Peripheral nerve injury results in loss of motor and sensory function distal to the nerve injury and is often permanent in nerve gaps longer than 5 cm. Autologous nerve grafts (nerve autografts) utilize patients’ own nerve tissue from another part of their body to repair the defect and are the gold standard in care. However, there is a limited autologous tissue supply, size mismatch between donor nerve and injured nerve, and morbidity at the site of nerve donation. Decellularized cadaveric nerve tissue alleviates some of these limitations and has demonstrated success clinically. We previously developed an alternative apoptosis-assisted decellularization process for nerve tissue. This new process may result in an ideal scaffold for peripheral nerve regeneration by gently removing cells and antigens while preserving delicate topographical cues. In addition, the apoptosis-assisted process requires less active processing time and is inexpensive. This study examines the utility of apoptosis-decellularized peripheral nerve scaffolds compared to detergent-decellularized peripheral nerve scaffolds and isograft controls in a rat nerve gap model. Results indicate that, at 8 weeks post-injury, apoptosis-decellularized peripheral nerve scaffolds perform similarly to detergent-decellularized and isograft controls in both functional (muscle weight recovery, gait analysis) and histological measures (neurofilament staining, macrophage infiltration). These new apoptosis-decellularized scaffolds hold great promise to provide a less expensive scaffold for nerve injury repair, with the potential to improve nerve regeneration and functional outcomes compared to current detergent-decellularized scaffolds.

Peripheral nerve injury is a serious clinical problem that can affect up to 200,000 people each year in the United States [Madison et al., 1992], resulting in up to 5 million disability days annually [Kelsey, 1997]. After injury to a peripheral nerve, the axon must regrow to the distal target to regain motor and sensory function [Ide, 1996]. For nerve injuries resulting in nerve gaps larger than 3 cm, the gold standard of care is the use of an autologous nerve graft (nerve autograft), which utilizes a sacrificial nerve from another part of the patient’s body [Pabari et al., 2010]. Nerve autografts have demonstrated substantial efficacy in supporting nerve regeneration because of their matched topography (basal lamina tubules) and inherent adhesion molecules (laminin) [Ide et al., 1990; Ide, 1996; Fu and Gordon, 1997]. However, there are many disadvantages of autografts including additional morbidity resulting from nerve harvest for autograft, size mismatch of donor nerve with injured nerve, and highly limited supply of sacrificial donor nerves [Pabari et al., 2010].

Over the past 25 years, an alternative scaffold sourced from cadaveric peripheral nerve tissue has been developed [Ide et al., 1990; Sondell et al., 1998; Hudson et al., 2004a, b]. Allogenic peripheral nerve tissue contains antigenic components; therefore, decellularization methods must be utilized to remove cellular components and avoid implant immune rejection. Decellularized nerve scaffolds are an alternative source to autografts for peripheral nerve repair. These scaffolds inherently contain the key benefits of autografts, such as the basal lamina tubule structure and laminin content, without having the limitations of comorbidity, mismatch, and limited supply [Ide et al., 1990; Sondell et al., 1998; Hudson et al., 2004a, b; Boriani et al., 2017]. Decellularized human nerve scaffolds are commercially available, and clinical studies indicate meaningful recovery in 87.8% of patients with gaps less than 3 cm, with substantial recovery in gaps up to 5 cm [Zhu et al., 2017]. However, at gaps larger than 5 cm, less meaningful recovery is attained resulting in incomplete motor and/or sensory recovery [Zhu et al., 2017; Li et al., 2019]. More ideal decellularization processing holds the potential to increase quality and length of nerve regeneration, while decreasing associated cost.

The majority of current decellularization processes require detergents that damage the tissue structure because of their method of cell lysis. Cell lysis is most commonly performed using a water wash or freeze-thaw to induce cell necrosis, thereby disrupting the cell membrane and distributing cellular debris throughout the surrounding ECM tissue [Sondell et al., 1998; Hudson et al., 2004a; Crapo et al., 2011; Boriani et al., 2017]. Because of this widespread distribution of antigens, damaging detergents are then required for removal. In the case of nerve tissue, many of these detergents can remove the delicate basal lamina tubule structure (which guides nerve regeneration) and laminin protein (which facilitates axonal adhesion) [Sondell et al., 1998; Hudson et al., 2004a]. Furthermore, these detergents can be expensive and the process can be labor-intensive [Sondell et al., 1998; Krekoski et al., 2001; Hudson et al., 2004a, b]. In addition, the use of detergent-decellularized nerve tissue has resulted in highly variable outcomes and immunological tolerance both in vitro and in vivo depending on type of detergents used for processing and residual compounds remaining [Hudson et al., 2004b; Brown et al., 2009; Szynkaruk et al., 2013; White et al., 2017; Lovati et al., 2018].

