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. Author manuscript; available in PMC: 2022 Jul 1.
Published in final edited form as: Plast Reconstr Surg. 2021 Jul 1;148(1):32e–41e. doi: 10.1097/PRS.0000000000008051

Long Acellular Nerve Allograft’s “Cap” Transected Nerve to Arrest Axon Regeneration and Alter Upstream Gene Expression in a Rat Neuroma Model

Deng Pan 1,*, Miles Bichanich 2,*, Ian S Wood 1, Daniel A Hunter 1, Scott M Tintle 3,4, Thomas A Davis 4, Matthew D Wood 1, Amy M Moore 5
PMCID: PMC8238802  NIHMSID: NIHMS1677051  PMID: 34014904

Abstract

Objective:

Treatments to manage painful neuroma are needed. An operative strategy that isolates and controls chaotic axonal growth could prevent neuroma. Using long acellular nerve allograft (ANA) to “cap” damaged nerve could control axonal regeneration, and in turn, regulate upstream gene expression patterns.

Material and Methods:

Rat sciatic nerve was transected, and the distal nerve end reversed and ligated to generate a model end-neuroma. Three groups were used to assess their effects immediately following this nerve injury: no treatment (control), traction neurectomy, or 5 cm ANA “cap” attached to the proximal nerve. Regeneration of axons from the injured nerve was assessed over 5 months and paired with concurrent measurements of gene expression from upstream affected dorsal root ganglia (DRG).

Results:

Both control and traction neurectomy groups demonstrated uncontrolled axon regeneration revealed using Thy1-GFP rat axon imaging and histomorphometric measures of regenerated axons within the most terminal region of regenerated tissue. The ANA group arrested axons within the ANA, where no axons reached the most terminal region even after 5 months. At 5 months, gene expression associated with regeneration and pain sensitization, including Bdnf, cfos, and Gal, was decreased within DRG obtained from the ANA group compared to control or traction neurectomy group DRG.

Conclusions:

Long ANAs to “cap” a severed nerve arrested axon regeneration within the ANA. This growth arrest corresponded with changes in regenerative and pain-related genes upstream. ANAs may be useful for surgical intervention of neuroma.

Introduction

Neuromas may form after any nerve injury whether trauma, traction injury, or a consequence of amputation. Stump- or end-neuromas develop from completely severing an affected nerve, where axon outgrowth from the unrepaired proximal nerve forms a bulbous swelling of axonal sprouts and scar tissue. While the incidence of end-neuromas varies by the affected area, as an example, it’s incidence is estimated at ~6-8% after finger amputation1,2. The treatment of neuroma is a challenging clinical problem. Symptomatic neuroma cause pain that severely impacts patient productivity and quality of life, especially in amputees as the neuromas can be stimulated by prosthesis and cause the patients to discontinue their use3-5. Both non-operative modalities, such as analgesics and other medications, and surgical interventions, such as traction neurectomy and transposition, have been employed in an attempt to treat or prevent neuroma formation and recurrence6-8. However, these forms of neuroma management have yielded inconsistent efficacy9-12. Consequently, experimental operative approaches are being currently explored, where data regarding their efficacy to manage neuroma is still ongoing13-17. Therefore, it is imperative that other technically simple, while effective, surgical strategies are researched to manage neuroma.

A rational operative approach to manage end-neuroma would utilize what is known regarding the etiology of their formation and mechanisms yielding pain. Unrepaired nerves produce chaotic regrowth of axons from the proximal stump into invading fibroblastic tissue that can form a non-neoplastic tumor (i.e. neuroma)18. While not all these neuromas are painful, the spontaneous and provoked firings due to traction or pressure of the nerve endings is implicated in pain associated with neuromas19. Moreover, it is known following nerve injury that retrograde axonal signaling through molecular kinases activates the expression of regeneration associated genes and growth factors, such as brain derived neurotrophic factor (BDNF) and nerve growth factor (NGF), proximally within sensory neurons and dorsal root ganglia (DRG)20. While expression of BDNF, NGF, and related genes promotes axon regeneration, they can also cause pain sensitization21. Given the disorganized nature of neuroma, axons may not only be more sensitive to spontaneous axon potentials, but also be trapped in a state of ongoing regeneration, leading to a sustained upregulation of growth associated genes in the DRG causing sensitization to pain. Thus, an operative strategy that both isolates these axons and arrests axonal regeneration may be able to prevent neuroma, by managing axonal outgrowth and changing gene expression within DRG from the arrested axon growth.

