A clinically relevant mouse model for traumatic peripheral nerve injury and repair

A clinically relevant mouse model for traumatic peripheral nerve injury and repair | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article A clinically relevant mouse model for traumatic peripheral nerve injury and repair Walter Unterberger, Lucas Rubisoier, Felix Schöffing, Egon Demetz, and 3 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-9254882/v1 This work is licensed under a CC BY 4.0 License Status: Under Review Version 1 posted 8 You are reading this latest preprint version Abstract In contrast to axons of the central nervous system, peripheral nerve fibers regenerate after traumatic injury, but often not to a clinically satisfactory extent, as only a small proportion of axons achieve long-distance regrowth. While autologous nerve transplantation remains the gold standard for tension-free repair, advances in biomaterials have led to increasing clinical use of synthetic nerve guidance conduits. Despite these developments, experimental research on traumatic peripheral nerve injury (PNI) continues to rely on crush lesions and direct end-to-end repair models, both of which have limited clinical relevance. Here, we present a mouse model of traumatic PNI that closely mimics clinical conditions and enables translationally relevant investigations. Our model combines sciatic nerve transection with conduit-based repair using a commercially available, standardized chitosan nerve guidance conduit that is longitudinally opened prior to implantation. This design allows for the direct application of soluble bioactive compounds to the lesion site. We further detail methods for visualizing and quantifying axonal regeneration at single-fiber resolution within the conduit and provide a time-course analysis of axonal regrowth and macrophage dynamics during Wallerian degeneration by making use of transgenic reporter mice. Functional recovery is assessed using sensory and motor performance tests, and time windows for applying bioactive compounds are defined. Together, this model recapitulates key features of clinical nerve injury and repair, and provides a platform for preclinical testing of regenerative therapies. We propose it as a standardized reference model for experimental PNI research. Biological sciences/Biotechnology Physical sciences/Engineering Health sciences/Medical research Biological sciences/Neuroscience Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 INTRODUCTION Unlike axons in the central nervous system (CNS), peripheral nerves are often assumed to regenerate spontaneously without the need for extensive medical intervention. While axonal regrowth does occur following Wallerian degeneration in the peripheral nervous system (PNS), this regenerative capacity is largely restricted to mild injuries such as compression or crush lesions. In contrast, traumatic nerve injuries involving complete transection do not regenerate spontaneously (Phillips et al., 2022 ). These severe injuries require surgical repair, and even then, functional outcomes are often poor, frequently resulting in lifelong sensory and motor deficits (Bergmeister et al., 2020 ; Ciaramitaro et al., 2010 ). A major limitation to recovery is the failure of long-distance axonal regeneration. Most severed axons either fail to regenerate or abort growth prematurely, often accompanied by axonal sprouting and neuroma formation (Allodi et al., 2012 ; Zhao et al., 2023 ). Traumatic peripheral nerve injury (PNI) most commonly affects the upper extremities and is frequently caused by occupational or traffic accidents (Bergmeister et al., 2020 ; Noble et al., 1998 ). In addition, warfare-related injuries are increasing in prevalence and often involve complex, multi-site nerve damage affecting upper and lower extremities simultaneously (Eckhoff et al., 2021 ; Howard et al., 2025 ). The current gold standard for tension-free nerve repair is autologous nerve grafting, typically using a segment of the sural nerve to bridge the lesion gap (Safa & Buncke, 2016 ). While this approach supports axonal regrowth, it is limited by donor-site morbidity and graft availability. Synthetic nerve guidance conduits have emerged as a promising alternative, offering standardized quality, unrestricted availability, and favorable biocompatibility (Redolfi Riva et al., 2024 ; Zhou et al., 2024 ). Despite these clinical advances, experimental models of PNI remain poorly aligned with clinical practice. Most studies rely on crush lesions or nerve transection followed by direct coaptation, neither of which adequately reflects current surgical approaches (Heinzel et al., 2020 ; Vela et al., 2020 ). Crush injuries are typically not treated surgically, and direct end-to-end repair is often avoided clinically due to tension and suboptimal outcomes compared to conduit-based repair (Berger & Millesi, 1978 ; Leis et al., 2024 ; M et al., 2014). Moreover, these models do not permit controlled local delivery of bioactive compounds to evaluate their therapeutic potential. In addition, crush lesions may confound interpretation, as a subset of axons can survive the injury and avoid Wallerian degeneration (Kim et al., 2023 ). To address these limitations, we developed a mouse model of traumatic PNI that more closely replicates clinically relevant injury and repair paradigms. The model combines sciatic nerve transection with conduit-based repair using a longitudinally opened chitosan nerve guidance conduit. This procedure involves both surgical intervention and reconstruction. Although microsurgical procedures are technically more demanding in mice than in rats due to smaller anatomic structures, the mouse offers access to a wide range of transgenic reporter lines (Cai et al., 2013 ; Feng et al., 2000 ). Chitosan conduits are commercially available, standardized, and increasingly used in clinical settings due to their favorable biocompatibility (Bocker et al., 2022 ; Meyer et al., 2016 ; Neubrech et al., 2016 ; Neubrech et al., 2018 ). This approach enables direct application of soluble bioactive compounds to the lesion site and their evaluation in a controlled microenvironment. Using an adapted imaging strategy for synthetic conduits, we are able to visualize and quantify regenerating axons at single-fiber resolution within the conduit and distal stump (Fogli et al., 2019 ). Here, we present this model in detail, describe microsurgical techniques and conduit processing, characterize axonal regeneration and macrophage dynamics over a time course from 2 days to 6 weeks post-injury, and assess functional recovery using sensory and motor assays. MATERIALS AND METHODS Animals Adult (2–6 month-old) female GFP-M mice (Feng et al., 2000 ) or Cx3cr1 CreERT2 ::ROSA26 tdTomatofl/fl mice (Petzer et al., 2018 ) were used for sciatic nerve surgery and imaging experiments. Adult (3 month-old) female C57BL/6N mice from Charles River Laboratories were used for sciatic nerve surgery and sensory-motor experiments. Animals were housed in individually ventilated cages (IVCs) in a temperature- and humidity-controlled room under a 12-hour light/12-hour dark cycle with access to food and water ad libitum . For time course experiments, homozygous GFP-M mice were crossbred with homozygous Cx3cr1 CreERT2 ::ROSA26 tdTomatofl/fl mice to obtain a heterozygous double GFP/Tomato reporter line. For imaging experiments involving the Cx3cr1 CreERT2 ::ROSA26 tdTomatofl/fl mouse line, normal food was exchanged with pellets containing tamoxifen (360 mg Tamoxifen citrate/kg; sniff #A11570360) one week before nerve-conduit dissection. All experimental protocols were approved by the Austrian Federal Ministry of Science and Research (GZ #2021 − 0.406.859) and complied with the European Convention for the Protection of Vertebrate Animals Used for Experimental and other Scientific Purposes (ETS no. 123). Sciatic nerve transection and conduit surgery Sterility was maintained throughout all surgical procedures. To provide a guidance structure for axonal regeneration, a Reaxon® chitosan conduit (2.1 mm × 14 mm) was used (Medovent #RD121). Prior to NaBH₄ treatment, the conduit was softened in distilled water and cut into six equal pieces by first dividing it transversally into three 4-mm tubes and subsequently splitting each tube longitudinally. The resulting conduit segments measured approximately 4 × 3 mm. If not used immediately, conduits were dried for 24 h in a laminar flow hood and stored in sealed Petri dishes. To reduce autofluorescence, conduits were incubated in 10 mg/mL NaBH₄ (Merck #213462) in 1× PBS for at least 40 h. Conduits were then washed five times by inversion in 40 mL sterile 1× PBS in a 50 mL tube under sterile conditions. After washing, conduits were stored in sterile 1× PBS in sealed Petri dishes. For pain management, a dual analgesic regimen was applied. Beginning one day before surgery, mice received 1.25 mg/mL metamizole (Metagelan®; G.L. Pharma #137760) in the drinking water. To improve palatability, the solution was supplemented with apple juice. Treatment was continued until postoperative day 2 and then replaced with autoclaved tap water. In addition, mice received daily subcutaneous injections of carprofen (0.5 mg/mL in sterile 1× PBS; Rimadyl®; Zoetis #10000319) at 10 µL per g body weight from the day of surgery until postoperative day 3. Anaesthesia was induced and maintained with isoflurane (IsoFlo®; Zoetis #400136.00.00) delivered via a mask at a flow rate of 0.3 L/min. Isoflurane concentration was maintained at 2.5% until nerve transection and subsequently reduced to 2.0% to limit overall anaesthetic exposure. Body temperature was continuously monitored using a rectal probe and maintained with a heating plate set to 37°C. After induction of anaesthesia and analgesic administration, the right hind limb was shaved, and the surgical area was disinfected with an iodine-containing antiseptic. A neurosurgical microscope (S100/OPMI pico; Carl Zeiss Meditec) and specialized microsurgical instruments were used for all surgical procedures. The right hind limb was extended and fixed to the surgical table to facilitate visualization of the femur. A longitudinal 4–5 mm skin incision was made just below the caudal border of the femur, and the subcutaneous tissue was separated using scissors. The underlying muscles were bluntly separated at the natural connective tissue plane to expose the sciatic nerve. Connective tissue surrounding the nerve was carefully removed, and the nerve was transected with spring scissors in a single cut. Following transection, the distal stump was sutured to the inner wall of the conduit using an Ethilon® 9 − 0 (Johnson & Johnson #EH7448G) suture. The needle was inserted paramedially from the outside of the conduit to the inside (approximately 1.5 mm from the edge), passed through the epineurium and back through the conduit wall, where a knot was tied (with three throws). This creates a loop that safely secures the nerve stump to the conduit. The same procedure was performed for the proximal stump, leaving a gap between the stumps of up to 1 mm. To close the conduit after fixation of the nerve stumps, the conduit was rotated within the surgical field to expose its corners. The distal end was first closed by connecting the conduit corners with a simple interrupted suture. A second distal suture was placed to enable reliable identification of the sample orientation after extraction of the nerve-conduit assembly. The proximal corners were then sutured in the same manner. This configuration left a small opening between the sutures that allowed matrix instillation. To minimize matrix loss during final closure, the closing suture was pre-positioned before matrix instillation. The viscous matrix, which was prepared freshly on the day of surgery, was instilled using a slightly cut-off tip to increase the opening diameter. The matrix composition was: 50 mM HEPES, 1x PBS, 0.85x PuraMatrix™ (Corning #CLS354250). The conduit was then fully closed with a simple interrupted suture. Finally, the nerve–conduit assembly was rotated back into its original position, and the muscles were closed using a mattress suture with Permahand® 5 − 0 (Johnson & Johnson #K890H) sutures to minimize tension. The skin was closed with either interrupted buried sutures or surgical clips. Tissue preparation and clearing Two weeks after surgery, mice were euthanized and peripherally perfused with 1× PBS and 4% PFA via an intravenous cannula inserted into the vena cava. Prior to perfusion, mice were deeply anaesthetized by intraperitoneal injection of approximately 100 µL of a 2:1 ketamine/xylacine mixture (100 mg/ml Ketasol®, aniMedica #8-00173; 20 mg/ml Xylasol®, aniMedica #8-00178). After insertion of the cannula, the right femoral vein (vena femoralis dextra) was punctured to allow blood drainage and facilitate removal of intravascular hemoglobin. Following perfusion, the skin above the operated area was removed, and the tissue was fixed in 4% paraformaldehyde at 4°C for 24 h. The nerve-conduit assembly was then carefully excised while preserving as much surrounding nerve tissue as possible. Samples were washed twice in 50 mL 1× PBS for 2 h on a tube roller wrapped in aluminum foil. Tissue clearing was performed using a modified CUBIC protocol (Susaki et al., 2014 ), in which reagent ratios were maintained while incubation times were adjusted. Briefly, CUBIC-1 was freshly prepared by dissolving urea (25 wt%), Quadrol® (25 wt%), and Triton™ X-100 (15 wt%) in distilled water (35 wt%) by stirring at 40–50°C, followed by degassing. CUBIC-2 was prepared similarly by dissolving sucrose (50 wt%), urea (25 wt%), triethanolamine (10 wt%), and Triton™ X-100 (0.1 vol%) in distilled water (15 wt%). All reagents were obtained from Merck. To minimize signal loss, samples were protected from light by wrapping tubes in aluminum foil. Samples were incubated in 10 mL CUBIC-1 on an incubation shaker at 55 rpm and 37°C for 2 days, with the reagent refreshed after the first day. Samples were subsequently washed three times for 2 h in 40 mL 1× PBS on a tube roller. During the washing steps, 50 ml flasks and continuous rotation were required to prevent precipitation within the nerve-conduit assembly. After washing, samples were transferred to 10 mL CUBIC-2 and incubated for 2 days while shaking at 55 rpm and 37°C. On day 5, samples were transferred to 2 mL microcentrifuge tubes filled with CUBIC-2 and shipped overnight for imaging (Schweigreiter et al., 2020 ), resulting in a total CUBIC-2 incubation time of approximately 2.5 days. Confocal microscopy Nerve–conduit assemblies were placed in sealed custom-made imaging chambers filled with CUBIC-2 solution. The chambers were based on standard microscope slide dimensions and contained a hollow cavity (15 mm diameter, 2 mm depth). Standard coverslips sealed with desiccator grease were used to close the cavity. Up to four nerves were mounted in parallel using a holder compatible with the Caco-2 standard well-plate insert of the motorized microscope stage. Imaging was performed using a Nikon A1R confocal microscope mounted on a Nikon Eclipse