Our group recently developed and patented an alternative decellularization process that lyses cells by leveraging cell apoptosis to initiate programmed cell death as opposed to necrosis [Cornelison et al., 2018; Schmidt et al., 2018]. During apoptosis, cells detach from the matrix and fragment into enclosed apoptotic bodies, which may be subsequently easily washed from the matrix [Bourgine et al., 2013]. Our recent work using camptothecin to induce apoptosis demonstrated apoptosis-assisted decellularization is able to remove >95% of DNA, while maintaining a delicate basal lamina tubule structure that is comparable to native nerve [Cornelison et al., 2018]. Moreover, there is reduced labor compared to detergent-based decellularization [Cornelison et al., 2018]. This apoptosis-assisted decellularization process is incredibly robust and yields a peripheral nerve scaffold with superior biocompatibility to detergent-based decellularized scaffolds [Cornelison et al., 2018]. Due to this more delicate processing that preserves the basal lamina tubule structure and has better immunological tolerance, the hypothesis of the present study is that apoptosis-decellularized peripheral nerve (ADPN) scaffolds will attain similar nerve regeneration and functional outcomes compared to detergent-decellularized peripheral nerve scaffolds (DDPN) and gold standard allograft in a nerve defect model [Hudson et al., 2004a, b; Nagao et al., 2011]. To test this hypothesis, ADPN, DDPN, and isograft implants were compared to sham animals in a rat sciatic nerve defect over an 8-week time course with histological analysis and functional assessment.

Tissue Harvest and Scaffold Processing

All animal studies were approved by the Institutional Animal Care and Use Committee at the University of Florida and performed in accordance with the Guide for the Care and Use of Laboratory Animals [National Research Council (US) Committee for the Update of the Guide for the Care and Use of Laboratory Animals, 2011]. All animals were housed in polysulfone cages (Allentown) on a ventilated rack with 1/8 inch corn cob bedding (Envigo) and fed diet 2918 (Envigo). Animals were kept on a 12-h reverse light cycle to facilitate behavioral data collection.

This study contained 3 scaffold implant groups: ADPN, DDPN, and fresh Lewis isograft controls. The ADPN and DDPN tissues were harvested from adult male Sprague Dawley rats and processed as previously published [Hudson et al., 2004a, b; Cornelison et al., 2018]. Briefly, the ADPN tissue was treated with 5 μM camptothecin (159732, MP Biomedicals) in DMEM:F12 (D6421, Sigma-Aldrich) with 1% penicillin-streptomycin (15140122, Gibco) at 37°C for 24 h to induce cellular apoptosis. Our prior publication indicates camptothecin induces robust apoptosis to allow subsequent washing of apoptotic bodies from the matrix [Cornelison et al., 2018]. ADPN tissue was subsequently washed with 4× phosphate-buffered saline (PBS) for 24 h to remove apoptotic bodies. Samples were then washed with 1× PBS and treated with DNase (D4527, Sigma-Aldrich) at 75 U/mL for 24 h to remove any remnant DNA. The final step for ADPN samples included treatment with chondroitinase ABC (C3667, Sigma-Aldrich) at 0.2 U/mL for 16 h to remove any neuroinhibitory chondroitin sulfate proteoglycans in the scaffold [Krekoski et al., 2001], followed by 3 washes of 1 h each in 1× PBS. DDPN tissue was treated with sequential detergent washes, including SB-10 (D4266, Sigma Aldrich) and SB-16 (H6883, Sigma-Aldrich)/Triton X-200 (Dow Chemicals) with intermittent 15-min buffered washes. The final treatment of DDPN samples was with chondroitinase ABC wash at 0.2 U/mL for 16 h, followed by 3 washes of 1 h in 1× PBS. ADPN and DDPN tissues were stored in 1× PBS at 4°C until surgical implantation. Macroscopic images of ADPN and DDPN samples were acquired using a digital camera (T3i with Canon EF 100 mm f/2.8 L macro lens, Canon). Fresh Lewis isograft samples were utilized to mimic the clinical gold standard autograft in humans. Isograft was selected because its performance in rats of the same genotype is nearly analogous to autograft without potential implications in functional outcomes due to motor deficit from autograft harvest. Fresh Lewis isografts were harvested from anesthetized donor rats at the time of surgical implantation.

Scaffold Implantation into Sciatic Nerve Defect

A rat 10-mm nerve defect model was utilized to determine differential regeneration potential of ADPN compared to DDPN, isograft, and sham controls [Angius et al., 2012]. Thirty-six male Lewis rats (250–300 g; Charles River Laboratories) were randomly divided into 4 treatment groups. Study end points of 4 weeks (n = 3/group) and 8 weeks (n = 6/group) were selected to observe histological and functional changes associated with nerve regeneration. Rats were anesthetized with 3–4% isoflurane (07-893-1389, Patterson Veterinary) and maintained at 2% isoflurane. Rats were placed in a prone position and the right hind limb was shaved and cleaned with chlorhexidine (07-847-5554, Patterson Veterinary) and sterile saline. Animals were administered buprenorphine (07-891-9756, Patterson Veterinary) subcutaneously at 0.03 mg/kg prior to incision. Using microscope guided dissection, a 2.5–3 cm incision was made longitudinally and the sciatic nerve was exposed using a gluteal approach. The sciatic nerve was dissected from the sciatic notch proximal to the branch point, and an 8-mm defect was created. Interrupted suturing with 10-0 nylon was utilized to implant a 10-mm scaffold of either ADPN, DDPN or fresh Lewis isograft. Macroscopic images post-implantation were acquired using a digital camera (T3i with Canon EF 100 mm f/2.8 L macro lens, Canon). Sham control animals had surgical visualization of the sciatic nerve but no transection. The gluteal muscle was closed using 5-0 Vicryl sutures, and the skin was closed using interrupted 5-0 nylon sutures. All animals recovered on a heating pad. All animals were administered buprenorphine subcutaneously at 0.03 mg/kg for pain every 12 h for 48 h post-surgery.