Previously, preclinical rat studies demonstrated the repair of nerve gaps using long (i.e. >4 cm) acellular nerve allografts (ANAs) have limited ability to promote axon regeneration22-24. While axons were capable of traversing short (< 3cm) ANAs to innervate distal targets, axons became arrested within long ANAs due in part to the altered regenerative environment within the long ANAs24. Building upon this work, long ANAs (5 cm) were then used to “cap” an injured nerve in rats, where these long ANAs arrested axonal regeneration within the ANA for up to 5 weeks25. However, it was unclear if axon regeneration would be arrested over a long period (i.e. months) necessary for an effective therapy. Furthermore, it was unclear if this approach caused gene changes at the level of the DRG that would suggest changes to overall nerve regeneration and pain sensitization.

In this study, we utilized a rat sciatic nerve end-neuroma model to determine if long ANAs used to “cap” transected nerve can arrest axonal regeneration for months, and if this axon arrest is associated with changes in pain and regenerative associated genes within the DRGs. We also compared this approach to traction neurectomy, a current clinical strategy utilized in major limb amputations for management of the nerves, to provide additional context to the results.

Materials and Methods

Animals.

Adult male Lewis rats (Charles River Laboratories; Wilmington, MA) and Thy1-GFP rats (raised in house) weighing 226-250 g were used in these studies. Thy1-GFP rats (generated by GenOway (Lyon, France) on a Sprague-Dawley rat background and previously characterized26-28) have axons that express green fluorescence protein (GFP) to provide optical imaging capabilities. Animals were housed in a central housing facility and received food (PicoLab rodent diet 20, Purina Animal Nutrition LLC; St. Louis, MO) and water ad libitum. Animals were monitored postoperatively for signs of weight loss and infection. All procedures were carried out in strict accordance with the guidelines set forth by the National Institutes of Health and were approved by the Division of Comparative Medicine at the institution. Specifically, surgical procedures and peri-operative care measures were conducted in compliance with the AAALAC accredited Washington University Institutional Animal Care and Use Committee (IACUC) under animal protocol #20150120.

Experimental Design.

Studies were carried out using two arms and three groups in both arms. Both rat strains were used in experimental groups and as donor rats to provide nerve for ANAs. Lewis (RT-11 MHC) and Sprague-Dawley (RT-1b MHC) rat strains are known to be MHC incompatible for use as allograft donors29. Therefore, experimental studies using Lewis rats utilized Thy1-GFP rats donors for ANAs, while studies using Thy1-GFP rats used Lewis rat donors for ANAs. In all experimental groups, rat sciatic nerve was transected and left unrepaired to generate an end-neuroma30,31. Immediately after these procedures, experimental treatments were performed. Group I (Control) served as control in which the sciatic nerve was transected but not treated. Group II (Traction Neurectomy) and Group III (ANA) had treatment applied to the transected proximal nerve. Full experimental surgery details are described in the next section.

The first arm used Thy1-GFP rats to assess GFP+ axon outgrowth in situ over time, which determined how treatments affected dynamic axon growth until the 120-day endpoint. In this arm, 6 Thy1-GFP rats were randomized to the three experimental groups (n=2 rats per group). At 30- and 120-days post injury, in situ fluorescent macroscopy on the sciatic nerve was conducted using a single, live rat from each treatment group. For the second arm, Lewis rats were used to assess the status of nerve regeneration in the long term. This arm quantified the extent to which treatments affected nerve regeneration from the proximal injured nerve, and upstream gene expression within dorsal root ganglia (DRG). For this arm, 15 Lewis rats were randomized to the three experimental groups (n=5 per group). At 120 days post nerve injury, the proximal sciatic nerves along with regenerated tissue or ANA were harvested en bloc for histomorphometric analysis, while the L4-L5 DRGs together were harvested for gene analysis. At all endpoints following procedures, donor animals were euthanized by intraperitoneal injection of sodium pentobarbital (200 mg/kg, Vortech Pharmaceutical Ltd; Dearborn, MI).