Ni-E microscope body. A Plan Apo 10× objective (NA 0.45, working distance 4.0 mm) was used to capture the full width of the nerve. Green and red fluorescence were excited using 488 nm and 561 nm lasers, respectively, with emission detected through 515/30 nm and 593/46 nm filters. To image the entire nerve–conduit assembly, tiles centered on the middle of the conduit were acquired with 15% overlap. Individual tiles were acquired at 1024 × 1024 pixels with a pixel size of 1.24 µm. Z-stacks of up to 800 µm were acquired per nerve with a step size of 2.45 µm. Multiple stage positions were defined for each sample holder to enable consecutive imaging of up to four nerves in a single session. Image processing Because the chitosan conduit retained residual autofluorescence despite quenching, a multistep image analysis workflow was implemented. First, image stacks were aligned and cropped to a standardized field of view using Fiji / ImageJ (v1.54p) (Schindelin et al., 2012 ). For segmentation, pixel classification was performed using ilastik (v1.4.0) (Berg et al., 2019 ; Kreshuk et al., 2011 ). Two classes were defined to distinguish axonal structures from everything else, including the conduit material, debris, and background. The resulting probability maps were exported to Fiji and thresholded to obtain a more stringent 3D binary mask (confidence > 60–95%), where background was 0 and detected axons were 1. Subsequently, the mask was multiplied with the original grayscale image stack to preserve the original fluorescence intensities while removing background and conduit autofluorescence. The processed image stacks were subsequently resliced to generate en face views of the nerve. Nerve fibers were detected in each resliced section using a custom GA3 spot-detection workflow implemented in NIS-Elements AR (v6.20.01), and the resulting counts were exported to Microsoft Excel. This approach generated a progressive, micrometer-resolved count of nerve fibers per pixel row along the length of the nerve-conduit assembly. von Frey test Mechanical sensitivity was assessed using a dynamic plantar aesthesiometer (Ugo Basile; model 37450) equipped with the touch stimulator (cat. no. 37400-002) and base platform (model 37000-003). On each testing day, the device was calibrated according to the manufacturer’s instructions using 5 g and 50 g calibration weights. Mice were first allowed to habituate to the testing room for at least 30 min in their home cages. Animals were then transferred to the testing chambers and allowed to habituate for approximately 2 h prior to testing. Each hind paw was stimulated three times with a gradually increasing force ranging from 0 to 10 g over a 20 s ramp period. Consecutive stimulations were separated by at least 120 s. For each hind paw, the mean of the three measurements was calculated. Outcome parameters were the force and latency at which the animal withdrew the hind paw. Baseline measurements were performed approximately one week before surgery. Postoperative measurements were conducted in the 4th and 6th week after surgery. CatWalk test Gait and paw-print parameters were analyzed using the CatWalk XT system (v10.6; Noldus Information Technology BV), largely following the protocol described in (Moritz et al., 2019 ) with minor modifications. After a 2-week habituation period to the facility, experiments were initiated. Mice were not trained to reach a goal box. Instead, after each run, they were returned to the beginning of the walkway or, after completing three compliant runs, put back to their home cage. Experiments were conducted in the dark. To improve paw-print contrast, mice were briefly placed in a cage containing a wet paper towel (tap water) immediately before testing to moisten the paws. The walkway floor and walls were cleaned with household cleaner between animals, or whenever urine or fecal boli were deposited on the walkway. For baseline measurements, mice were trained in the CatWalk system at 10 weeks of age on two consecutive days and tested two days later to generate the “before surgery” dataset. The following acquisition settings were used during training and testing. The camera was positioned 27 cm below the walkway. Minimum and maximum run durations were set to 0.5 s and 5 s, respectively. The minimum number of compliant runs per animal was set to three, and the maximum allowed speed variation was set to 60%. Camera gain was set to 12.9 dB, the green intensity threshold to 0.1, the red ceiling illumination to 17.7 V, and the green walkway illumination to 16.5 V. Postoperative measurements were performed in the 5th and 7th week after surgery to generate the corresponding post-surgery datasets. For paw-print analysis, runs were included with a minimum number of consecutive steps of 5, average speed from 10.0 to 40.0 cm/second, and maximum allowed speed variation of 60%. For gait cycle analysis, runs were included with a minimum number of consecutive steps of 10, average speed from 10.0 to 40.0 cm/second, and maximum allowed speed variation of 50%. Paw-prints were controlled manually to ensure correct identification of the affected right hind paw. Statistical analysis Statistical analyses were performed using GraphPad Prism (v10.6.1; GraphPad Software). Data were assessed for normality using the Kolmogorov–Smirnov test, and no significant deviations from normal distribution were detected. For paw-print parameters obtained from CatWalk analyses, two-way analysis of variance (ANOVA) was used to evaluate the effects of experimental group (left versus right hind paw) and time point. When missing values resulted in unequal sample sizes, a mixed-effects model was applied instead of two-way ANOVA. Gait parameters were analyzed using one-way ANOVA. When significant effects were detected, pairwise comparisons were performed using Bonferroni’s multiple-comparisons post hoc test. A significance threshold of p < 0.05 was applied. Exact sample sizes and data presentation (mean ± SD) are indicated in the respective figure legends. No statistical methods were used to predetermine sample size. RESULTS Chitosan conduit-based repair of the transected sciatic nerve and tissue clearing The sciatic nerve was transected in the upper hind limb proximal to its trifurcation into the peroneal, tibial, and sural branches. Immediately after transection, the stumps retracted due to intrinsic axial tension. Commercial chitosan nerve guidance conduits are designed for clinical use, and even the smallest available diameter (2.1 mm) exceeds that of the mouse sciatic nerve. To adapt the conduit for mice, we cut it in half and positioned one half beneath the nerve stumps. Before placement, the conduit was soaked in NaBH₄ to quench autofluorescence. Each stump was fixed to the conduit with tensionless epineural sutures, leaving a 0.5–1 mm gap between the stumps. The conduit was subsequently closed at the ends and the center, and the cavity filled with PuraMatrix hydrogel, which can be supplemented with biologically active compounds to promote axonal regeneration and functional recovery (Fig. 1 A-D). Following wound closure, animals were allowed to recover for a period appropriate to the readout: 2 weeks for imaging axonal regeneration, when the majority of regenerating axons had crossed the gap and pioneering fibers had begun extending into the distal stump, and 4–7 weeks for functional assessments of sensory and motor recovery. For imaging, animals were perfused with PBS followed by PFA. Perfusion was necessary to flush blood from the lesion site, particularly in cases of surgery-induced vasculogenesis. Because conventional cardiac perfusion does not efficiently clear blood from the upper hind limb, we implemented a “peripheral perfusion” protocol, with perfusion through the vena cava and drainage via the femoral vein. The nerve-conduit assembly was then dissected, immersion-fixed for 24 hours, and washed thoroughly with PBS. At this stage, samples could be stored at 4°C in the dark to preserve GFP fluorescence. Tissue clearing was performed using an adapted CUBIC protocol (Susaki et al., 2014 ), with 2 days in CUBIC 1 followed by 2.5 days in CUBIC 2 (Fig. 1 E-F). Extended incubation in CUBIC 2 was found to diminish GFP fluorescence; imaging was therefore performed without delay. (A) Sciatic nerve immediately after transection. Retraction of the proximal and distal nerve stumps is visible. (B) Placement of the chitosan conduit beneath the transected nerve. Bulbous nerve ends are formed after transection. The proximal and distal nerve stumps are fixed to the conduit using epineural sutures without tension, leaving a gap of approximately 0.5–1 mm between the nerve ends. (C) The conduit is wrapped and closed at the ends. Before closing the central portion, a hydrogel matrix is pipetted into the conduit; regeneration-promoting agents can be added to this matrix. (D) The conduit is fully closed, and the surgical site is subsequently closed with muscle and skin sutures. (E) Two weeks after surgery, animals were euthanized, and the nerve-conduit assembly was dissected following peripheral perfusion and fixation. (F) The nerve-conduit assembly after the CUBIC-based clearing protocol. Scale bar, 500 µm. Imaging and quantifying regenerative axonal growth within the nerve-conduit assembly While light sheet fluorescence microscopy (LSFM) is commonly used for brain and nerve imaging, prior work indicated that LSFM is unsuitable for synthetic nerve conduits due to light scattering by the conduit walls, which prevents visualization of axons within the conduit (Fogli et al., 2019 ). We therefore used confocal microscopy with a 10x long-working distance objective to image the centimeter-scale nerve-conduit assemblies. Confocal imaging provided high spatial resolution, enabling precise quantification of individual fibers, including small-diameter C-type axons that may be unresolved by LSFM. A multi-position holder allowed automated imaging of up to four samples in a single overnight session. Reconstructed 3D image stacks were processed using ilastik, a machine learning–based classification and segmentation tool (Berg et al., 2019 ). Residual conduit autofluorescence and background were subtracted by thresholding to isolate axonal GFP signals. To quantify regeneration, the z-stack was re-sliced orthogonally for an en face view of the nerve. Individual axons were detected and counted along the nerve axis using a spot detection algorithm, highlighting that only a small fraction of axons initiate regenerative growth — one reason for poor functional recovery after traumatic PNI (Fig. 2 A-E). (A) Combined transmission light and EGFP fluorescence confocal imaging of the nerve-conduit assembly two weeks after PNI using a 10x long-working distance objective. A GFP-M mouse, which expresses GFP in a mosaic pattern in sensory and motor fibers (approximately 1% of all axons; (Feng et al., 2000 )), was used. Dashed white lines indicate the proximal and distal nerve stumps within the conduit. (B) GFP fluorescence channel alone. The yellow arrow indicates the typical drop in fluorescence at the conduit edge. (C) Maximum intensity projection (MIP) of the grayscale image stack after pixel classification and subtraction of conduit autofluorescence. (D) MIP of the grayscale image stack after pixel classification and thresholding. Regenerating nerve fibers in the distal nerve stump are indicated by yellow arrowheads. Orthogonal slices of the proximal and distal stumps are shown at positions marked by blue lines. (E) Quantification of GFP-positive intersections per orthogonal slice, corresponding to axon number, along the nerve-conduit axis. The yellow arrow indicates the typical drop in fluorescence at the conduit edge. The light-blue curve presents data at each incremental position. The dark-blue curve has been smoothed over 50 µm intervals. Scale bar, 500 µm. Time course of axon regeneration and macrophage dynamics after nerve transection and conduit-based repair While skin reinnervation after sciatic nerve crush has been investigated immunohistochemically (Navarro et al., 1997 ), systematic morphological data for nerve transection followed by conduit repair are lacking. To address this, we performed an imaging time series at intervals ranging from 2 days to 6 weeks post-injury. Macrophages play a key role in Wallerian degeneration and axonal regeneration, clearing myelin and axonal debris and, together with repair Schwann cells, contributing to the formation of bands of Büngner (Li et al., 2022 ). To visualize macrophage dynamics, we crossed GFP-M mice with Cx3cr1 CreERT2 ::ROSA26 tdTomatofl/fl animals, enabling tamoxifen-inducible tdTomato expression in the Cx3cr1⁺ macrophage lineage (Petzer et al., 2018 ). Axonal reorganization was first evident 2 days post-surgery, including growth cone formation, consistent with the approximately 48-hour latency required for neurons to reprogram into a regenerative state (Griffin et al., 2010 ). Bulk axonal regrowth initiated after 1 week, once Wallerian degeneration was largely complete. By 2 weeks, most axons had crossed the gap, with pioneering fibers extending into the distal stump; this pattern was consolidated at 4 and 6 weeks. Notably, chitosan autofluorescence quenching diminished at later time points, particularly in the 488 nm channel, limiting observations beyond 2 weeks (Fig. 3 A). In intact nerves, a resident macrophage population was observed. Following sciatic nerve injury (SNI), macrophage recruitment commenced within 1 week, peaking around 2 weeks in the conduit and reaching maximal density in the distal stump by 4 weeks. By 6 weeks, macrophage numbers had begun to decline, indicating clearance of macrophages from the lesion site (Fig. 3 B). In summary, our time course captures all major events after nerve transection and conduit repair: macrophage recruitment within 1 week, initiation of axonal regrowth after 1 week, consolidation of axonal regeneration by 2 weeks, and onset of macrophage clearance by 6 weeks. We identify 2 weeks post-lesion as the optimal time point to analyze axonal regeneration within the conduit, coinciding with maximal macrophage density (Fig. 3 C). A double-reporter mouse line generated by crossing GFP-M mice with the macrophage-specific tamoxifen-inducible tdTomato reporter line Cx3cr1 CreERT2 ::ROSA26 tdTomatofl/fl was used to visualize axon and macrophage dynamics for up to six weeks after SNI. MIPs are shown for each time point. (A) Progressive axonal fragmentation in the distal nerve stump, indicative of Wallerian degeneration, is visible at 4 days and 1 week after surgery (white arrow heads). By 2 weeks after SNI, all transected axons have been degraded, and the first regenerative fibers extend into the distal nerve stump (white arrows). (B) Macrophages are present in the intact nerve and increase in number several days after SNI due to recruitment. By 6 weeks after SNI, macrophages are largely cleared from the lesion site. (C) Combined axonal and macrophage signals. Scale bar, 500 µm. p.s., post-surgery. Sensory-motor recovery after nerve transection and conduit-based repair Although axonal