Muscle Harvest and Weights

The sciatic nerve innervates both the tibialis anterior and the gastrocnemius muscles in the hind limb. After creation of a nerve gap in the sciatic nerve, innervation of the tibialis anterior and the gastrocnemius is severed, which results in a loss of neural input and motor drive and subsequent loss of muscle weight. As regeneration and re-innervation occur over time, this muscle weight is recovered. Therefore, wet weights of the tibialis anterior and the gastrocnemius muscles of each animal were measured and compared to the muscle weights in the contralateral limb at 4 weeks (n = 3 per group) and 8 weeks (n = 6 per group) post-surgery. All subsequent data collection and analysis was conducted by blinded observers.

Gait Data Collection and Analysis

Gait analysis was used to monitor changes in functional outputs over time due to sciatic nerve regeneration through the nerve gap to the distal target. Prior to study commencement, each animal was acclimated to the gait arena (Fig. 3a) for 2 weeks, with a minimum of 1 acclimation session per week. During acclimation, animals were allowed to freely explore the gait arena without any intervention for 10 min. Using an ultra-high speed video camera (Red Lake M3, ITC, San Diego, CA), a minimum of 5 gait trials were collected prior to surgery, and at 2, 4, 6, and 8 weeks post-surgery. A gait trial consisted of an animal freely walking across the track without stopping.

Analysis of gait was performed utilizing previously developed MATLAB code to assess stride length, velocity, temporal symmetry, stance time, imbalance, and duty factor [Jacobs et al., 2018]. In-depth definitions of gait parameters are discussed in Jacobs et al. [2014]. Briefly, stride length is the distance between consecutive strikes of the same foot. Velocity is how fast the animal progresses across the track. Temporal symmetry quantifies synchronicity of foot strikes in a given gait cycle; for example, a temporal symmetry of 0.5 indicates a left foot strike occurs halfway (i.e., 0.5) between 2 right foot strikes. Stance time imbalance is a ratio describing how much time is spent on one foot versus the other. Duty factor is equal to the stance time of a limb divided by the stride time of the limb, giving the percent of time spent in contact with the ground for that limb. Stride length and duty factor are highly correlated to the velocity of gait, and therefore, these data were plotted versus velocity and residual measurements were calculated [Kloefkorn et al., 2015]. Taken together, these parameters provide an indication of abnormalities in each animal’s gait pattern and can describe compensations relating to injury, treatment type, and time.

Immunohistochemical and Histological Analyses

Immunohistochemical and histological analyses were performed at cross sections in the mid-point of the nerve gap or sham nerve to determine degree of nerve regeneration and immune response. At 4 weeks or 8 weeks post-surgery, animals were perfused with 4% paraformaldehyde (441244, Sigma-Aldrich) by cardiac perfusion. After perfusion, animals were euthanized using carbon dioxide inhalation confirmed with bilateral thoracotomy. All samples were secondarily fixed in 4% paraformaldehyde overnight. Samples were then soaked in 30% sucrose for 24 h, embedded in optimal cutting temperature medium (4586, Tissue-Tek), frozen, and cryosectioned (10 μm) before staining. All images were acquired on a Zeiss Axio Observer Z1 microscope with a grayscale camera (Carl Zeiss Microimaging, Inc.). All image analysis was performed using Zen Image Analysis software (Carl Zeiss Microimaging, Inc.).

Nerve regeneration was visualized using an antibody for neurofilament-H (NF-H; RT97, Developmental Studies Hybridoma Bank, 1:500), followed by a secondary fluorescent antibody (Alexa Fluor 568, A11011, Fisher Scientific, 1:500). A complete cross-section at the mid-gap of each sample from each group (ADPN, DDPN, isograft, sham) at each time point (4 or 8 weeks) was imaged and quantified. Within each cross section, the image was converted to binary, and the area of positive NF-H was summated to determine degree of nerve regeneration at the mid-gap in each sample.

Macrophage infiltration was determined by staining with an antibody for all macrophages (CD68, MCA341R, Abd Serotec, 1:200) and secondary fluorescent antibody (Alexa Fluor 568, 1:500). Sections were subsequently stained with 4′,6-diamidino-2-phenylindole (DAPI, D1306, Thermo Fisher Scientific, 1:1,000) to visualize nuclei. A complete cross-section at the mid-gap of each sample from each group (ADPN, DDPN, isograft, sham) at each time point (4 or 8 weeks) was imaged. Total numbers of nuclei were counted. The total positive CD68 staining area was calculated and normalized to the cross-sectional area.

Statistical Methods

All statistical analyses were performed using GraphPad Prism 7 Software. Two-way analysis of variance (ANOVA) was performed for all studies with Tukey’s post-hoc analysis to determine differences between groups or time points, where applicable. Significance was defined at p < 0.05.