ANA Procurement and Processing.

Donor rats were anesthetized with a subcutaneous injection of ketamine (75 mg/kg, Ketaset®, Fort Dodge Animal Health; Fort Dodge, IA) and medetomidine (0.5 mg/kg, Dormitor®, Orion Corporation; Espoo, Finland) and the sciatic nerves were harvested bilaterally. The donor animals were then euthanized by intraperitoneal injection of sodium pentobarbital (200 mg/kg). ANAs were decellularized using a modified series of detergents in the method described previously22,23. Briefly, nerves were repeatedly washed in deionized water and three detergents in a sodium phosphate buffer: Triton X-100, sulfobetaine-16 (SB-16), and sulfobetaine-10 (SB-10). All grafts were washed and stored in 10 mM phosphate-buffered 50 mM sodium solution at 4°C and used within 3 days.

Surgical Procedures.

Surgical procedures were performed using aseptic conditions with the aid of an operating microscope (JEDMED/KAPS, St. Louis, MO). Animals were identically anesthetized as for ANA Procurement detail above. The right hindquarter was shaved and prepped using betadine and ethanol, and the skin divided parallel to the femur. Under 10-25x magnification using sterilized micro-instruments, the right sciatic nerve was exposed through a gluteal muscle-splitting incision and the nerve transected 5 mm proximal to the trifurcation with surgical scissors. The distal nerve stump was ligated with 4-0 silk suture (Ethicon; Somerville, NJ) and then pulled away from the proximal stump and sutured with 9-0 nylon (Surgical Specialties Corporation; Wyomissing, PA) to nearby muscle. This maneuver was used to decrease possible neurotropic influence. For the traction neurectomy group, the proximal nerve stump was pulled taut before being transected again to remove 2.5mm of nerve. The nerve was allowed to retract into the proximal muscle. For the ANA group, ANAs were cut to 5 cm and coapted in reverse orientation via the proximal nerve epineurium with four to five 9-0 nylon sutures. The distal end of the ANA was then secured by a subcutaneous pocket created anteriorly using 9-0 nylon to secure the ANA and keep it away from both the proximal stump and ligated distal stump. Upon completion of the procedures, the incision was irrigated with bacteriostatic 0.9% sodium chloride (Hospira, Inc.; Lake Forest, IL) and the muscle fascia was closed with 4-0 Vicryl suture (Ethicon) and skin was closed in layers using 4-0 nylon suture (Ethicon). The animals were recovered with atipamezole hydrochloride (1 mg/kg, Antisedan®, Orion Corporation; Espoo, Finland) on warmed heating station and closely monitored before returning to a central housing facility. Postoperative pain was managed using Buprenorphine SR™ (0.05 mg/kg; ZooPharm, Windsor, CO) q 8-12 hrs prn.

Thy1-GFP Fluorescence Macroscopy.

The growth of axons from Thy1-GFP rats was measured at post-operative days 0, 30, and 120 using a stereo-enabled macroscope with fluorescent light source (Olympus MVX10, Olympus Corporation; Center Valley, PA). Rats were anesthetized and prepped as previously, then the sciatic nerve exposed to include the sciatic notch proximally and all the area of tissue outgrowth. The re-exposed nerve and surrounding tissue were imaged in live animals using brightfield and fluorescence illumination (FITC filtercube). Image acquisitions were standardized according to magnification (overall 6.3X) and exposure time.

Histomorphometry.