regeneration is actively ongoing within the conduit at 2 weeks, target reinnervation takes additional time. Functional recovery in this model has not been previously characterized. Based on in vivo axonal growth rates in young adult mice, approximately 1.9 mm/day for motor fibers and approximately 2.9 mm/day for sensory fibers (Verdu et al., 2000 ), sensory axons are expected to reinnervate targets around 3 weeks post-injury, and motor fibers by about 4 weeks. Accounting for the initial 1–2 week phase required to traverse the gap and enter the distal stump, we selected a 4–7 week window to monitor sensory-motor performance, which should also align with the period most suitable for evaluating regenerative interventions at the lesion site. Sensory recovery was assessed longitudinally using the von Frey test from baseline to 6 weeks post-surgery. In the 6th week, operated animals exhibited responses comparable to baseline, whereas in the 4th week, the operated right hind limb showed significantly reduced responsiveness relative to the intact left hind limb for both withdrawal force and latency (Fig. 4 A, B). These results indicate that 3–4 weeks post-lesion is the optimal window to evaluate sensory recovery. Longitudinal von Frey testing was performed from baseline to the 6th week post-surgery. (A) Force required to elicit hind paw withdrawal. (B) Latency to hind paw withdrawal. Individual data points represent the mean of three measurements per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 6 in the 4th week post-surgery; n = 6 in the 6th week post-surgery. Statistical significance and p-value are indicated for p ≤ 0.05. ns, not significant. p.s., post-surgery; LH, left hind paw; RH, right hind paw. Motor recovery was assessed longitudinally using the CatWalk system from baseline to 7 weeks post-surgery. After SNI, animals shifted from a digitigrade to a plantigrade gait, reflected in increased print length (Fig. 5 A). Print width decreased due to the absence of outer toe prints, preventing calculation of toe spread (TS) and hence the sciatic functional index (SFI), a finding at odds with crush models where these measures are typically obtained (Fey et al., 2010 ; Vogelaar et al., 2004 ). Most paw-print parameters, including duty cycle, print area, intermediate toe spread, maximum intensity (at %), and maximum contact area (at %), improved between the 5th and 7th week but remained significantly impaired relative to baseline, indicating mid-phase motor recovery (Fig. 5 C). Paw-print parameters from a longitudinal CatWalk analysis performed from baseline to the 7th week post-surgery. (A) Representative paw-prints of the right hind limb before surgery and in the 5th and 7th week post-surgery. Print length (PL), toe spread (TS), and intermediate toe spread (ITS) are shown by white lines. The red arrow marks the tarsal (heel) pad, which is typically visible only in operated animals. (B) Schematic ventral view of the operated animal with the SNI located in the right hind limb. The mouse schematic was adjusted from BioRender. (C) Paw-print parameters of the affected right hind limb compared with the intact left hind limb. Individual data points represent the mean of three compliant runs per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 6 in the 5th week post-surgery (except ITS); n = 6 in the 7th week post-surgery (except ITS). Statistical significance and p-value are indicated for p ≤ 0.05. p.s., post-surgery; LH, left hind paw; RH, right hind paw. Dynamic gait analysis of all four limbs showed substantial deviations in phasing from baseline in the 5th week, with significant improvement in the 7th week (Fig. 6 A, B, D, F). Regularity indices of phase dispersions remained stable, suggesting coordinated, rather than chaotic, gait disruption (Fig. 6 C, E, G). Gait parameters from a longitudinal CatWalk analysis performed from baseline to the 7th week post-surgery. (A) Girdle phase dispersion LH-RH (theoretical baseline value 50%). (B) Schematic ventral view of the operated animal with the SNI located in the right hind limb and the four paws marked with different colours. The mouse schematic was adjusted from BioRender. (C) Regularity index of the girdle phase dispersion LH-RH (ideal value 1.00). (D) Diagonal phase dispersion LF-RH (theoretical baseline value 0%). (E) Regularity index of the diagonal phase dispersion LF-RH (ideal value 1.00). (F) Ipsilateral phase dispersion RF-RH (theoretical baseline value 50%). (G) Regularity index of the ipsilateral phase dispersion RF-RH (ideal value 1.00). Individual data points represent the mean of three compliant runs per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 5 in the 5th week post-surgery; n = 6 in the 7th week post-surgery. Statistical significance and p-value are indicated for p ≤ 0.05. p.s., post-surgery; LF, left front paw; RF, right front paw; LH, left hind paw; RH, right hind paw. Parameters unaffected by SNI included stride length, step sequence regularity, and base of support of the fore- and hind limbs (Suppl. Figure 1A-E). Supplemental Fig. 1 CatWalk test: Additional gait parameters Additional gait parameters are shown from the longitudinal CatWalk analysis performed from baseline to the 7th week post-surgery. (A) stride length. (B) Run speed. (C) Regularity index of the step sequence (ideal value 1.00). (D) Base of support (BoS) of the front paws. (E) BoS of the hind paws. Individual data points represent the mean of three compliant runs per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 5–6 in the 5th week post-surgery; n = 6 in the 7th week post-surgery. Statistical significance and p-value are indicated for p ≤ 0.05. p.s., post-surgery; LH, left hind paw; RH, right hind paw. Overall, 5–7 weeks post-lesion represents an appropriate time window to assess motor recovery, capturing ongoing regeneration before the plateau phase, and allowing room to detect improvements from potential regenerative interventions. DISCUSSION We introduce and characterize a mouse model of PNI that mimics severe lesions involving neurotmesis and incorporates clinically relevant microsurgical repair strategies. By combining nerve transection with conduit-based repair, this model bridges the gap between growing clinical practice and experimental research. Despite the increasing clinical use of synthetic nerve guidance conduits, experimental PNI research still relies on crush lesions or direct end-to-end coaptation after transection, both of which have limited clinical relevance. A systematic review of nerve transection studies identified 49 animal studies employing various coaptation techniques, but none using conduit-based repair (Vela et al., 2020 ). Similarly, a systematic review of experimental PNI models and motor recovery cited 223 studies, almost exclusively based on crush or direct ligation paradigms (Heinzel et al., 2020 ). In contrast, our nerve transection and conduit-based repair model closely reflects clinical practice and provides a framework for testing the therapeutic potential of bioactive compounds. While novel conduit designs, such as 3D-printed scaffolds, are actively being developed, these approaches remain exploratory and lack standardization and commercial availability (Kim et al., 2024 ; Larijani et al., 2024 ; Park et al., 2022 ; Wan et al., 2025 ; Wang et al., 2024 ). Chitosan nerve guidance conduits provide a practical solution to this need. They are commercially available, standardized, and exhibit excellent biocompatibility and biodegradability, which has led to their increasing use in clinical settings (Bocker et al., 2022 ; Neubrech et al., 2018 ). Chitosan interacts favorably with cells of the neural microenvironment and undergoes gradual degradation after implantation. In contrast to clinical application, we modified the conduit by halving it prior to implantation. This adjustment not only accommodates the smaller diameter of the mouse sciatic nerve but also enables the delivery of soluble, bioactive compounds directly to the lesion site via a hydrogel matrix. In our hands, PuraMatrix supported axonal regeneration more effectively than Matrigel (data not shown), making it a suitable substrate for this application. A technical challenge of this approach is the intrinsic autofluorescence of chitosan. Chitosan is a chitin derivative that was originally prepared from crab tendons (Yamaguchi et al., 2003 ) and has been developed for its application as a material for nerve guidance conduits (Boecker et al., 2019 ). As a polysaccharide, chitosan is not susceptible to lipid-based clearing protocols. We developed a protocol to quench conduit autofluorescence, allowing visualization of regenerating axons within the conduit using long-working-distance confocal microscopy (Fogli et al., 2019 ). However, this quenching effect is transient; autofluorescence gradually re-emerges over time, particularly in the 488 nm channel, limiting imaging at later time points. To improve signal-to-noise ratio, we developed a “peripheral perfusion” technique, in which perfusion via the vena cava and drainage through the femoral vein enhances blood removal from the hind limb compared to conventional transcardiac perfusion. Combined with transgenic fluorescent reporters and a shortened CUBIC clearing protocol (Susaki et al., 2014 ), this approach enables robust imaging of axonal and cellular dynamics while preserving GFP fluorescence. Functional recovery is a central outcome measure in experimental PNI models. While electrophysiological and electromyographic methods have traditionally been used (Navarro, 2016 ), computational gait analysis has become increasingly important, with the CatWalk system providing a precise and detailed quantitative assessment of locomotion (Chen et al., 2017 ; Heinzel et al., 2020 ). Based on known time courses of nerve regeneration, we selected a 4–7 week post-injury window to assess sensory and motor recovery. Previous studies have shown that reinnervation of neuromuscular junctions can occur as early as 2 weeks after injury in both crush and transection/coaptation models (Bauder & Ferguson, 2012 ; Vannucci et al., 2019 ), with sensory recovery preceding motor recovery (Navarro & Kennedy, 1991 ; Navarro et al., 1994 ). In general, axonal regeneration is expected to proceed more slowly after nerve transection and coaptation than after a crush lesion. A recent study in the rat directly compared these paradigms and found that functional recovery following transection and coaptation was delayed by approximately one week relative to a crush lesion. Specifically, sensory recovery began about 2 weeks after the crush lesion, whereas motor recovery did not commence before 3 weeks post-lesion (Wang et al., 2023 ). Consistent with these findings and taking into account an approximately one-week initiation phase required for axons to navigate the conduit microenvironment, we observed progressive functional improvement over a period of 4–7 weeks post-surgery. Within this window, sensory recovery was best assessed at 3–4 weeks, whereas motor recovery was most appropriately evaluated at 5–7 weeks after injury. Importantly, both time windows fall within the dynamic phase of regeneration, prior to the plateau phase, thereby providing sensitivity to detect therapeutic effects. A key difference between our model and conventional crush injury models lies in motor recovery patterns. Following transection and conduit repair, animals adopted a plantigrade gait and failed to use their outer toes even at later time points, precluding calculation of the sciatic functional index (SFI). In contrast, crush injury models typically exhibit rapid and near-complete recovery due to preservation of epineurium and endoneurial guidance structures, with functional restoration achieved within 3–4 weeks, including recovery of toe spread (Fey et al., 2010 ; Vogelaar et al., 2004 ). In the transection model, regenerating axons must bridge a physical gap and re-establish target connections without intact guidance structures, resulting in delayed motor recovery. Impaired intrinsic foot muscle function likely underlies the absence of toe spreading in our model. This highlights an important distinction between injury paradigms and suggests that foot muscle activity is particularly sensitive to severe nerve injuries. At the same time, restoration of toe spread is indicative of fine motor recovery and may therefore serve as a sensitive endpoint for evaluating therapeutic interventions. CONCLUSION The severity of traumatic peripheral nerve injury is often underestimated, partly due to the assumption that peripheral axons regenerate readily after injury. However, in cases of neurotmesis, successful regeneration depends on microsurgical repair and remains inefficient, frequently resulting in incomplete sensory and motor recovery and long-term deficits. This reflects the limited proportion of regenerating axons that successfully reach their targets and re-establish functional connections. Consequently, there is a pressing need for therapeutic strategies that enhance axonal regeneration and functional recovery. Progress in this field is hindered by the lack of experimental models that adequately reflect clinical conditions and allow for the effective delivery of bioactive compounds to the lesion site. To address these limitations, we developed a mouse model of PNI that closely mimics clinically relevant injury and repair paradigms. By combining sciatic nerve transection with tension-free, conduit-based repair, this lesion model also enables local delivery of bioactive agents to the lesion site, thereby providing a platform for preclinical testing of therapeutic interventions. All components of this model, including the transgenic reporter lines, microsurgical procedures, conduit preparation, tissue processing, and imaging workflows, are readily implementable (Fig. 7 ). We propose this paradigm as a standardized and clinically relevant reference model for experimental PNI research. Schematic overview of the experimental paradigm described in this study, including sciatic nerve transection, conduit implantation, tissue processing, imaging, and axon quantification. Declarations COMPETING INTERESTS The authors declare no competing interests. FUNDING The Vienna BioCenter Core Facilities (VBCF) Preclinical Phenotyping Facility acknowledges funding from the Austrian Federal Ministry of Education, Science & Research, and the City of Vienna. SM was supported by VIB, the VIB BioImaging Core Leuven, and the Research Foundation – Flanders (FWO) through grants FWOI000123N and FWOI001322N. Additionally, SM was supported by KU Leuven (KA/24/041). RS was supported by a research project grant from the Austrian Science Fund FWF (P33411-B). Author Contribution W.U., L.R., and F.S. refined and performed the nerve transection protocol, including conduit-based repair, and carried out tissue processing and clearing; S.M. imaged the nerve samples and processed images; S.M. and R.S. analysed the imaging data; S.B. carried out sensory-motor tests and analysed the data; W.U. and R.S. analysed sensory-motor data; E.D. contributed to the perfusion protocol and to macrophage imaging; S.M. and R.S. designed experiments and managed the project; R.S. conceived the study and wrote the manuscript. All authors reviewed the manuscript. Acknowledgement We thank Klaus Kraitsy for excellent assistance with aspects of the nerve transection experiments. We are grateful to Matilde Bongio (somersault18:24) for outstanding graphical rendering. The GFP-M mouse line was originally generated in the laboratories of Joshua R. Sanes and Jeff W. Lichtman. Data Availability All data are available from the authors upon request. 