Surgical Implantation and Outcomes

All implantation procedures were successful, and all animals recovered from surgery without complications. Isograft, DDPN, and ADPN implants maintained macroscopic integrity out to 8 weeks (Fig. 1d). The DDPN implant appeared noticeably more translucent and smaller in diameter than isograft and ADPN scaffolds at implant (Fig. 1b, c) and explant (Fig. 1d).

Fig. 1.

a Illustration of differential processing of detergent-decellularized peripheral nerves (DDPN) versus apoptosis-decellularized peripheral nerves (ADPN) demonstrating less harsh and costly chemicals, reduced labor, and slightly reduced processing time. b Macroscopic images of DDPN (left) and ADPN (right) after completion of processing prior to trimming to length. c, d Macroscopic images of surgical implant (c) and surgical explant (d) of isograft, DDPN, and ADPN samples. DDPN was noticeably more translucent after processing, at implant and at explant.

Fig. 1.

a Illustration of differential processing of detergent-decellularized peripheral nerves (DDPN) versus apoptosis-decellularized peripheral nerves (ADPN) demonstrating less harsh and costly chemicals, reduced labor, and slightly reduced processing time. b Macroscopic images of DDPN (left) and ADPN (right) after completion of processing prior to trimming to length. c, d Macroscopic images of surgical implant (c) and surgical explant (d) of isograft, DDPN, and ADPN samples. DDPN was noticeably more translucent after processing, at implant and at explant.

Close modal

Functional Recovery

Muscle Weight Recovery

From 4 to 8 weeks, muscle weight recovered in all treated animals (Fig. 2). In all treatment groups at 8 weeks post-injury, animals exhibited a significantly reduced muscle weight in their injured limb muscle weights compared to their contralateral control muscle weights. The tibialis anterior exhibited a significant increase in muscle weight over time in all treatment groups (isograft, DDPN, and ADPN). By 8 weeks post-implantation, these values reached close to 50% of the contralateral limb in all treated groups (Fig. 2a). However, in the gastrocnemius, this temporal increase in muscle weight was only significant in the ADPN treatment group (Fig. 2b). No significant differences were observed between any of the groups at either time point.

Fig. 2.

Recovery of tibialis anterior (a) and gastrocnemius (b) weight over time in treatment groups (isograft, DDPN, ADPN) normalized to the contralateral muscle weight. Significant increases in tibialis anterior muscle weight were observed for all treatment groups between 4 and 8 weeks. However, only the ADPN gastrocnemius muscle exhibited a significant increase between 4 and 8 weeks. No significant differences were observed between any of the groups over time. n= 3/group for 4 weeks and n= 6/group for 8 weeks. Error bars are 95% confidence intervals. * p< 0.05.

Fig. 2.

Recovery of tibialis anterior (a) and gastrocnemius (b) weight over time in treatment groups (isograft, DDPN, ADPN) normalized to the contralateral muscle weight. Significant increases in tibialis anterior muscle weight were observed for all treatment groups between 4 and 8 weeks. However, only the ADPN gastrocnemius muscle exhibited a significant increase between 4 and 8 weeks. No significant differences were observed between any of the groups over time. n= 3/group for 4 weeks and n= 6/group for 8 weeks. Error bars are 95% confidence intervals. * p< 0.05.

Close modal

Gait Analysis

Gait data were analyzed at 2, 4, 6, and 8 weeks post-treatment and compared to sham controls. Gait data from week 2 were highly variable likely due to variability in acute post-operative pain, and therefore this time point was not included in the final analysis. All treated animals (isograft, DDPN, and ADPN) exhibited asymmetry (symmetry <0.5) and imbalance (imbalance >0.0) throughout the course of this study (Fig. 3c, d). By 8 weeks post-surgery, all treatment groups exhibited significantly different temporal symmetry (Fig. 3c) and imbalance (Fig. 3d) compared to the sham control (p < 0.05). Within each treatment group, symmetry and imbalance did not change significantly over time. There were no significant differences between treatment groups (isograft, DDPN, ADPN) in symmetry or imbalance at any time point, demonstrating similar performance of decellularized scaffolds (DDPN and ADPN) to the gold standard isograft.

Fig. 3.

a Illustration of single rat traversing gait chamber with mirror reflection of foot falls. b Plot of velocity-corrected stride length residuals between groups over time indicates that all treatment groups are moving at velocity-corrected stride lengths similar to the sham group by 8 weeks. c, d Temporal symmetry (c) and stance time imbalance (d) data demonstrate that by 8 weeks post-surgery, all treated groups exhibited significantly altered symmetry of the foot-strike sequence and imbalance compared to the sham control, but not compared to other treated groups. n= 6/group. Error bars are 95% confidence intervals. * p< 0.05.