Nerves from Lewis rats, including the regenerating tissue from the proximal nerve end or ANA, were harvested and marked with a proximal suture, then stored in 3% glutaraldehyde (Electron Microscopy Sciences; Hatfield, PA) in 0.1 M pH 7.2 phosphate buffer (Fisher Scientific; Fair Lawn, NJ) at 4° C until processing. The explanted tissues were processed and analyzed as described previously32. Briefly, the tissues were post-fixed in 1% osmium tetroxide (Polysciences, Inc., Warrington, PA), and serially dehydrated in 90% ethanol (Thermo Fischer Scientific, Winmill Hill, UK). The tissues were then embedded in grade 502 epoxy (Polysciences, Inc., Warrington, PA) and sectioned to 1 μm with an ultramicrotome. The slides were stained with 1% toluidine blue and analyzed under light microscopy at 1000x (Leitz Laborlux S, Leica; Buffalo Grove, IL) using the IA32 Image Analysis System (Leco; St. Joseph, MI) in order to quantify nerve fiber counts, fiber width, fiber density, and percent neural tissue. One hundred twenty (120) days after the initial nerve injury, the number of myelinated axons within the proximal, middle and distal segment of regenerated tissue, as well as proximal to the initial injury were examined. All analysis was conducted by a blinded observer to the experimental groups.

Quantitative RT-PCR.

At the time of nerve harvest in Lewis rats, the L4 and L5 DRG were harvested. Total RNA was isolated from the pooled L4-L5 DRG from each animal using Trizol (Life Technologies), chloroform, and a RNeasy Kit (Qiagen, Valencia, CA) according to manufacturer’s protocol. DRGs from 5 rats per group were analyzed. RNA concentration was determined on a NanoDrop 1000 Spectrophotometer (Thermo Scientific, Wilmington, DE). The cDNA was generated with SuperScript II reverse transcriptase (Invitrogen, Carlsbad, CA). Real-time PCR was performed using a Step One Plus thermocycler (Applied Biosystems, Foster City, CA) using Taqman Master Mix (Applied Biosystems) reagents with Taqman gene expression probe for Bdnf, Galactin, and cfos (Thermofisher Scientific, Waltham, MA). The expression of genes were normalized to Actb in each group. Data were analyzed using Step One Software v2.2.2 (Applied Biosystems, Foster City, CA). The gene expression of BDNF, Galactin, and cFOS was measured as representative genes of interest based upon the literature33-35. Gene expression from DRGs of uninjured rats to control and experimental groups was compared.

Statistical Analysis.

All data was compiled as mean ± standard deviation and tested for normality using the Kolmogorov-Smirnov test. One-way analysis of variance (ANOVA) was used to examine means from the histomorphometry data. If analysis demonstrated significant differences, a Tukey’s post-hoc test was performed to isolate these differences while correcting for multiple comparisons. Statistical analysis of nerve data was performed using GraphPad Prism (Version 7.0; GraphPad Software, La Jolla, CA). Significance was set at α = 0.05 (P < 0.05).

Results

To assess how the treatments affected axon outgrowth following injury, the growth of GFP+ axons was visualized using Thy1-GFP rats up to 120 days following initial injury (Fig. 1). The Control group demonstrated evidence of GFP+ axon growth distal to the site of injury at post-operative days 30 and 120. Similarly, the Traction Neurectomy group demonstrated similar behavior, where axon extension was also observed at post-operative days 30 and 120. For the ANA group, axon extension was observed dynamically over time, but axons did not reach the pectineus muscle and remained within the attached ANA. Interestingly, the control and traction neurectomy groups, in contrast to the ANA group, showed bulbus nerve near the site of initial injury indicative of neuroma (Fig.1).

Figure 1. An ANA “cap” arrests axon regeneration within the integrated ANA tissue.

Figure 1.

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Fluorescence images captured from Thy1-GFP rats with an end-neuroma model followed by interventions. Images from day 0, 30, and 120 are shown. Note axon regeneration to the most terminal region of Control and Traction Neurectomy groups. Arrowhead: bulbous nerve region within the transected proximal nerve. Note its absence in the ANA capped nerve.

In independent long-term studies (120 days after the initial nerve injury and applicable treatment), the extent of myelinated axon regeneration using histology with histomorphometry was assessed revealing axon growth arrest within the ANA group. All groups showed a reduction in the number of myelinated axons as a function of distance to site of injury (Fig. 2A). However, the ANA group showed complete lack of regenerated axons in the distal segment of the ANA, while the Control and Traction Neurectomy groups contained axons within this regenerated distal segment (P<0.05; Fig. 2B). Furthermore, there was no difference between Control vs Traction Neurectomy myelinated axons numbers within this most distal segment.

