein (MBP), and membrane associated proteins like myelin associated glycoprotein (MAG), PMP22, and periaxin.

As different as Remak and myelin cells appear, these cells nevertheless originate developmentally from a common cell, the immature Schwann cell, present in rodent nerves in the perinatal period (**Figure 2**). These cells, in turn, derive from a distinct glial cell of embryonic nerves, the Schwann cell precursor. The conversion of Schwann cell precursors to Schwann cells is controlled by transcription factors such as AP2α, Zeb2, and Notch, and extracellular signals including endothelin and axon-associated neuregulin, which is essential for precursor survival (reviewed in Jessen and Mirsky, 2005; Jessen et al., 2015b; Quintes and Brinkmann, 2017).

Schwann cell precursors unambiguously exhibit a glial phenotype, expressing characteristic glial features, such as ensheathing axons within nerves and expressing Schwann cell associated mRNAs, such as those encoding for the major myelin protein MPZ and desert hedgehog (Dhh). It is interesting, however, that these cells also retain one notable feature of the neural crest cells from which they originate, namely they have a broad developmental potential. Thus, Schwann cell precursors give rise to cells such as melanocytes, endoneurial fibroblasts and neurons, in addition to Schwann cells, during embryonic development (reviewed in Jessen et al., 2015b; Kastriti and Adameyko, 2017).

One of the most interesting biological properties of Schwann cells is their plasticity (reviewed in Boerboom et al., 2017; Castelnovo et al., 2017; Jacob, 2017; Ma and Svaren, 2018). The phenotype adopted by Schwann cells, such as the Remak or myelin phenotypes described above, is strikingly dependent on signals in the cellular environment. Thus, all the evidence indicates that if a Remak cell was placed in contact with large diameter axon it would adopt the myelin phenotype and conversely, a myelin cell would convert to a Remak cell if associated with small diameter axons. This phenotypic instability may pre-dispose Schwann cells to demyelinating disease, since myelin Schwann cells may regress from myelin maintenance relatively easily in response to mutations that disturb cellular homeostasis. After nerve damage, Schwann cell plasticity and the ready response to environmental signals initially helps to provide nerves with strong regenerative potential, but at later times after injury, these features contribute to regenerative failure, as repair supportive Schwann cells are not sustained during the long times required for nerve repair in humans.

After nerve injury, the Schwann cells distal to the damaged area lose contact with axons as they degenerate. This represents

a radical change in the signaling environment, because key signals for the control Schwann cell phenotype come from axons. Subsequently, further change is provided by a cocktail of bioactive factors secreted by macrophages, which invade injured nerves in large numbers. In response to these perturbations, Schwann cells show a remarkably fortunate response. This is the conversion of Remak and myelin cells, within their original basal lamina tubes, to a Schwann cell phenotype, the repair Schwann cell, which is specialized to encourage regeneration (reviewed in Jessen and Mirsky, 2016; **Figure 2**). These cells execute a repair program, a partly overlapping sequence of phenotypic changes involving myelin autophagy, expression of cytokines that call in macrophages for later stages of myelin clearance, activation of trophic factor expression and cellular elongation and branching to form regeneration tracks, called Bungner Bands. In this way, repair cells clear myelin, support the survival of injured neurons, axon regeneration and target innervation. These are the cells present in the distal stump of injured nerves along which regenerating axons navigate after injury, often for months or even years in humans due to the slow rate of axon growth.

One of the main problems in human nerve repair is that the phenotype of repair cells is not stable, but fades with time as the cells fail to maintain expression of trophic factors that promote axonal growth, likely due to gradual changes in the signaling environment within the chronically denervated distal stump (reviewed in Höke, 2006b; Sulaiman and Gordon, 2009). Furthermore, the repair cell population is not stable, their number eventually declining to very low levels. The reasons for the deterioration of repair cells are poorly understood, although the transcription factors STAT3 and c-Jun have been implicated in this process.

Importantly, PNS neurons also respond to axonal damage by activating an extensive gene program that facilitates axonal regeneration, a response classically referred to as the cell body response or the signaling to growth mode switch (Allodi et al., 2012; Blesch et al., 2012; reviewed in Fu and Gordon, 1997; Doron-Mandel et al., 2015).

Therefore, in response to damage, both neurons and Schwann cells convert to cell states that are specialized to deal with injury and promote healing. Comparable reprogramming of differentiated cells in response to injury can also be seen in other systems. This includes the conversion of fibroblasts to myofibroblasts during wound healing, and of pigmented epithelial cells to lens cells in the eye, the conversions of supportive cells to hair cells in the ear, hepatocytes to biliary epithelial cells in the liver, and of endocrine α to β cells in the pancreatic isles. In all of these cases, differentiated cells change identity in order to promote tissue homeostasis and repair. This type of change has therefore been termed adaptive cellular reprogramming (reviewed in Jessen et al., 2015a).

 The repair Schwann cells are therefore a transient population, which exists only while they are needed.

We will now discuss some of the issues aired above in more detail.

#### THE SCHWANN CELL INJURY RESPONSE: THE GENERATION OF REPAIR CELLS

Unfortunately, in humans most nerve injuries involve nerve transection rather than the more easily repaired nerve crush. After a nerve crush, the basal lamina around each axon/Schwann cell unit remains intact and an axon stays within its basal lamina tube as it grows across the injury site to reach the distal stump. It therefore has a favorable possibility to reconnect with its original target tissue and restore function.