Fig. 3.

a Illustration of single rat traversing gait chamber with mirror reflection of foot falls. b Plot of velocity-corrected stride length residuals between groups over time indicates that all treatment groups are moving at velocity-corrected stride lengths similar to the sham group by 8 weeks. c, d Temporal symmetry (c) and stance time imbalance (d) data demonstrate that by 8 weeks post-surgery, all treated groups exhibited significantly altered symmetry of the foot-strike sequence and imbalance compared to the sham control, but not compared to other treated groups. n= 6/group. Error bars are 95% confidence intervals. * p< 0.05.

Close modal

Residual analysis to correct for velocity effects on stride length revealed that by 8 weeks post-surgery, all treatment groups were not significantly different from the sham control (Fig. 3b). DDPN velocity-corrected stride length was significantly lower than sham at 4 weeks; however, this difference was recovered by the 6-week time point. There were no significant differences between the sham, isograft, or ADPN groups over time. The DDPN velocity-corrected stride length significantly increased from 4 to 6 weeks. These data indicate that all treatment groups used stride lengths that were not significantly different from the sham group by 6 weeks post-implantation and maintained until study completion at 8 weeks.

For the velocity-corrected duty factors, both the right and left hind limbs were significantly different than the sham control at 8 weeks post-surgery for all treatment groups compared (p < 0.05, data not shown). However, no significant differences exist between any of the treatment groups at any time point. These data indicate that all treated animals adopted similar compensatory gaits during injury healing, and these compensatory gaits with ADPN and DDPN are comparable to the isograft gold standard by 8 weeks post-implantation. Additionally, these data demonstrate that an 8-week time frame is not long enough for complete nerve regeneration necessary to restore treated animals to a functional gait similar to the sham control.

Histological Assessment of Regeneration

Degree of Nerve Regrowth

Robust NF-H staining was observed at both 4 and 8 weeks post-implantation in all groups at the mid-gap cross-section (Fig. 4a). By 8 weeks post-implantation, NF-H staining of treatment groups (isograft, DDPN, ADPN) began to exhibit morphology more indicative of neuron bundles (Fig. 4a). Quantification of positive NF-H area revealed there was an increase in NF-H+ area between 4 and 8 weeks; however, this difference was only significant in the DDPN group (Fig. 4b). At 8 weeks post-implantation, the isograft-treated group was the only group with NF-H+ area still significantly reduced from the sham control. DDPN and ADPN NF-H+ area was not significantly different than the sham control.

Fig. 4.

aRepresentative neurofilament-H (NF-H) images at 4 and 8 weeks post-implantation. b Quantified NF-H-positive area at 4 and 8 weeks post-implantation demonstrating increases in NF-H+ staining over time in all groups, but only significant in DDPN. By 8 weeks post-implantation, DDPN and ADPN NF-H+ area is not significantly different than sham, but isograft is still significantly reduced. n= 3/group for 4 weeks and n= 6/group for 8 weeks. Error bars are 95% confidence intervals. * p< 0.05.

Fig. 4.

aRepresentative neurofilament-H (NF-H) images at 4 and 8 weeks post-implantation. b Quantified NF-H-positive area at 4 and 8 weeks post-implantation demonstrating increases in NF-H+ staining over time in all groups, but only significant in DDPN. By 8 weeks post-implantation, DDPN and ADPN NF-H+ area is not significantly different than sham, but isograft is still significantly reduced. n= 3/group for 4 weeks and n= 6/group for 8 weeks. Error bars are 95% confidence intervals. * p< 0.05.

Close modal

Immune Response

Four weeks post-implantation, all treatment groups (isograft, DDPN, ADPN) exhibited significantly increased total nuclear counts compared to the sham control (Fig. 5a). DDPN samples had significantly more nuclei at 4 weeks compared to the ADPN and isograft-treated animals. By 8 weeks post-implantation, only the DDPN treatment groups maintained a significantly increased number of nuclei compared to the sham control (Fig. 5b), suggesting an increase in either recruitment or retention of invading cells. Between 4 and 8 weeks only, the DDPN had a significant decrease in nuclei.

Fig. 5.

aRepresentative images of sham, isograft, DDPN, and ADPN nuclei (blue) and macrophage (red) at 4 weeks post-implantation. b Quantification of total number of nuclei revealed all samples had significantly more infiltrating cells than sham at both time points, however DDPN was also significantly greater infiltrating cells than both isograft and ADPN samples. c Quantification of positive macrophage staining per area revealed that DDPN had significantly increased macrophage staining at 4 and 8 weeks compared to the sham, whereas ADPN was significantly lower than sham at 4 weeks. Error bars are 95% confidence intervals. * p< 0.05.

Fig. 5.

aRepresentative images of sham, isograft, DDPN, and ADPN nuclei (blue) and macrophage (red) at 4 weeks post-implantation. b Quantification of total number of nuclei revealed all samples had significantly more infiltrating cells than sham at both time points, however DDPN was also significantly greater infiltrating cells than both isograft and ADPN samples. c Quantification of positive macrophage staining per area revealed that DDPN had significantly increased macrophage staining at 4 and 8 weeks compared to the sham, whereas ADPN was significantly lower than sham at 4 weeks. Error bars are 95% confidence intervals. * p< 0.05.