Figure 2. An ANA “cap” inhibits regeneration of myelinated axons at the most terminal regenerated tissue region.

Figure 2.

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A) Five months (120 days) after interventions, the number of myelinated axons in the proximal nerve and the proximal, middle, and distal region of regenerated tissue or ANA was quantified. Mean ± standard deviation shown; P value compare each regenerated tissue region to proximal nerve. B) Representative histological images and quantification of number of myelinated axons in the distal region of regenerated tissue of Control, Traction Neurectomy, and ANA groups. Mean ± standard deviation shown; N=5 per group; scale bars represent 20 μm.

In addition to the number of regenerating myelinated axons in the most terminal region (distal segment), examination of the mid-segment with regenerating axons revealed that the axon area, myelin area, and fiber area (axon + myelin) for all three groups were not different (Table 1). These findings suggest that axons that regenerated into ANAs were not different compared to other groups.

Table 1:

Histomorphometric data of mid-segment regenerated tissue or ANA harvested 120 days post-sciatic nerve injury and treatment.

Control Traction
Neurectomy		 ANA p value
Axon area (μm2) 3.5285±0.238 3.4535±0.435 3.354±0.401 0.841
Myelin area (μm2) 5.171±0.571 5.131±0.230 5.287±0.482 0.825
Fiber area (μm2) 8.699±0.747 8.584±0.412 8.641±0.592 0.946
G-ratio 0.571±0.017 0.567±0.033 0.563±0.036 0.958
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Note: Abbreviations include ANA: acellular nerve allograft; p value derived from one-way ANOVA of all groups.

DRG harvested at 120 days from these same animals used in the long-term histological analysis of nerve regeneration revealed elevated expression of Bdnf, cfos, and Galectin among the Control and Traction Neurectomy groups compared to uninjured nerve DRG (Fig. 3). Expression levels from DRG of these genes from the ANA group were statistically decreased (P<0.05) compared to both Control and Traction Neurectomy groups. As well, the expression levels from the ANA group were not statistically different from the uninjured nerve DRG levels. Overall, the data demonstrated the ANA treatment led to changes in gene expression suggestive of a cessation of regeneration and less risk of pain sensitization.

Figure 3. An ANA “cap” alters gene expression within affected DRGs.

Figure 3.

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Expression levels of Bdnf, cfos, and Gal were measured from pooled L4-L5 DRGs harvested at 120 weeks post-injury and treatment. Uninjured nerve DRG served as relative comparison. P values of Control vs other groups are shown. Mean ± standard deviation shown; N=5 for all groups.

Discussion

Surgical treatments to minimize neuroma formation and alleviate resulting pain are desired, where previous studies suggested that the use of ANAs of sufficient length (i.e. 5 cm) could act as a tool to manage neuroma formation by controlling axonal regeneration. However, the results of those studies were limited to conclude if long ANAs could control axon regeneration stably over time, as an endpoint of 5 weeks was used to demonstrate this first proof of principle of axon growth arrest25. In this current work, our studies confirmed that long ANAs successfully “capped” transected nerves arresting axon growth for at least 5 months and altered DRG gene expression suggesting that axon regeneration is permanently arrested within the ANA.

Previous mechanistic studies have found a major role for the central influence of pain following nerve injury 36,37. In this regard, axons send retrograde signals to the neuron cell body during active regeneration38. Long ANAs have been shown to inhibit regeneration23,39. We found, consistent with our previous data, that long ANAs inhibited axon regeneration. In fact, no myelinated axons were detected in the most terminal region (distal segment). Overall, these data demonstrate that long ANAs were effective to arrest axon growth in a controlled manner, which is a major component to potentially change gene expression within upstream neurons, such as sensory neuron bodies within DRG.