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U., Sun, C., Li, S., Zhang, N., Chen, H., Han, C. C., Xu, S., & Liu, Y. (2024, Apr). Perspectives on the Novel Multifunctional Nerve Guidance Conduits: From Specific Regenerative Procedures to Motor Function Rebuilding. Adv Mater , 36 (14), e2307805. https://doi.org/10.1002/adma.202307805 Additional Declarations No competing interests reported. Cite Share Download PDF Status: Under Review Version 1 posted Reviews received at journal 18 May, 2026 Reviews received at journal 06 May, 2026 Reviewers agreed at journal 30 Apr, 2026 Reviewers agreed at journal 26 Apr, 2026 Reviewers invited by journal 31 Mar, 2026 Editor assigned by journal 31 Mar, 2026 Submission checks completed at journal 31 Mar, 2026 First submitted to journal 28 Mar, 2026 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-9254882","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Article","associatedPublications":[],"authors":[{"id":616703812,"identity":"b2cff2ef-f58a-44ec-8781-b4b5f4ccb362","order_by":0,"name":"Walter Unterberger","email":"","orcid":"","institution":"Innsbruck Medical University","correspondingAuthor":false,"prefix":"","firstName":"Walter","middleName":"","lastName":"Unterberger","suffix":""},{"id":616703813,"identity":"ac52116d-a439-4fe3-a278-300cc4a0e617","order_by":1,"name":"Lucas Rubisoier","email":"","orcid":"","institution":"Innsbruck Medical University","correspondingAuthor":false,"prefix":"","firstName":"Lucas","middleName":"","lastName":"Rubisoier","suffix":""},{"id":616703814,"identity":"9d6e828e-f79c-4c96-99e9-de0a1f30e6e9","order_by":2,"name":"Felix Schöffing","email":"","orcid":"","institution":"Innsbruck Medical University","correspondingAuthor":false,"prefix":"","firstName":"Felix","middleName":"","lastName":"Schöffing","suffix":""},{"id":616703815,"identity":"cc0307c9-9d61-4e93-844c-c7f945fea5e6","order_by":3,"name":"Egon Demetz","email":"","orcid":"","institution":"Innsbruck Medical University","correspondingAuthor":false,"prefix":"","firstName":"Egon","middleName":"","lastName":"Demetz","suffix":""},{"id":616703816,"identity":"3f98418b-9c0a-4d9e-b4b2-d6dd007d5bd9","order_by":4,"name":"Sylvia Badurek","email":"","orcid":"","institution":"Vienna BioCenter Core Facilities (VBCF)","correspondingAuthor":false,"prefix":"","firstName":"Sylvia","middleName":"","lastName":"Badurek","suffix":""},{"id":616703817,"identity":"fae351de-dec6-437b-872b-c50893007303","order_by":5,"name":"Sebastian Munck","email":"","orcid":"","institution":"VIB-KU Leuven Center for Neurosciences","correspondingAuthor":false,"prefix":"","firstName":"Sebastian","middleName":"","lastName":"Munck","suffix":""},{"id":616703818,"identity":"754a0808-a855-4253-bd8f-9fc75ee951b2","order_by":6,"name":"Rüdiger Schweigreiter","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAABT0lEQVRIie2RMWuDQBTHn4tZDtzKK4bkK1gCSQpBv4py4JSStUMoHkKyhMwpFPoxunS4INhF2tXiohQyp0sIQdI+aSExduhYqD/wvON/P5/3DqCm5o+C9tdbLeb0KB4NAwbfy18pghz3SFGrCkA5KcoER1FZ6S34Mk53gxFoweqNXffNh6k/Td8fX5pwFixjGJug3XvHSjN2ed+Zu5ceur0Oi5DfRUshFquEgU4RhBwwLP8UDrvozAIDkKn61QQ5oiN8JhNmJRSBKgFOldGGlA8DtIiUPSntTPi5fKYqFMFeQrtSRUV7Kw2AISkemoiK8EFKUihSJhKME4WtOn3H44aKblfPQ7SROeJ2JnlxFormnF2Edklp8Ox1m5uGRh07X4xvLGw8peutNC3QeRavN2arFcjKxSiTQ++dUkOBvs8q+wvyw9T6cUNNTU3Nv+YTgeZvnkQ4f1gAAAAASUVORK5CYII=","orcid":"","institution":"Innsbruck Medical University","correspondingAuthor":true,"prefix":"","firstName":"Rüdiger","middleName":"","lastName":"Schweigreiter","suffix":""}],"badges":[],"createdAt":"2026-03-28 19:23:03","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-9254882/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-9254882/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":106257347,"identity":"98eb0ae1-9aa9-4acc-8d2a-8af414ee5934","added_by":"auto","created_at":"2026-04-06 19:26:14","extension":"jpg","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":409829,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eMicrosurgery and clearing in the conduit-based nerve transection lesion model\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003e Sciatic nerve immediately after transection. Retraction of the proximal and distal nerve stumps is visible. \u003cstrong\u003e(B)\u003c/strong\u003e Placement of the chitosan conduit beneath the transected nerve. Bulbous nerve ends are formed after transection. The proximal and distal nerve stumps are fixed to the conduit using epineural sutures without tension, leaving a gap of approximately 0.5–1 mm between the nerve ends. \u003cstrong\u003e(C)\u003c/strong\u003eThe conduit is wrapped and closed at the ends. Before closing the central portion, a hydrogel matrix is pipetted into the conduit; regeneration-promoting agents can be added to this matrix. \u003cstrong\u003e(D)\u003c/strong\u003e The conduit is fully closed, and the surgical site is subsequently closed with muscle and skin sutures. \u003cstrong\u003e(E)\u003c/strong\u003eTwo weeks after surgery, animals were euthanized, and the nerve-conduit assembly was dissected following peripheral perfusion and fixation. \u003cstrong\u003e(F)\u003c/strong\u003eThe nerve-conduit assembly after the CUBIC-based clearing protocol. Scale bar, 500 µm.\u003c/p\u003e","description":"","filename":"Picture1.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/7b4e87fa7170d354bb7b3a61.jpg"},{"id":106257342,"identity":"4379cd53-6c05-4f3a-a4da-dd668888a608","added_by":"auto","created_at":"2026-04-06 19:26:13","extension":"jpg","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":505318,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eImaging and quantification of axons within the chitosan conduit\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A) \u003c/strong\u003eCombined transmission light and EGFP fluorescence confocal imaging of the nerve-conduit assembly two weeks after PNI using a 10x long-working distance objective. A GFP-M mouse, which expresses GFP in a mosaic pattern in sensory and motor fibers (approximately 1% of all axons; (Feng et al., 2000)), was used. Dashed white lines indicate the proximal and distal nerve stumps within the conduit. \u003cstrong\u003e(B)\u003c/strong\u003e GFP fluorescence channel alone. The yellow arrow indicates the typical drop in fluorescence at the conduit edge. \u003cstrong\u003e(C)\u003c/strong\u003e Maximum intensity projection (MIP) of the grayscale image stack after pixel classification and subtraction of conduit autofluorescence. \u003cstrong\u003e(D)\u003c/strong\u003e MIP of the grayscale image stack after pixel classification and thresholding. Regenerating nerve fibers in the distal nerve stump are indicated by yellow arrowheads. Orthogonal slices of the proximal and distal stumps are shown at positions marked by blue lines. \u003cstrong\u003e(E)\u003c/strong\u003eQuantification of GFP-positive intersections per orthogonal slice, corresponding to axon number, along the nerve-conduit axis. The yellow arrow indicates the typical drop in fluorescence at the conduit edge. The light-blue curve presents data at each incremental position. The dark-blue curve has been smoothed over 50 µm intervals. Scale bar, 500 µm.\u003c/p\u003e","description":"","filename":"Picture2.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/1d899e5c5056c9f62e053fc4.jpg"},{"id":106257343,"identity":"e60f712a-a036-4d1b-b9c1-58c9d08ed0ee","added_by":"auto","created_at":"2026-04-06 19:26:13","extension":"jpg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":328850,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eTime course of axonal and macrophage dynamics after PNI\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eA double-reporter mouse line generated by crossing GFP-M mice with the macrophage-specific tamoxifen-inducible tdTomato reporter line Cx3cr1\u003csup\u003eCreERT2\u003c/sup\u003e::ROSA26\u003csup\u003etdTomatofl/fl\u003c/sup\u003e was used to visualize axon and macrophage dynamics for up to six weeks after SNI. MIPs are shown for each time point. \u003cstrong\u003e(A) \u003c/strong\u003eProgressive axonal fragmentation in the distal nerve stump, indicative of Wallerian degeneration, is visible at 4 days and 1 week after surgery (white arrow heads). By 2 weeks after SNI, all transected axons have been degraded, and the first regenerative fibers extend into the distal nerve stump (white arrows). \u003cstrong\u003e(B)\u003c/strong\u003e Macrophages are present in the intact nerve and increase in number several days after SNI due to recruitment. By 6 weeks after SNI, macrophages are largely cleared from the lesion site. \u003cstrong\u003e(C)\u003c/strong\u003e Combined axonal and macrophage signals. Scale bar, 500 µm. p.s., post-surgery.\u003c/p\u003e","description":"","filename":"Picture3.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/cc1964eb763ae1fc510c9a37.jpg"},{"id":106402633,"identity":"ea6c82f9-094c-4043-8d4c-a1254df60cdd","added_by":"auto","created_at":"2026-04-08 09:12:28","extension":"jpg","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":71801,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003evon Frey test\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eLongitudinal von Frey testing was performed from baseline to the 6\u003csup\u003eth\u003c/sup\u003e week post-surgery.\u003cstrong\u003e (A) \u003c/strong\u003eForce required to elicit hind paw withdrawal. \u003cstrong\u003e(B)\u003c/strong\u003e Latency to hind paw withdrawal. Individual data points represent the mean of three measurements per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 6 in the 4\u003csup\u003eth\u003c/sup\u003e week post-surgery; n = 6 in the 6\u003csup\u003eth\u003c/sup\u003e week post-surgery. Statistical significance and p-value are indicated for p ≤ 0.05. ns, not significant. p.s., post-surgery; LH, left hind paw; RH, right hind paw.\u003c/p\u003e","description":"","filename":"Picture4.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/eeeb8f6aed1efe9f1359e3e8.jpg"},{"id":106257344,"identity":"38573c76-caec-46bc-b17d-c2013b9d8a1d","added_by":"auto","created_at":"2026-04-06 19:26:13","extension":"jpg","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":233924,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eCatWalk test: Paw-print parameters\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003ePaw-print parameters from a longitudinal CatWalk analysis performed from baseline to the 7\u003csup\u003eth\u003c/sup\u003e week post-surgery.\u003cstrong\u003e (A) \u003c/strong\u003eRepresentative paw-prints of the right hind limb before surgery and in the 5\u003csup\u003eth\u003c/sup\u003e and 7\u003csup\u003eth\u003c/sup\u003e week post-surgery. Print length (PL), toe spread (TS), and intermediate toe spread (ITS) are shown by white lines. The red arrow marks the tarsal (heel) pad, which is typically visible only in operated animals. \u003cstrong\u003e(B)\u003c/strong\u003e Schematic ventral view of the operated animal with the SNI located in the right hind limb. The mouse schematic was adjusted from BioRender. \u003cstrong\u003e(C)\u003c/strong\u003e Paw-print parameters of the affected right hind limb compared with the intact left hind limb. Individual data points represent the mean of three compliant runs per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 6 in the 5\u003csup\u003eth\u003c/sup\u003e week post-surgery (except ITS); n = 6 in the 7\u003csup\u003eth\u003c/sup\u003e week post-surgery (except ITS). Statistical significance and p-value are indicated for p ≤ 0.05. p.s., post-surgery; LH, left hind paw; RH, right hind paw.\u003c/p\u003e","description":"","filename":"Picture5.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/2c9774440e3ed8c56dd1350e.jpg"},{"id":106402859,"identity":"d41494e0-8430-48b9-9317-50705c5a3822","added_by":"auto","created_at":"2026-04-08 09:13:03","extension":"jpg","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":171736,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eCatWalk test: Gait parameters\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eGait parameters from a longitudinal CatWalk analysis performed from baseline to the 7\u003csup\u003eth\u003c/sup\u003e week post-surgery. \u003cstrong\u003e(A)\u003c/strong\u003e Girdle phase dispersion LH-RH (theoretical baseline value 50%). \u003cstrong\u003e(B)\u003c/strong\u003e Schematic ventral view of the operated animal with the SNI located in the right hind limb and the four paws marked with different colours. The mouse schematic was adjusted from BioRender. \u003cstrong\u003e(C)\u003c/strong\u003e Regularity index of the girdle phase dispersion LH-RH (ideal value 1.00). \u003cstrong\u003e(D)\u003c/strong\u003e Diagonal phase dispersion LF-RH (theoretical baseline value 0%). \u003cstrong\u003e(E)\u003c/strong\u003e Regularity index of the diagonal phase dispersion LF-RH (ideal value 1.00). \u003cstrong\u003e(F)\u003c/strong\u003e Ipsilateral phase dispersion RF-RH (theoretical baseline value 50%). \u003cstrong\u003e(G)\u003c/strong\u003e Regularity index of the ipsilateral phase dispersion RF-RH (ideal value 1.00). Individual data points represent the mean of three compliant runs per animal. Data are presented as mean ± SD; n = 12 before surgery; n = 5 in the 5\u003csup\u003eth\u003c/sup\u003e week post-surgery; n = 6 in the 7\u003csup\u003eth\u003c/sup\u003e week post-surgery. Statistical significance and p-value are indicated for p ≤ 0.05. p.s., post-surgery; LF, left front paw; RF, right front paw; LH, left hind paw; RH, right hind paw.\u0026nbsp;\u0026nbsp;\u0026nbsp;\u0026nbsp;\u0026nbsp;\u0026nbsp;\u003c/p\u003e","description":"","filename":"Picture6.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/107ccf664ec026af60e3826d.jpg"},{"id":106257346,"identity":"c5ddda99-4750-4026-9f8e-7fcc9cca2891","added_by":"auto","created_at":"2026-04-06 19:26:13","extension":"jpg","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":232573,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSchematic overview of the conduit-based sciatic nerve transection lesion paradigm in the mouse\u003c/strong\u003e\u003c/p\u003e","description":"","filename":"Picture7.jpg","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/90be286fa6c15a65583abd27.jpg"},{"id":106405794,"identity":"b695d325-17fe-4660-b3b0-6f2355b0a190","added_by":"auto","created_at":"2026-04-08 09:28:33","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":3014684,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-9254882/v1/1eab6a2b-26a2-4663-8ce0-5c109b65dcad.