Close modal

Our results indicate that ADPN samples perform similarly to gold standard isograft controls in nearly all functional and histological assessments performed in a rat defect of the sciatic nerve over an 8-week time course. These data suggest the ADPN decellularization process and resulting ADPN scaffolds have the potential for widespread use as a scaffold for peripheral nerve injury repair in place of the current clinical gold standard autograft.

Overall, the gait data analysis revealed that by 8 weeks post-implantation all treated groups (isograft, DDPN, ADPN) exhibited velocity-corrected stride lengths that were indistinguishable from the sham control, suggesting no permanent deficits in velocity or stride length from injury. Further, by 8 weeks post-implantation both decellularized scaffolds (DDPN and ADPN) exhibited similar asymmetry, imbalance, and adaptive gait style to the gold standard isograft. However, the treated groups’ asymmetry and imbalance had not returned to sham control values by 8 weeks post-implantation. This deficit is likely caused by incomplete regeneration of the nerve to the distal target. A longer term study is necessary to determine if all gait parameters return to sham levels after complete nerve regeneration occurs. However, the similarity of the ADPN to the gold standard isograft at 8 weeks post-implantation suggests a likelihood of promising long-term outcomes.

The muscle weight data further support the hypothesis of incomplete nerve regeneration to the distal target at 8 weeks. Muscle weight will increase over time as nerve re-innervation occurs and should return to contralateral muscle weight levels after complete re-innervation. Between 4 and 8 weeks, there was a significant increase in muscle weight in all treated groups. By 8 weeks, the ipsilateral tibialis anterior muscle weight of the ADPN sample was 44% of the contralateral limb, and the DDPN sample was 41% of the contralateral limb. A recent review by Lovati et al. [2018] highlights similar findings in 8 papers of sciatic nerve defect with autograft treatment, where muscle weights returned to a mean of 46.4% of the contralateral limb by 6–9 weeks post-implantation. Further, these data demonstrate that in all detergent-decellularized processes, the average muscle weight observed was 35.4% at 6–9 weeks post-implantation [Lovati et al., 2018]. Taken together, our data suggest that the ADPN scaffold has muscle re-innervation that exceeds other detergent decellularization processes and is nearing that of the gold standard autograft. Complete restoration of muscle weight would be expected to occur after complete re-innervation of the muscle.

Histological analysis revealed NF-H+ staining within the nerve gap increased for all treated groups (isograft, DDPN, ADPN) from 4 to 8 weeks. The positive staining/area in the ADPN and DDPN was not significantly different from sham controls by 8 weeks, indicating extensive nerve regeneration across the gap by 8 weeks. Previous work predominantly presents axon diameter to determine axonal regeneration, however, due to extensive regeneration and merging axonal fibers, we analyzed the NF-H+ staining per area of cross-section to determine axonal regeneration. The trends we observed were similar to those in previous decellularized nerve publications with an increase in fiber number over time and isograft or autograft often lagging compared to decellularized scaffolds [Szynkaruk et al., 2013; Lovati et al., 2018]. These data suggest the nerve fibers have completely regenerated across the gap in all treated groups by 8 weeks.

Taken together, the functional and histological data suggest the nerve has not yet fully regenerated to the distal target. These results match previous publications that suggest the nerve reaches the distal target by 6–12 weeks post-implantation as evidenced by increases in muscle mass by 40–60% [Adams et al., 2017; McGrath et al., 2018]. Further, additional studies demonstrate muscle mass begins to reach 80–90% of control muscles by 15–16 weeks post-implantation, but isometric evoked muscle forces still remain less than 50% of the control limb suggesting incomplete functional recovery [di Summa et al., 2011; Moore et al., 2011]. Full innervation and myelination may take as long as 6 months, however, few studies go beyond 16 weeks post-implantation in rodent sciatic nerve defect models.

Major differences in macrophage infiltration were observed between DDPN and ADPN. The DDPN samples exhibited increased total nuclei at both 4 and 8 weeks, and increased macrophage staining at 4 weeks compared to isograft and ADPN samples. These data suggest the DDPN caused a more pronounced immune response than either the isograft or ADPN implants. These differences may be caused by incomplete antigen removal, detergent residuals left behind from the DDPN scaffolds or alteration of ECM that occurred from use of these detergents on DDPN scaffolds. In addition, our results are comparable to those of our previous publication in which the ADPN and DDPN were implanted subcutaneously, and inflammatory infiltrate and stromal remodeling were assessed [Cornelison et al., 2018]. ADPN demonstrated decreased inflammatory infiltrate and stromal remodeling compared to the DDPN at all time points from 1 to 4 weeks [Cornelison et al., 2018]. Very few papers assess the immunological effect of decellularized nerve directly, however, most observe a slight increase in cells during early time points compared to the autograft [Szynkaruk et al., 2013; Lovati et al., 2018]. Recent work has demonstrated differences in the timing of the immune response in autografts compared to allografts which may indicate a different healing cascade [Roballo and Bushman, 2019]. An appropriate host immune response to implanted biomaterials is essential to nerve regeneration, and our data in this paper and our previous publication [Cornelison et al., 2018] suggest the ADPN process may be more tolerated by the immune system.