Along with controlled termination of axon regeneration, there is a desire to modify gene expression upstream in neuronal bodies to alter pain sensitization. Following peripheral nerve injury, there is an orchestrated upregulation of regenerative associated genes in the neuron cell body40. These genes include BDNF, NGF, Galectin, cFOS, iNOS, CCL2, and others41-46. While upregulation of these genes is important for regeneration of peripheral nerve, they can also cause sensitization to stimuli, leading to pain47. We found that ANA treated nerves had reduced expression of select regenerative associated genes Bdnf, cfos, and Galectin. Conversely, the control and traction neurectomy groups did not hinder axonal regeneration, while in the same instance contained DRG expressing levels of the genes significantly greater than uninjured nerve DRG. Therefore, these data suggest a direct link between arresting axon regeneration and modulating upstream gene expression. While limited at present, this link suggests that arresting axon regeneration could have an impact on pain and/or pain sensitization by modulating upstream gene expression in neuronal bodies. Future studies will need to address whether this hypothetical link exists and can use a long ANA “cap” as a tool to address this new hypothesis.

Several limitations exist in these studies. First, no evaluation for pain-related behaviors was performed. This obstacle is a common challenge in preclinical studies, as pain-related behaviors can be subjective to measure in rodents and are sensitive to the injury model chosen. The sciatic nerve injury model used allows for measurement of axon growth over long time and distances, but may not be the best model to observe neuroma-like pain behaviors from the afflicted nerve, as it is surrounded by muscle tissue and coordinates multiple motor and sensory functions. Future studies could benefit from adapting these studies to an animal model of neuroma that allows for direct measurements from the afflicted nerve, like the tibial neuroma transposition model48. Another limitation of this study is the current methodology used to measure gene expression, which was non-specific to the cell type, as the entire DRG were used for analysis. Thus, changes in gene expression of Bdnf, cfos, and Galectin may be caused by changes of these genes in other cells, such as macrophage that infiltrate DRG following nerve injury. However, since BDNF is a secreted factor, the overall level of expression, rather than source of these factors, may be more important. Future studies could isolate neuronal bodies and other cell populations to measure cell-specific gene expression patterns. Third, these studies considered the prevention of neuroma formation using an end-neuroma model. Other animal models of neuroma, such as a traumatic neuroma in continuity injury model49,50, could be considered. As neuroma prevention vs treatment are different concepts, our studies address the former. Studies to address neuroma treatment using appropriate animal models should be considered for future work. Finally, these studies used an in-house processed ANA as opposed to a commercially available product that could serve as a “cap”. Select commercially available products have demonstrated axon growth across these products for distances over 5 cm51; however, this regeneration occurs when both the proximal and distal nerve is connected to the product. In our studies, the ANA was connected to only the proximal nerve. This modified approach has been previously shown to alter the ability of axons to regenerate across a nerve graft over the same distance comparable distance if connected to a distal nerve52. Therefore, future studies would benefit from determining if this approach using a commercially available product to “cap” the severed proximal nerve can also manage neuroma.

Conclusions

Grafting of long ANAs following nerve transection injury limits regeneration of axons, leading to down-regulation of regenerative associated genes within DRGs that are also associated with pain induction. The use of ANAs for treatment of painful neuroma should therefore be further investigated. Overall, use of ANAs could be a viable clinical option to manage neuroma.

Acknowledgement of Financial and Grant Support:

This work was funded by a research grant from the Department of Defense (DoD) Congressionally Directed Medical Research Programs (CDMRP) Peer Reviewed Orthopaedic Research Program (PRORP) under the Translational Science Award for the project entitled: “Macroscopic Management of Neuromas in Residual Limbs,” W81XWH-15-1-0625 (AMM); and in part by the National Institutes of Neurological Disorders and Stroke of the National Institutes of Health (NIH) under award number R01 NS115960 (MDW) to Washington University.

Footnotes

Conflict of Interest: MDW has been the recipient of sponsored research agreements from Checkpoint Surgical, Inc. and has consulted for Foundry Therapeutics, LLC and The Foundry, LLC. No personal compensation was provided. None of the other authors have any conflicts of interest to disclose.

Disclaimer: The opinions or assertions contained herein are the private ones of the author/speaker and are not to be construed as official or reflecting the views of the Department of Defense, the Uniformed Services University of the Health Sciences or any other agency of the U.S. Government.

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