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"A clinically relevant mouse model for traumatic peripheral nerve injury and repair","fulltext":[{"header":"INTRODUCTION","content":"\u003cp\u003eUnlike axons in the central nervous system (CNS), peripheral nerves are often assumed to regenerate spontaneously without the need for extensive medical intervention. While axonal regrowth does occur following Wallerian degeneration in the peripheral nervous system (PNS), this regenerative capacity is largely restricted to mild injuries such as compression or crush lesions. In contrast, traumatic nerve injuries involving complete transection do not regenerate spontaneously (Phillips et al., \u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). These severe injuries require surgical repair, and even then, functional outcomes are often poor, frequently resulting in lifelong sensory and motor deficits (Bergmeister et al., \u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e2020\u003c/span\u003e; Ciaramitaro et al., \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e2010\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eA major limitation to recovery is the failure of long-distance axonal regeneration. Most severed axons either fail to regenerate or abort growth prematurely, often accompanied by axonal sprouting and neuroma formation (Allodi et al., \u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e2012\u003c/span\u003e; Zhao et al., \u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e2023\u003c/span\u003e). Traumatic peripheral nerve injury (PNI) most commonly affects the upper extremities and is frequently caused by occupational or traffic accidents (Bergmeister et al., \u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e2020\u003c/span\u003e; Noble et al., \u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e1998\u003c/span\u003e). In addition, warfare-related injuries are increasing in prevalence and often involve complex, multi-site nerve damage affecting upper and lower extremities simultaneously (Eckhoff et al., \u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e2021\u003c/span\u003e; Howard et al., \u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e2025\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eThe current gold standard for tension-free nerve repair is autologous nerve grafting, typically using a segment of the sural nerve to bridge the lesion gap (Safa \u0026amp; Buncke, \u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e2016\u003c/span\u003e). While this approach supports axonal regrowth, it is limited by donor-site morbidity and graft availability. Synthetic nerve guidance conduits have emerged as a promising alternative, offering standardized quality, unrestricted availability, and favorable biocompatibility (Redolfi Riva et al., \u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e2024\u003c/span\u003e; Zhou et al., \u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e2024\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eDespite these clinical advances, experimental models of PNI remain poorly aligned with clinical practice. Most studies rely on crush lesions or nerve transection followed by direct coaptation, neither of which adequately reflects current surgical approaches (Heinzel et al., \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e2020\u003c/span\u003e; Vela et al., \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). Crush injuries are typically not treated surgically, and direct end-to-end repair is often avoided clinically due to tension and suboptimal outcomes compared to conduit-based repair (Berger \u0026amp; Millesi, \u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e1978\u003c/span\u003e; Leis et al., \u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e2024\u003c/span\u003e; M et al., 2014). Moreover, these models do not permit controlled local delivery of bioactive compounds to evaluate their therapeutic potential. In addition, crush lesions may confound interpretation, as a subset of axons can survive the injury and avoid Wallerian degeneration (Kim et al., \u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e2023\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eTo address these limitations, we developed a mouse model of traumatic PNI that more closely replicates clinically relevant injury and repair paradigms. The model combines sciatic nerve transection with conduit-based repair using a longitudinally opened chitosan nerve guidance conduit. This procedure involves both surgical intervention and reconstruction. Although microsurgical procedures are technically more demanding in mice than in rats due to smaller anatomic structures, the mouse offers access to a wide range of transgenic reporter lines (Cai et al., \u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e2013\u003c/span\u003e; Feng et al., \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e2000\u003c/span\u003e). Chitosan conduits are commercially available, standardized, and increasingly used in clinical settings due to their favorable biocompatibility (Bocker et al., \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e2022\u003c/span\u003e; Meyer et al., \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e2016\u003c/span\u003e; Neubrech et al., \u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e2016\u003c/span\u003e; Neubrech et al., \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e). This approach enables direct application of soluble bioactive compounds to the lesion site and their evaluation in a controlled microenvironment. Using an adapted imaging strategy for synthetic conduits, we are able to visualize and quantify regenerating axons at single-fiber resolution within the conduit and distal stump (Fogli et al., \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e2019\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eHere, we present this model in detail, describe microsurgical techniques and conduit processing, characterize axonal regeneration and macrophage dynamics over a time course from 2 days to 6 weeks post-injury, and assess functional recovery using sensory and motor assays.\u003c/p\u003e"},{"header":"MATERIALS AND METHODS","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eAnimals\u003c/h2\u003e \u003cp\u003eAdult (2\u0026ndash;6 month-old) female GFP-M mice (Feng et al., \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e2000\u003c/span\u003e) or Cx3cr1\u003csup\u003eCreERT2\u003c/sup\u003e::ROSA26\u003csup\u003etdTomatofl/fl\u003c/sup\u003e mice (Petzer et al., \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e2018\u003c/span\u003e) were used for sciatic nerve surgery and imaging experiments. Adult (3 month-old) female C57BL/6N mice from Charles River Laboratories were used for sciatic nerve surgery and sensory-motor experiments. Animals were housed in individually ventilated cages (IVCs) in a temperature- and humidity-controlled room under a 12-hour light/12-hour dark cycle with access to food and water \u003cem\u003ead libitum\u003c/em\u003e. For time course experiments, homozygous GFP-M mice were crossbred with homozygous Cx3cr1\u003csup\u003eCreERT2\u003c/sup\u003e::ROSA26\u003csup\u003etdTomatofl/fl\u003c/sup\u003e mice to obtain a heterozygous double GFP/Tomato reporter line. For imaging experiments involving the Cx3cr1\u003csup\u003eCreERT2\u003c/sup\u003e::ROSA26\u003csup\u003etdTomatofl/fl\u003c/sup\u003e mouse line, normal food was exchanged with pellets containing tamoxifen (360 mg Tamoxifen citrate/kg; sniff #A11570360) one week before nerve-conduit dissection. All experimental protocols were approved by the Austrian Federal Ministry of Science and Research (GZ #2021\u0026thinsp;\u0026minus;\u0026thinsp;0.406.859) and complied with the European Convention for the Protection of Vertebrate Animals Used for Experimental and other Scientific Purposes (ETS no. 123).\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eSciatic nerve transection and conduit surgery\u003c/h3\u003e\n\u003cp\u003eSterility was maintained throughout all surgical procedures. To provide a guidance structure for axonal regeneration, a Reaxon\u0026reg; chitosan conduit (2.1 mm \u0026times; 14 mm) was used (Medovent #RD121). Prior to NaBH₄ treatment, the conduit was softened in distilled water and cut into six equal pieces by first dividing it transversally into three 4-mm tubes and subsequently splitting each tube longitudinally. The resulting conduit segments measured approximately 4 \u0026times; 3 mm. If not used immediately, conduits were dried for 24 h in a laminar flow hood and stored in sealed Petri dishes.\u003c/p\u003e \u003cp\u003eTo reduce autofluorescence, conduits were incubated in 10 mg/mL NaBH₄ (Merck #213462) in 1\u0026times; PBS for at least 40 h. Conduits were then washed five times by inversion in 40 mL sterile 1\u0026times; PBS in a 50 mL tube under sterile conditions. After washing, conduits were stored in sterile 1\u0026times; PBS in sealed Petri dishes.\u003c/p\u003e \u003cp\u003eFor pain management, a dual analgesic regimen was applied. Beginning one day before surgery, mice received 1.25 mg/mL metamizole (Metagelan\u0026reg;; G.L. Pharma #137760) in the drinking water. To improve palatability, the solution was supplemented with apple juice. Treatment was continued until postoperative day 2 and then replaced with autoclaved tap water. In addition, mice received daily subcutaneous injections of carprofen (0.5 mg/mL in sterile 1\u0026times; PBS; Rimadyl\u0026reg;; Zoetis #10000319) at 10 \u0026micro;L per g body weight from the day of surgery until postoperative day 3.\u003c/p\u003e \u003cp\u003eAnaesthesia was induced and maintained with isoflurane (IsoFlo\u0026reg;; Zoetis #400136.00.00) delivered via a mask at a flow rate of 0.3 L/min. Isoflurane concentration was maintained at 2.5% until nerve transection and subsequently reduced to 2.0% to limit overall anaesthetic exposure. Body temperature was continuously monitored using a rectal probe and maintained with a heating plate set to 37\u0026deg;C. After induction of anaesthesia and analgesic administration, the right hind limb was shaved, and the surgical area was disinfected with an iodine-containing antiseptic.\u003c/p\u003e \u003cp\u003eA neurosurgical microscope (S100/OPMI pico; Carl Zeiss Meditec) and specialized microsurgical instruments were used for all surgical procedures. The right hind limb was extended and fixed to the surgical table to facilitate visualization of the femur. A longitudinal 4\u0026ndash;5 mm skin incision was made just below the caudal border of the femur, and the subcutaneous tissue was separated using scissors. The underlying muscles were bluntly separated at the natural connective tissue plane to expose the sciatic nerve. Connective tissue surrounding the nerve was carefully removed, and the nerve was transected with spring scissors in a single cut.\u003c/p\u003e \u003cp\u003eFollowing transection, the distal stump was sutured to the inner wall of the conduit using an Ethilon\u0026reg; 9\u0026thinsp;\u0026minus;\u0026thinsp;0 (Johnson \u0026amp; Johnson #EH7448G) suture. The needle was inserted paramedially from the outside of the conduit to the inside (approximately 1.5 mm from the edge), passed through the epineurium and back through the conduit wall, where a knot was tied (with three throws). This creates a loop that safely secures the nerve stump to the conduit. The same procedure was performed for the proximal stump, leaving a gap between the stumps of up to 1 mm. To close the conduit after fixation of the nerve stumps, the conduit was rotated within the surgical field to expose its corners. The distal end was first closed by connecting the conduit corners with a simple interrupted suture. A second distal suture was placed to enable reliable identification of the sample orientation after extraction of the nerve-conduit assembly. The proximal corners were then sutured in the same manner. This configuration left a small opening between the sutures that allowed matrix instillation. To minimize matrix loss during final closure, the closing suture was pre-positioned before matrix instillation. The viscous matrix, which was prepared freshly on the day of surgery, was instilled using a slightly cut-off tip to increase the opening diameter. The matrix composition was: 50 mM HEPES, 1x PBS, 0.85x PuraMatrix\u0026trade; (Corning #CLS354250). The conduit was then fully closed with a simple interrupted suture. Finally, the nerve\u0026ndash;conduit assembly was rotated back into its original position, and the muscles were closed using a mattress suture with Permahand\u0026reg; 5\u0026thinsp;\u0026minus;\u0026thinsp;0 (Johnson \u0026amp; Johnson #K890H) sutures to minimize tension. The skin was closed with either interrupted buried sutures or surgical clips.\u003c/p\u003e\n\u003ch3\u003eTissue preparation and clearing\u003c/h3\u003e\n\u003cp\u003eTwo weeks after surgery, mice were euthanized and peripherally perfused with 1\u0026times; PBS and 4% PFA via an intravenous cannula inserted into the vena cava. Prior to perfusion, mice were deeply anaesthetized by intraperitoneal injection of approximately 100 \u0026micro;L of a 2:1 ketamine/xylacine mixture (100 mg/ml Ketasol\u0026reg;, aniMedica #8-00173; 20 mg/ml Xylasol\u0026reg;, aniMedica #8-00178). After insertion of the cannula, the right femoral vein (vena femoralis dextra) was punctured to allow blood drainage and facilitate removal of intravascular hemoglobin. Following perfusion, the skin above the operated area was removed, and the tissue was fixed in 4% paraformaldehyde at 4\u0026deg;C for 24 h. The nerve-conduit assembly was then carefully excised while preserving as much surrounding nerve tissue as possible. Samples were washed twice in 50 mL 1\u0026times; PBS for 2 h on a tube roller wrapped in aluminum foil.\u003c/p\u003e \u003cp\u003eTissue clearing was performed using a modified CUBIC protocol (Susaki et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2014\u003c/span\u003e), in which reagent ratios were maintained while incubation times were adjusted. Briefly, CUBIC-1 was freshly prepared by dissolving urea (25 wt%), Quadrol\u0026reg; (25 wt%), and Triton\u0026trade; X-100 (15 wt%) in distilled water (35 wt%) by stirring at 40\u0026ndash;50\u0026deg;C, followed by degassing. CUBIC-2 was prepared similarly by dissolving sucrose (50 wt%), urea (25 wt%), triethanolamine (10 wt%), and Triton\u0026trade; X-100 (0.1 vol%) in distilled water (15 wt%). All reagents were obtained from Merck. To minimize signal loss, samples were protected from light by wrapping tubes in aluminum foil. Samples were incubated in 10 mL CUBIC-1 on an incubation shaker at 55 rpm and 37\u0026deg;C for 2 days, with the reagent refreshed after the first day. Samples were subsequently washed three times for 2 h in 40 mL 1\u0026times; PBS on a tube roller. During the washing steps, 50 ml flasks and continuous rotation were required to prevent precipitation within the nerve-conduit assembly. After washing, samples were transferred to 10 mL CUBIC-2 and incubated for 2 days while shaking at 55 rpm and 37\u0026deg;C. On day 5, samples were transferred to 2 mL microcentrifuge tubes filled with CUBIC-2 and shipped overnight for imaging (Schweigreiter et al., \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e2020\u003c/span\u003e), resulting in a total CUBIC-2 incubation time of approximately 2.5 days.