Some limitations of this study include the limited time course of only 8 weeks and nerve gap length. A longer time course out to 12 weeks or more would likely demonstrate more robust recovery of regeneration and thus allow better assessment of differences between treatment groups. Additional experiments using longer nerve gaps and/or a large animal model may be required to verify clinical relevancy of this scaffold. Further, additional analysis of type of macrophage phenotype and changes to longer time points would allow for more robust assessment of the immune response. Future studies will pursue longer time points and more robust immune assessment as well as an assessment of Schwann cell presence and phenotype. Despite these limitations it is still possible to draw some conclusions from this study that are worthwhile. ADPN grafts demonstrated performance similar to or exceeding DDPN grafts and the clinical gold standard isograft up to 8 weeks post-implantation in a rat nerve defect model. Functionally, injured muscle weights were similar between all treated groups (isograft, ADPN, DDPN) at 4 and 8 weeks post-implantation, and gait results revealed ADPN and DDPN animals had gait asymmetries, imbalances, and adaptive gait patterns not significantly different from isograft animals by 8 weeks. Histologically, nerve re-growth through the gap in ADPN and DDNP samples were not significantly different than sham by 8 weeks, however, they exceeded isograft re-growth. Finally, ADPN exhibited reduced cellular infiltration and macrophage number compared to DDPN samples, suggesting a reduced immune response. The outcomes of the study demonstrate that ADPN nerve grafts perform similarly to the gold standard isograft with potential for enhanced performance in longer term studies due to maintenance of topographical cues and a reduced immune response. Taken together, these data suggest that ADPN grafts have the potential for efficacy in long nerve gaps with comparable outcomes to the gold standard isograft. Together, the novel decellularization process and new nerve grafts have potential to enhance patient lives by increasing nerve regeneration across long gaps, thereby more completely restoring motor and sensory function. Further, this method has the potential to be used in other organ systems as a novel method of tissue decellularization.

This study was approved by the Institutional Animal Care and Use Committee at the University of Florida under approval number 201609443.

Christine E. Schmidt, Rebecca A. Wachs, and R. Chase Cornelison have been awarded a patent for this apoptosis processing approach (Schmidt CE, Wachs RA, and Cornelison RC. “Tissue decellularization methods,” US 10898609 B2). Christine E. Schmidt and Young Hye Song have an additional provisional patent submitted (Agrawal N, Griffin J, Schmidt CE, McCrary MW, Bousalis D, Song YH. “Decellularized tissues, hydrogels thereof, and uses thereof.” US Patent PCT No. US2019/019001, filed February 21, 2019). The authors have no additional conflicts of interest to declare.

This work was supported by the National Science Foundation (CBET 1605223).

Rebecca A. Wachs designed and conducted experiments, analyzed data, wrote manuscript. Steven M. Wellman designed conducted experiments, analyzed data, and edited manuscript. Stacy L. Porvasnik and Young Hye Song performed surgeries, analyzed data, and edited manuscript. Emily H. Lakes and Kyle D. Allen provided gait equipment, helped with gait data collection and analysis, and edited manuscript. R. Chase Cornelison and Christine E. Schmidt helped with experimental design and edited manuscript.

All data generated or analyzed during this study are included in this article. Further enquiries can be directed to the corresponding author.