\u003c/p\u003e\n\u003ch3\u003eConfocal microscopy\u003c/h3\u003e\n\u003cp\u003eNerve\u0026ndash;conduit assemblies were placed in sealed custom-made imaging chambers filled with CUBIC-2 solution. The chambers were based on standard microscope slide dimensions and contained a hollow cavity (15 mm diameter, 2 mm depth). Standard coverslips sealed with desiccator grease were used to close the cavity. Up to four nerves were mounted in parallel using a holder compatible with the Caco-2 standard well-plate insert of the motorized microscope stage.\u003c/p\u003e \u003cp\u003eImaging was performed using a Nikon A1R confocal microscope mounted on a Nikon Eclipse Ni-E microscope body. A Plan Apo 10\u0026times; objective (NA 0.45, working distance 4.0 mm) was used to capture the full width of the nerve. Green and red fluorescence were excited using 488 nm and 561 nm lasers, respectively, with emission detected through 515/30 nm and 593/46 nm filters. To image the entire nerve\u0026ndash;conduit assembly, tiles centered on the middle of the conduit were acquired with 15% overlap. Individual tiles were acquired at 1024 \u0026times; 1024 pixels with a pixel size of 1.24 \u0026micro;m. Z-stacks of up to 800 \u0026micro;m were acquired per nerve with a step size of 2.45 \u0026micro;m. Multiple stage positions were defined for each sample holder to enable consecutive imaging of up to four nerves in a single session.\u003c/p\u003e\n\u003ch3\u003eImage processing\u003c/h3\u003e\n\u003cp\u003eBecause the chitosan conduit retained residual autofluorescence despite quenching, a multistep image analysis workflow was implemented. First, image stacks were aligned and cropped to a standardized field of view using Fiji / ImageJ (v1.54p) (Schindelin et al., \u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e2012\u003c/span\u003e). For segmentation, pixel classification was performed using ilastik (v1.4.0) (Berg et al., \u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e2019\u003c/span\u003e; Kreshuk et al., \u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e2011\u003c/span\u003e). Two classes were defined to distinguish axonal structures from everything else, including the conduit material, debris, and background. The resulting probability maps were exported to Fiji and thresholded to obtain a more stringent 3D binary mask (confidence\u0026thinsp;\u0026gt;\u0026thinsp;60\u0026ndash;95%), where background was 0 and detected axons were 1. Subsequently, the mask was multiplied with the original grayscale image stack to preserve the original fluorescence intensities while removing background and conduit autofluorescence.\u003c/p\u003e \u003cp\u003eThe processed image stacks were subsequently resliced to generate \u003cem\u003een face\u003c/em\u003e views of the nerve. Nerve fibers were detected in each resliced section using a custom GA3 spot-detection workflow implemented in NIS-Elements AR (v6.20.01), and the resulting counts were exported to Microsoft Excel. This approach generated a progressive, micrometer-resolved count of nerve fibers per pixel row along the length of the nerve-conduit assembly.\u003c/p\u003e \u003cdiv id=\"Sec8\" class=\"Section2\"\u003e \u003ch2\u003evon Frey test\u003c/h2\u003e \u003cp\u003eMechanical sensitivity was assessed using a dynamic plantar aesthesiometer (Ugo Basile; model 37450) equipped with the touch stimulator (cat. no. 37400-002) and base platform (model 37000-003). On each testing day, the device was calibrated according to the manufacturer\u0026rsquo;s instructions using 5 g and 50 g calibration weights. Mice were first allowed to habituate to the testing room for at least 30 min in their home cages. Animals were then transferred to the testing chambers and allowed to habituate for approximately 2 h prior to testing. Each hind paw was stimulated three times with a gradually increasing force ranging from 0 to 10 g over a 20 s ramp period. Consecutive stimulations were separated by at least 120 s. For each hind paw, the mean of the three measurements was calculated. Outcome parameters were the force and latency at which the animal withdrew the hind paw. Baseline measurements were performed approximately one week before surgery. Postoperative measurements were conducted in the 4th and 6th week after surgery.\u003c/p\u003e \u003c/div\u003e\n\u003ch3\u003eCatWalk test\u003c/h3\u003e\n\u003cp\u003eGait and paw-print parameters were analyzed using the CatWalk XT system (v10.6; Noldus Information Technology BV), largely following the protocol described in (Moritz et al., \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e2019\u003c/span\u003e) with minor modifications. After a 2-week habituation period to the facility, experiments were initiated. Mice were not trained to reach a goal box. Instead, after each run, they were returned to the beginning of the walkway or, after completing three compliant runs, put back to their home cage. Experiments were conducted in the dark. To improve paw-print contrast, mice were briefly placed in a cage containing a wet paper towel (tap water) immediately before testing to moisten the paws. The walkway floor and walls were cleaned with household cleaner between animals, or whenever urine or fecal boli were deposited on the walkway.\u003c/p\u003e \u003cp\u003eFor baseline measurements, mice were trained in the CatWalk system at 10 weeks of age on two consecutive days and tested two days later to generate the \u0026ldquo;before surgery\u0026rdquo; dataset. The following acquisition settings were used during training and testing. The camera was positioned 27 cm below the walkway. Minimum and maximum run durations were set to 0.5 s and 5 s, respectively. The minimum number of compliant runs per animal was set to three, and the maximum allowed speed variation was set to 60%. Camera gain was set to 12.9 dB, the green intensity threshold to 0.1, the red ceiling illumination to 17.7 V, and the green walkway illumination to 16.5 V. Postoperative measurements were performed in the 5th and 7th week after surgery to generate the corresponding post-surgery datasets. For paw-print analysis, runs were included with a minimum number of consecutive steps of 5, average speed from 10.0 to 40.0 cm/second, and maximum allowed speed variation of 60%. For gait cycle analysis, runs were included with a minimum number of consecutive steps of 10, average speed from 10.0 to 40.0 cm/second, and maximum allowed speed variation of 50%. Paw-prints were controlled manually to ensure correct identification of the affected right hind paw.\u003c/p\u003e \u003cdiv id=\"Sec10\" class=\"Section2\"\u003e \u003ch2\u003eStatistical analysis\u003c/h2\u003e \u003cp\u003eStatistical analyses were performed using GraphPad Prism (v10.6.1; GraphPad Software). Data were assessed for normality using the Kolmogorov\u0026ndash;Smirnov test, and no significant deviations from normal distribution were detected. For paw-print parameters obtained from CatWalk analyses, two-way analysis of variance (ANOVA) was used to evaluate the effects of experimental group (left versus right hind paw) and time point. When missing values resulted in unequal sample sizes, a mixed-effects model was applied instead of two-way ANOVA. Gait parameters were analyzed using one-way ANOVA. When significant effects were detected, pairwise comparisons were performed using Bonferroni\u0026rsquo;s multiple-comparisons post hoc test.\u003c/p\u003e \u003cp\u003eA significance threshold of \u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026lt;\u0026thinsp;0.05 was applied. Exact sample sizes and data presentation (mean\u0026thinsp;\u0026plusmn;\u0026thinsp;SD) are indicated in the respective figure legends. No statistical methods were used to predetermine sample size.\u003c/p\u003e \u003c/div\u003e"},{"header":"RESULTS","content":"\u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003eChitosan conduit-based repair of the transected sciatic nerve and tissue clearing\u003c/h2\u003e \u003cp\u003eThe sciatic nerve was transected in the upper hind limb proximal to its trifurcation into the peroneal, tibial, and sural branches. Immediately after transection, the stumps retracted due to intrinsic axial tension. Commercial chitosan nerve guidance conduits are designed for clinical use, and even the smallest available diameter (2.1 mm) exceeds that of the mouse sciatic nerve. To adapt the conduit for mice, we cut it in half and positioned one half beneath the nerve stumps. Before placement, the conduit was soaked in NaBH₄ to quench autofluorescence. Each stump was fixed to the conduit with tensionless epineural sutures, leaving a 0.5\u0026ndash;1 mm gap between the stumps. The conduit was subsequently closed at the ends and the center, and the cavity filled with PuraMatrix hydrogel, which can be supplemented with biologically active compounds to promote axonal regeneration and functional recovery (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA-D). Following wound closure, animals were allowed to recover for a period appropriate to the readout: 2 weeks for imaging axonal regeneration, when the majority of regenerating axons had crossed the gap and pioneering fibers had begun extending into the distal stump, and 4\u0026ndash;7 weeks for functional assessments of sensory and motor recovery.\u003c/p\u003e \u003cp\u003e For imaging, animals were perfused with PBS followed by PFA. Perfusion was necessary to flush blood from the lesion site, particularly in cases of surgery-induced vasculogenesis. Because conventional cardiac perfusion does not efficiently clear blood from the upper hind limb, we implemented a \u0026ldquo;peripheral perfusion\u0026rdquo; protocol, with perfusion through the vena cava and drainage via the femoral vein. The nerve-conduit assembly was then dissected, immersion-fixed for 24 hours, and washed thoroughly with PBS. At this stage, samples could be stored at 4\u0026deg;C in the dark to preserve GFP fluorescence. Tissue clearing was performed using an adapted CUBIC protocol (Susaki et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2014\u003c/span\u003e), with 2 days in CUBIC 1 followed by 2.5 days in CUBIC 2 (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eE-F). Extended incubation in CUBIC 2 was found to diminish GFP fluorescence; imaging was therefore performed without delay.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cb\u003e(A)\u003c/b\u003e Sciatic nerve immediately after transection. Retraction of the proximal and distal nerve stumps is visible. \u003cb\u003e(B)\u003c/b\u003e Placement of the chitosan conduit beneath the transected nerve. Bulbous nerve ends are formed after transection. The proximal and distal nerve stumps are fixed to the conduit using epineural sutures without tension, leaving a gap of approximately 0.5\u0026ndash;1 mm between the nerve ends. \u003cb\u003e(C)\u003c/b\u003e The conduit is wrapped and closed at the ends. Before closing the central portion, a hydrogel matrix is pipetted into the conduit; regeneration-promoting agents can be added to this matrix. \u003cb\u003e(D)\u003c/b\u003e The conduit is fully closed, and the surgical site is subsequently closed with muscle and skin sutures. \u003cb\u003e(E)\u003c/b\u003e Two weeks after surgery, animals were euthanized, and the nerve-conduit assembly was dissected following peripheral perfusion and fixation. \u003cb\u003e(F)\u003c/b\u003e The nerve-conduit assembly after the CUBIC-based clearing protocol. Scale bar, 500 \u0026micro;m.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec13\" class=\"Section2\"\u003e \u003ch2\u003eImaging and quantifying regenerative axonal growth within the nerve-conduit assembly\u003c/h2\u003e \u003cp\u003eWhile light sheet fluorescence microscopy (LSFM) is commonly used for brain and nerve imaging, prior work indicated that LSFM is unsuitable for synthetic nerve conduits due to light scattering by the conduit walls, which prevents visualization of axons within the conduit (Fogli et al., \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e2019\u003c/span\u003e). We therefore used confocal microscopy with a 10x long-working distance objective to image the centimeter-scale nerve-conduit assemblies. Confocal imaging provided high spatial resolution, enabling precise quantification of individual fibers, including small-diameter C-type axons that may be unresolved by LSFM.\u003c/p\u003e \u003cp\u003eA multi-position holder allowed automated imaging of up to four samples in a single overnight session. Reconstructed 3D image stacks were processed using ilastik, a machine learning\u0026ndash;based classification and segmentation tool (Berg et al., \u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e2019\u003c/span\u003e). Residual conduit autofluorescence and background were subtracted by thresholding to isolate axonal GFP signals. To quantify regeneration, the z-stack was re-sliced orthogonally for an \u003cem\u003een face\u003c/em\u003e view of the nerve. Individual axons were detected and counted along the nerve axis using a spot detection algorithm, highlighting that only a small fraction of axons initiate regenerative growth \u0026mdash; one reason for poor functional recovery after traumatic PNI (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA-E).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003e \u003cb\u003e(A)\u003c/b\u003e Combined transmission light and EGFP fluorescence confocal imaging of the nerve-conduit assembly two weeks after PNI using a 10x long-working distance objective. A GFP-M mouse, which expresses GFP in a mosaic pattern in sensory and motor fibers (approximately 1% of all axons; (Feng et al., \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e2000\u003c/span\u003e)), was used. Dashed white lines indicate the proximal and distal nerve stumps within the conduit. \u003cb\u003e(B)\u003c/b\u003e GFP fluorescence channel alone. The yellow arrow indicates the typical drop in fluorescence at the conduit edge. \u003cb\u003e(C)\u003c/b\u003e Maximum intensity projection (MIP) of the grayscale image stack after pixel classification and subtraction of conduit autofluorescence. \u003cb\u003e(D)\u003c/b\u003e MIP of the grayscale image stack after pixel classification and thresholding. Regenerating nerve fibers in the distal nerve stump are indicated by yellow arrowheads. Orthogonal slices of the proximal and distal stumps are shown at positions marked by blue lines. \u003cb\u003e(E)\u003c/b\u003e Quantification of GFP-positive intersections per orthogonal slice, corresponding to axon number, along the nerve-conduit axis. The yellow arrow indicates the typical drop in fluorescence at the conduit edge. The light-blue curve presents data at each incremental position. The dark-blue curve has been smoothed over 50 \u0026micro;m intervals. Scale bar, 500 \u0026micro;m.