1.
Adams A, VanDusen K, Kostrominova T, Mertens J, Larkin L. Scaffoldless tissue-engineered nerve conduit promotes peripheral nerve regeneration and functional recovery after tibial nerve injury in rats. Neural Regen Res. 2017;12(9):1529.
2.
Angius D, Wang H, Spinner RJ, Gutierrez-Cotto Y, Yaszemski MJ, Windebank AJ, et al. A systematic review of animal models used to study nerve regeneration in tissue-engineered scaffolds. Biomaterials. 2012;33(32):8034–9.
3.
Boriani F, Fazio N, Fotia C, Savarino L, Nicoli Aldini N, Martini L, et al. A novel technique for decellularization of allogenic nerves and in vivo study of their use for peripheral nerve reconstruction. J Biomed Mater Res A. 2017;105(8):2228–40.
4.
Bourgine PE, Pippenger BE, Todorov A Jr, Tchang L, Martin I. Tissue decellularization by activation of programmed cell death. Biomaterials. 2013;34(26):6099–108.
5.
Brown BN, Valentin JE, Stewart-Akers AM, McCabe GP, Badylak SF. Macrophage phenotype and remodeling outcomes in response to biologic scaffolds with and without a cellular component. Biomaterials. 2009;30(8):1482–91.
6.
Cornelison R, Wellman S, Park J, Porvasnik S, Song Y, Wachs R, et al. Development of an apoptosis-assisted decellularization method for maximal preservation of nerve tissue structure. Acta Biomater. 2018;77:116–26.
7.
Crapo PM, Gilbert TW, Badylak SF. An overview of tissue and whole organ decellularization processes. Biomaterials. 2011;32(12):3233–43.
8.
di Summa PG, Kalbermatten DF, Pralong E, Raffoul W, Kingham PJ, Terenghi G, et al. Long-term in vivo regeneration of peripheral nerves through bioengineered nerve grafts. Neuroscience. 2011;181:278–91.
9.
Fu SY, Gordon T. The cellular and molecular basis of peripheral nerve regeneration. Mol Neurobiol. 1997;14(1‐2):67–116.
10.
Hudson TW, Liu SY, Schmidt CE. Engineering an improved acellular nerve graft via optimized chemical processing. Tissue Eng. 2004a;10(9-10):1346–58.
11.
Hudson TW, Zawko S, Deister C, Lundy S, Hu CY, Lee K, et al. Optimized acellular nerve graft is immunologically tolerated and supports regeneration. Tissue Eng. 2004b;10(11‐12):1641–51.
12.
Ide C. Peripheral nerve regeneration. Neurosci Res. 1996;25(2):101–21.
13.
Ide C, Osawa T, Tohyama K. Nerve regeneration through allogeneic nerve grafts, with special reference to the role of the Schwann cell basal lamina. Prog Neurobiol. 1990;34(1):1–38.
14.
Jacobs BY, Kloefkorn HE, Allen KD. Gait analysis methods for rodent models of osteoarthritis. Curr Pain Headache Rep. 2014;18(10):456.
15.
Jacobs BY, Lakes EH, Reiter AJ, Lake SP, Ham TR, Leipzig ND, et al. The open source GAITOR suite for rodent gait analysis. Sci Rep. 2018;8(1):9797.
16.
Kelsey JL. Upper extremity disorders: frequency, impact, and cost. Churchill Livingstone; 1997.
17.
Kloefkorn HE, Jacobs BY, Loye AM, Allen KD. Spatiotemporal gait compensations following medial collateral ligament and medial meniscus injury in the rat: correlating gait patterns to joint damage. Arthritis Res Ther. 2015;17(1):287.
18.
Krekoski CA, Neubauer D, Zuo J, Muir D. Axonal regeneration into acellular nerve grafts is enhanced by degradation of chondroitin sulfate proteoglycan. J Neurosci. 2001;21(16):6206–13.
19.
Li L, Yang J, Qin B, Wang H, Yang Y, Fang J, et al. Analysis of human acellular nerve allograft combined with contralateral C7 nerve root transfer for restoration of shoulder abduction and elbow flexion in brachial plexus injury: a mean 4-year follow-up. J Neurosurgery. 2020;132(6):1914–24.
20.
Lovati AB, D'Arrigo D, Odella S, Tos P, Geuna S, Raimondo S, et al. Nerve repair using decellularized nerve grafts in rat models. A review of the literature. Front Cell Neurosci. 2018;12:427.
21.
Madison R, Archibald S, Krarup C. Peripheral nerve injury. In: Cohen IK, Diegelman F, Lindblad WJ , editors. Wound healing: biochemical and clinical aspects. Philadelphia: Saunders; 1992. p. 450–80.
22.
McGrath AM, Brohlin M, Wiberg R, Kingham PJ, Novikov LN, Wiberg M, et al. Long-term effects of fibrin conduit with human mesenchymal stem cells and immunosuppression after peripheral nerve repair in a xenogenic model. Cell Med. 2018;10:215517901876032.
23.
Moore AM, Macewan M, Santosa KB, Chenard KE, Ray WZ, Hunter DA, et al. Acellular nerve allografts in peripheral nerve regeneration: a comparative study. Muscle Nerve. 2011;44(2):221–34.
24.
Nagao RJ, Lundy S, Khaing ZZ, Schmidt CE. Functional characterization of optimized acellular peripheral nerve graft in a rat sciatic nerve injury model. Neurol Res. 2011;33(6):600–8.
25.
National Research Council (US) Committee for the Update of the Guide for the Care and Use of Laboratory Animals. Guide for the Care and Use of Laboratory Animals, ed 8. Washington (DC): National Academies Press (US); 2011.
26.
Pabari A, Yang SY, Seifalian AM, Mosahebi A. Modern surgical management of peripheral nerve gap. J Plast Reconstr Aesthet Surg. 2010;63(12):1941–8.
27.
Roballo KCS, Bushman J. Evaluation of the host immune response and functional recovery in peripheral nerve autografts and allografts. Transpl Immunol. 2019;53:61–71.
28.
Schmidt CE, Wachs RA, Cornelison RC. Tissue decellularization methods. Google Patents; 2018.
29.
Sondell M, Lundborg G, Kanje M. Regeneration of the rat sciatic nerve into allografts made acellular through chemical extraction. Brain Res. 1998;795(1‐2):44–54.
30.
Szynkaruk M, Kemp SW, Wood MD, Gordon T, Borschel GH. Experimental and clinical evidence for use of decellularized nerve allografts in peripheral nerve gap reconstruction. Tissue Eng Part B Rev. 2013;19(1):83–96.
31.
White LJ, Taylor AJ, Faulk DM, Keane TJ, Saldin LT, Reing JE, et al. The impact of detergents on the tissue decellularization process: a ToF-SIMS study. Acta Biomater. 2017;50:207–19.
32.
Zhu S, Liu J, Zheng C, Gu L, Zhu Q, Xiang J, et al. Analysis of human acellular nerve allograft reconstruction of 64 injured nerves in the hand and upper extremity: a 3 year follow‐up study. J Tissue Eng Regen Med. 2017;11(8):2314–22.