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003ch2\u003eTime course of axon regeneration and macrophage dynamics after nerve transection and conduit-based repair\u003c/h2\u003e \u003cp\u003eWhile skin reinnervation after sciatic nerve crush has been investigated immunohistochemically (Navarro et al., \u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e1997\u003c/span\u003e), systematic morphological data for nerve transection followed by conduit repair are lacking. To address this, we performed an imaging time series at intervals ranging from 2 days to 6 weeks post-injury. Macrophages play a key role in Wallerian degeneration and axonal regeneration, clearing myelin and axonal debris and, together with repair Schwann cells, contributing to the formation of bands of B\u0026uuml;ngner (Li et al., \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e2022\u003c/span\u003e). To visualize macrophage dynamics, we crossed GFP-M mice with Cx3cr1\u003csup\u003eCreERT2\u003c/sup\u003e::ROSA26\u003csup\u003etdTomatofl/fl\u003c/sup\u003e animals, enabling tamoxifen-inducible tdTomato expression in the Cx3cr1⁺ macrophage lineage (Petzer et al., \u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e2018\u003c/span\u003e).\u003c/p\u003e \u003cp\u003eAxonal reorganization was first evident 2 days post-surgery, including growth cone formation, consistent with the approximately 48-hour latency required for neurons to reprogram into a regenerative state (Griffin et al., \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e2010\u003c/span\u003e). Bulk axonal regrowth initiated after 1 week, once Wallerian degeneration was largely complete. By 2 weeks, most axons had crossed the gap, with pioneering fibers extending into the distal stump; this pattern was consolidated at 4 and 6 weeks. Notably, chitosan autofluorescence quenching diminished at later time points, particularly in the 488 nm channel, limiting observations beyond 2 weeks (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA).\u003c/p\u003e \u003cp\u003eIn intact nerves, a resident macrophage population was observed. Following sciatic nerve injury (SNI), macrophage recruitment commenced within 1 week, peaking around 2 weeks in the conduit and reaching maximal density in the distal stump by 4 weeks. By 6 weeks, macrophage numbers had begun to decline, indicating clearance of macrophages from the lesion site (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eB).\u003c/p\u003e \u003cp\u003eIn summary, our time course captures all major events after nerve transection and conduit repair: macrophage recruitment within 1 week, initiation of axonal regrowth after 1 week, consolidation of axonal regeneration by 2 weeks, and onset of macrophage clearance by 6 weeks. We identify 2 weeks post-lesion as the optimal time point to analyze axonal regeneration within the conduit, coinciding with maximal macrophage density (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eA double-reporter mouse line generated by crossing GFP-M mice with the macrophage-specific tamoxifen-inducible tdTomato reporter line Cx3cr1\u003csup\u003eCreERT2\u003c/sup\u003e::ROSA26\u003csup\u003etdTomatofl/fl\u003c/sup\u003e was used to visualize axon and macrophage dynamics for up to six weeks after SNI. MIPs are shown for each time point. \u003cb\u003e(A)\u003c/b\u003e Progressive axonal fragmentation in the distal nerve stump, indicative of Wallerian degeneration, is visible at 4 days and 1 week after surgery (white arrow heads). By 2 weeks after SNI, all transected axons have been degraded, and the first regenerative fibers extend into the distal nerve stump (white arrows). \u003cb\u003e(B)\u003c/b\u003e Macrophages are present in the intact nerve and increase in number several days after SNI due to recruitment. By 6 weeks after SNI, macrophages are largely cleared from the lesion site. \u003cb\u003e(C)\u003c/b\u003e Combined axonal and macrophage signals. Scale bar, 500 \u0026micro;m. p.s., post-surgery.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec15\" class=\"Section2\"\u003e \u003ch2\u003eSensory-motor recovery after nerve transection and conduit-based repair\u003c/h2\u003e \u003cp\u003eAlthough axonal regeneration is actively ongoing within the conduit at 2 weeks, target reinnervation takes additional time. Functional recovery in this model has not been previously characterized. Based on in vivo axonal growth rates in young adult mice, approximately 1.9 mm/day for motor fibers and approximately 2.9 mm/day for sensory fibers (Verdu et al., \u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e2000\u003c/span\u003e), sensory axons are expected to reinnervate targets around 3 weeks post-injury, and motor fibers by about 4 weeks. Accounting for the initial 1\u0026ndash;2 week phase required to traverse the gap and enter the distal stump, we selected a 4\u0026ndash;7 week window to monitor sensory-motor performance, which should also align with the period most suitable for evaluating regenerative interventions at the lesion site.\u003c/p\u003e \u003cp\u003eSensory recovery was assessed longitudinally using the von Frey test from baseline to 6 weeks post-surgery. In the 6th week, operated animals exhibited responses comparable to baseline, whereas in the 4th week, the operated right hind limb showed significantly reduced responsiveness relative to the intact left hind limb for both withdrawal force and latency (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA, B). These results indicate that 3\u0026ndash;4 weeks post-lesion is the optimal window to evaluate sensory recovery.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eLongitudinal von Frey testing was performed from baseline to the 6th week post-surgery. \u003cb\u003e(A)\u003c/b\u003e Force required to elicit hind paw withdrawal. \u003cb\u003e(B)\u003c/b\u003e Latency to hind paw withdrawal. Individual data points represent the mean of three measurements per animal. Data are presented as mean\u0026thinsp;\u0026plusmn;\u0026thinsp;SD; n\u0026thinsp;=\u0026thinsp;12 before surgery; n\u0026thinsp;=\u0026thinsp;6 in the 4th week post-surgery; n\u0026thinsp;=\u0026thinsp;6 in the 6th week post-surgery. Statistical significance and p-value are indicated for p\u0026thinsp;\u0026le;\u0026thinsp;0.05. ns, not significant. p.s., post-surgery; LH, left hind paw; RH, right hind paw.\u003c/p\u003e \u003cp\u003eMotor recovery was assessed longitudinally using the CatWalk system from baseline to 7 weeks post-surgery. After SNI, animals shifted from a digitigrade to a plantigrade gait, reflected in increased print length (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA). Print width decreased due to the absence of outer toe prints, preventing calculation of toe spread (TS) and hence the sciatic functional index (SFI), a finding at odds with crush models where these measures are typically obtained (Fey et al., \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e2010\u003c/span\u003e; Vogelaar et al., \u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e2004\u003c/span\u003e). Most paw-print parameters, including duty cycle, print area, intermediate toe spread, maximum intensity (at %), and maximum contact area (at %), improved between the 5th and 7th week but remained significantly impaired relative to baseline, indicating mid-phase motor recovery (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eC).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003ePaw-print parameters from a longitudinal CatWalk analysis performed from baseline to the 7th week post-surgery. \u003cb\u003e(A)\u003c/b\u003e Representative paw-prints of the right hind limb before surgery and in the 5th and 7th week post-surgery. Print length (PL), toe spread (TS), and intermediate toe spread (ITS) are shown by white lines. The red arrow marks the tarsal (heel) pad, which is typically visible only in operated animals. \u003cb\u003e(B)\u003c/b\u003e Schematic ventral view of the operated animal with the SNI located in the right hind limb. The mouse schematic was adjusted from BioRender. \u003cb\u003e(C)\u003c/b\u003e Paw-print parameters of the affected right hind limb compared with the intact left hind limb. Individual data points represent the mean of three compliant runs per animal. Data are presented as mean\u0026thinsp;\u0026plusmn;\u0026thinsp;SD; n\u0026thinsp;=\u0026thinsp;12 before surgery; n\u0026thinsp;=\u0026thinsp;6 in the 5th week post-surgery (except ITS); n\u0026thinsp;=\u0026thinsp;6 in the 7th week post-surgery (except ITS). Statistical significance and p-value are indicated for p\u0026thinsp;\u0026le;\u0026thinsp;0.05. p.s., post-surgery; LH, left hind paw; RH, right hind paw.\u003c/p\u003e \u003cp\u003eDynamic gait analysis of all four limbs showed substantial deviations in phasing from baseline in the 5th week, with significant improvement in the 7th week (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA, B, D, F). Regularity indices of phase dispersions remained stable, suggesting coordinated, rather than chaotic, gait disruption (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eC, E, G).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eGait parameters from a longitudinal CatWalk analysis performed from baseline to the 7th week post-surgery. \u003cb\u003e(A)\u003c/b\u003e Girdle phase dispersion LH-RH (theoretical baseline value 50%). \u003cb\u003e(B)\u003c/b\u003e Schematic ventral view of the operated animal with the SNI located in the right hind limb and the four paws marked with different colours. The mouse schematic was adjusted from BioRender. \u003cb\u003e(C)\u003c/b\u003e Regularity index of the girdle phase dispersion LH-RH (ideal value 1.00). \u003cb\u003e(D)\u003c/b\u003e Diagonal phase dispersion LF-RH (theoretical baseline value 0%). \u003cb\u003e(E)\u003c/b\u003e Regularity index of the diagonal phase dispersion LF-RH (ideal value 1.00). \u003cb\u003e(F)\u003c/b\u003e Ipsilateral phase dispersion RF-RH (theoretical baseline value 50%). \u003cb\u003e(G)\u003c/b\u003e Regularity index of the ipsilateral phase dispersion RF-RH (ideal value 1.00). Individual data points represent the mean of three compliant runs per animal. Data are presented as mean\u0026thinsp;\u0026plusmn;\u0026thinsp;SD; n\u0026thinsp;=\u0026thinsp;12 before surgery; n\u0026thinsp;=\u0026thinsp;5 in the 5th week post-surgery; n\u0026thinsp;=\u0026thinsp;6 in the 7th week post-surgery. Statistical significance and p-value are indicated for p\u0026thinsp;\u0026le;\u0026thinsp;0.05. p.s., post-surgery; LF, left front paw; RF, right front paw; LH, left hind paw; RH, right hind paw.\u003c/p\u003e \u003cp\u003eParameters unaffected by SNI included stride length, step sequence regularity, and base of support of the fore- and hind limbs (Suppl. Figure\u0026nbsp;1A-E).\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec16\" class=\"Section2\"\u003e \u003ch2\u003eSupplemental Fig.\u0026nbsp;1 CatWalk test: Additional gait parameters\u003c/h2\u003e \u003cp\u003eAdditional gait parameters are shown from the longitudinal CatWalk analysis performed from baseline to the 7th week post-surgery. \u003cb\u003e(A)\u003c/b\u003e stride length. \u003cb\u003e(B)\u003c/b\u003e Run speed. \u003cb\u003e(C)\u003c/b\u003e Regularity index of the step sequence (ideal value 1.00). \u003cb\u003e(D)\u003c/b\u003e Base of support (BoS) of the front paws. \u003cb\u003e(E)\u003c/b\u003e BoS of the hind paws. Individual data points represent the mean of three compliant runs per animal. Data are presented as mean\u0026thinsp;\u0026plusmn;\u0026thinsp;SD; n\u0026thinsp;=\u0026thinsp;12 before surgery; n\u0026thinsp;=\u0026thinsp;5\u0026ndash;6 in the 5th week post-surgery; n\u0026thinsp;=\u0026thinsp;6 in the 7th week post-surgery. Statistical significance and p-value are indicated for p\u0026thinsp;\u0026le;\u0026thinsp;0.05. p.s., post-surgery; LH, left hind paw; RH, right hind paw.\u003c/p\u003e \u003cp\u003eOverall, 5\u0026ndash;7 weeks post-lesion represents an appropriate time window to assess motor recovery, capturing ongoing regeneration before the plateau phase, and allowing room to detect improvements from potential regenerative interventions.\u003c/p\u003e \u003c/div\u003e"},{"header":"DISCUSSION","content":"\u003cp\u003eWe introduce and characterize a mouse model of PNI that mimics severe lesions involving neurotmesis and incorporates clinically relevant microsurgical repair strategies. By combining nerve transection with conduit-based repair, this model bridges the gap between growing clinical practice and experimental research.\u003c/p\u003e \u003cp\u003eDespite the increasing clinical use of synthetic nerve guidance conduits, experimental PNI research still relies on crush lesions or direct end-to-end coaptation after transection, both of which have limited clinical relevance. A systematic review of nerve transection studies identified 49 animal studies employing various coaptation techniques, but none using conduit-based repair (Vela et al., \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). Similarly, a systematic review of experimental PNI models and motor recovery cited 223 studies, almost exclusively based on crush or direct ligation paradigms (Heinzel et al., \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). In contrast, our nerve transection and conduit-based repair model closely reflects clinical practice and provides a framework for testing the therapeutic potential of bioactive compounds.\u003c/p\u003e \u003cp\u003eWhile novel conduit designs, such as 3D-printed scaffolds, are actively being developed, these approaches remain exploratory and lack standardization and commercial availability (Kim et al., \u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e2024\u003c/span\u003e; Larijani et al., \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e2024\u003c/span\u003e; Park et al., \u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e2022\u003c/span\u003e; Wan et al., \u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e2025\u003c/span\u003e; Wang et al., \u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e2024\u003c/span\u003e). Chitosan nerve guidance conduits provide a practical solution to this need. They are commercially available, standardized, and exhibit excellent biocompatibility and biodegradability, which has led to their increasing use in clinical settings (Bocker et al., \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e2022\u003c/span\u003e; Neubrech et al., \u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e2018\u003c/span\u003e). Chitosan interacts favorably with cells of the neural microenvironment and undergoes gradual degradation after implantation. In contrast to clinical application, we modified the conduit by halving it prior to implantation. This adjustment not only accommodates the smaller diameter of the mouse sciatic nerve but also enables the delivery of soluble, bioactive compounds directly to the lesion site via a hydrogel matrix. In our hands, PuraMatrix supported axonal regeneration more effectively than Matrigel (data not shown), making it a suitable substrate for this application.\u003c/p\u003e \u003cp\u003eA technical challenge of this approach is the intrinsic autofluorescence of chitosan. Chitosan is a chitin derivative that was originally prepared from crab tendons (Yamaguchi et al., \u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e2003\u003c/span\u003e) and has been developed for its application as a material for nerve guidance conduits (Boecker et al., \u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e2019\u003c/span\u003e). As a polysaccharide, chitosan is not susceptible to lipid-based clearing protocols. We developed a protocol to quench conduit autofluorescence, allowing visualization of regenerating axons within the conduit using long-working-distance confocal microscopy (Fogli et al., \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e2019\u003c/span\u003e). However, this quenching effect is transient; autofluorescence gradually re-emerges over time, particularly in the 488 nm channel, limiting imaging at later time points. To improve signal-to-noise ratio, we developed a \u0026ldquo;peripheral perfusion\u0026rdquo; technique, in which perfusion via the vena cava and drainage through the femoral vein enhances blood removal from the hind limb compared to conventional transcardiac perfusion. Combined with transgenic fluorescent reporters and a shortened CUBIC clearing protocol (Susaki et al., \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e2014\u003c/span\u003e), this approach enables robust imaging of axonal and cellular dynamics while preserving GFP fluorescence.\u003c/p\u003e \u003cp\u003eFunctional recovery is a central outcome measure in experimental PNI models. While electrophysiological and electromyographic methods have traditionally been used (Navarro, \u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e2016\u003c/span\u003e), computational gait analysis has become increasingly important, with the CatWalk system providing a precise and detailed quantitative assessment of locomotion (Chen et al., \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e2017\u003c/span\u003e; Heinzel et al., \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e2020\u003c/span\u003e). Based on known time courses of nerve regeneration, we selected a 4\u0026ndash;7 week post-injury window to assess sensory and motor recovery. Previous studies have shown that reinnervation of neuromuscular junctions can occur as early as 2 weeks after injury in both crush and transection/coaptation models (Bauder \u0026amp; Ferguson, \u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2012\u003c/span\u003e; Vannucci et al., \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e2019\u003c/span\u003e), with sensory recovery preceding motor recovery (Navarro \u0026amp; Kennedy, \u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e1991\u003c/span\u003e; Navarro et al., \u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e1994\u003c/span\u003e). In general, axonal regeneration is expected to proceed more slowly after nerve transection and coaptation than after a crush lesion. A recent study in the rat directly compared these paradigms and found that functional recovery following transection and coaptation was delayed by approximately one week relative to a crush lesion. Specifically, sensory recovery began about 2 weeks after the crush lesion, whereas motor recovery did not commence before 3 weeks post-lesion (Wang et al., \u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e2023\u003c/span\u003e). Consistent with these findings and taking into account an approximately one-week initiation phase required for axons to navigate the conduit microenvironment, we observed progressive functional improvement over a period of 4\u0026ndash;7 weeks post-surgery. Within this window, sensory recovery was best assessed at 3\u0026ndash;4 weeks, whereas motor recovery was most appropriately evaluated at 5\u0026ndash;7 weeks after injury. Importantly, both time windows fall within the dynamic phase of regeneration, prior to the plateau phase, thereby providing sensitivity to detect therapeutic effects.\u003c/p\u003e \u003cp\u003eA key difference between our model and conventional crush injury models lies in motor recovery patterns. Following transection and conduit repair, animals adopted a plantigrade gait and failed to use their outer toes even at later time points, precluding calculation of the sciatic functional index (SFI). In contrast, crush injury models typically exhibit rapid and near-complete recovery due to preservation of epineurium and endoneurial guidance structures, with functional restoration achieved within 3\u0026ndash;4 weeks, including recovery of toe spread (Fey et al., \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e2010\u003c/span\u003e; Vogelaar et al., \u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e2004\u003c/span\u003e). In the transection model, regenerating axons must bridge a physical gap and re-establish target connections without intact guidance structures, resulting in delayed motor recovery. Impaired intrinsic foot muscle function likely underlies the absence of toe spreading in our model. This highlights an important distinction between injury paradigms and suggests that foot muscle activity is particularly sensitive to severe nerve injuries. At the same time, restoration of toe spread is indicative of fine motor recovery and may therefore serve as a sensitive endpoint for evaluating therapeutic interventions.\u003c/p\u003e"},{"header":"CONCLUSION","content":"\u003cp\u003eThe severity of traumatic peripheral nerve injury is often underestimated, partly due to the assumption that peripheral axons regenerate readily after injury. However, in cases of neurotmesis, successful regeneration depends on microsurgical repair and remains inefficient, frequently resulting in incomplete sensory and motor recovery and long-term deficits. This reflects the limited proportion of regenerating axons that successfully reach their targets and re-establish functional connections. Consequently, there is a pressing need for therapeutic strategies that enhance axonal regeneration and functional recovery.\u003c/p\u003e \u003cp\u003eProgress in this field is hindered by the lack of experimental models that adequately reflect clinical conditions and allow for the effective delivery of bioactive compounds to the lesion site. To address these limitations, we developed a mouse model of PNI that closely mimics clinically relevant injury and repair paradigms. By combining sciatic nerve transection with tension-free, conduit-based repair, this lesion model also enables local delivery of bioactive agents to the lesion site, thereby providing a platform for preclinical testing of therapeutic interventions.\u003c/p\u003e \u003cp\u003eAll components of this model, including the transgenic reporter lines, microsurgical procedures, conduit preparation, tissue processing, and imaging workflows, are readily implementable (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e). We propose this paradigm as a standardized and clinically relevant reference model for experimental PNI research.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eSchematic overview of the experimental paradigm described in this study, including sciatic nerve transection, conduit implantation, tissue processing, imaging, and axon quantification.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e \u003ch2\u003eCOMPETING INTERESTS\u003c/h2\u003e \u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e \u003c/p\u003e\u003ch2\u003eFUNDING\u003c/h2\u003e \u003cp\u003eThe Vienna BioCenter Core Facilities (VBCF) Preclinical Phenotyping Facility acknowledges funding from the Austrian Federal Ministry of Education, Science \u0026amp; Research, and the City of Vienna. SM was supported by VIB, the VIB BioImaging Core Leuven, and the Research Foundation \u0026ndash; Flanders (FWO) through grants FWOI000123N and FWOI001322N. Additionally, SM was supported by KU Leuven (KA/24/041). RS was supported by a research project grant from the Austrian Science Fund FWF (P33411-B).\u003c/p\u003e\u003ch2\u003eAuthor Contribution\u003c/h2\u003e\u003cp\u003eW.U., L.R., and F.S. refined and performed the nerve transection protocol, including conduit-based repair, and carried out tissue processing and clearing; S.M. imaged the nerve samples and processed images; S.M. and R.S. analysed the imaging data; S.B. carried out sensory-motor tests and analysed the data; W.U. and R.S. analysed sensory-motor data; E.D. contributed to the perfusion protocol and to macrophage imaging; S.M. and R.S. designed experiments and managed the project; R.S. conceived the study and wrote the manuscript. All authors reviewed the manuscript.\u003c/p\u003e\u003ch2\u003eAcknowledgement\u003c/h2\u003e\u003cp\u003eWe thank Klaus Kraitsy for excellent assistance with aspects of the nerve transection experiments. We are grateful to Matilde Bongio (somersault18:24) for outstanding graphical rendering. The GFP-M mouse line was originally generated in the laboratories of Joshua R. Sanes and Jeff W. Lichtman.\u003c/p\u003e\u003ch2\u003eData Availability\u003c/h2\u003e\u003cp\u003eAll data are available from the authors upon request.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\u003cli\u003e\u003cspan\u003eAllodi, I., Udina, E., \u0026amp; Navarro, X. (2012, Jul). Specificity of peripheral nerve regeneration: interactions at the axon level. \u003cem\u003eProg Neurobiol\u003c/em\u003e, \u003cem\u003e98\u003c/em\u003e(1), 16\u0026ndash;37. \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://doi.org/10.1016/j.pneurobio.2012.05.005\u003c/span\u003e\u003cspan address=\"10.1016/j.pneurobio.2012.05.005\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBauder, A. 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Perspectives on the Novel Multifunctional Nerve Guidance Conduits: From Specific Regenerative Procedures to Motor Function Rebuilding. \u003cem\u003eAdv Mater\u003c/em\u003e, \u003cem\u003e36\u003c/em\u003e(14), e2307805. \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003ehttps://doi.org/10.1002/adma.202307805\u003c/span\u003e\u003cspan address=\"10.1002/adma.202307805\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e\u003c/span\u003e\u003c/li\u003e\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"npj-regenerative-medicine","isNatureJournal":false,"hasQc":false,"allowDirectSubmit":false,"externalIdentity":"npjregenmed","sideBox":"Learn more about [npj Regenerative Medicine](http://www.nature.com/npjregenmed/)","snPcode":"41536","submissionUrl":"https://mts-npjregenmed.nature.com/cgi-bin/main.plex","title":"npj Regenerative Medicine","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"ejp","reportingPortfolio":"NPJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"","lastPublishedDoi":"10.21203/rs.3.rs-9254882/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-9254882/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eIn contrast to axons of the central nervous system, peripheral nerve fibers regenerate after traumatic injury, but often not to a clinically satisfactory extent, as only a small proportion of axons achieve long-distance regrowth. While autologous nerve transplantation remains the gold standard for tension-free repair, advances in biomaterials have led to increasing clinical use of synthetic nerve guidance conduits. Despite these developments, experimental research on traumatic peripheral nerve injury (PNI) continues to rely on crush lesions and direct end-to-end repair models, both of which have limited clinical relevance. Here, we present a mouse model of traumatic PNI that closely mimics clinical conditions and enables translationally relevant investigations. Our model combines sciatic nerve transection with conduit-based repair using a commercially available, standardized chitosan nerve guidance conduit that is longitudinally opened prior to implantation. This design allows for the direct application of soluble bioactive compounds to the lesion site. We further detail methods for visualizing and quantifying axonal regeneration at single-fiber resolution within the conduit and provide a time-course analysis of axonal regrowth and macrophage dynamics during Wallerian degeneration by making use of transgenic reporter mice. Functional recovery is assessed using sensory and motor performance tests, and time windows for applying bioactive compounds are defined. Together, this model recapitulates key features of clinical nerve injury and repair, and provides a platform for preclinical testing of regenerative therapies. We propose it as a standardized reference model for experimental PNI research.\u003c/p\u003e","manuscriptTitle":"A clinically relevant mouse model for traumatic peripheral nerve injury and repair","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-04-06 19:26:09","doi":"10.21203/rs.3.rs-9254882/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"editorInvitedReview","content":"","date":"2026-05-18T09:35:28+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-05-06T09:54:02+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"69253130414863330997170644662633218609","date":"2026-04-30T07:35:46+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"149475945251858538104970093479240303662","date":"2026-04-27T02:20:27+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2026-03-31T14:27:40+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2026-03-31T12:52:02+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2026-03-31T05:32:21+00:00","index":"","fulltext":""},{"type":"submitted","content":"npj Regenerative Medicine","date":"2026-03-28T19:06:09+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"npj-regenerative-medicine","isNatureJournal":false,"hasQc":false,"allowDirectSubmit":false,"externalIdentity":"npjregenmed","sideBox":"Learn more about [npj Regenerative Medicine](http://www.nature.com/npjregenmed/)","snPcode":"41536","submissionUrl":"https://mts-npjregenmed.nature.com/cgi-bin/main.plex","title":"npj Regenerative Medicine","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"ejp","reportingPortfolio":"NPJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"29f039e7-6cce-4b84-90e6-81c366a825a1","owner":[],"postedDate":"April 6th, 2026","published":true,"recentEditorialEvents":[{"type":"editorInvitedReview","content":"","date":"2026-05-18T09:35:28+00:00","index":62,"fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-05-06T09:54:02+00:00","index":61,"fulltext":""},{"type":"reviewerAgreed","content":"69253130414863330997170644662633218609","date":"2026-04-30T07:35:46+00:00","index":60,"fulltext":""}],"rejectedJournal":[],"revision":"","amendment":"","status":"under-review","subjectAreas":[{"id":65627000,"name":"Biological sciences/Biotechnology"},{"id":65627001,"name":"Physical sciences/Engineering"},{"id":65627002,"name":"Health sciences/Medical research"},{"id":65627003,"name":"Biological sciences/Neuroscience"}],"tags":[],"updatedAt":"2026-04-06T19:26:09+00:00","versionOfRecord":[],"versionCreatedAt":"2026-04-06 19:26:09","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-9254882","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-9254882","identity":"rs-9254882","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}