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Method Article

Novel Surgical Rodent Model for Studying Neuroma Pain Treatment Options using Targeted Muscle Reinnervation Through the Saphenous Nerve

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DOI:

10.3791/69152

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November 14th, 2025

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1Department of Plastic and Reconstructive Surgery, University Medical Center Utrecht, Utrecht University

In This Article

Summary

This protocol presents a reproducible rat model using sensory nerves to mimic human neuroma pain and assess surgical treatments, offering a valuable tool for translational neuroma research.

Abstract

This protocol describes a novel surgical model to study nerve pain and surgical treatment in rats. Traditional neuroma models use mixed nerves, although it is most often the case that sensory nerves give rise to painful neuromas in the clinical setting. By selecting the saphenous nerve, this approach reflects the clinical reality of neuroma formation and provides a relevant platform for evaluating surgical strategies. This method aims to replicate the pathophysiology of human neuroma formation and to evaluate a common surgical intervention for neuroma pain, particularly targeted muscle reinnervation (TMR), a widely used intervention for neuroma-associated pain. The article provides a detailed, step-by-step description of the procedure, beginning with animal preparation and perioperative care. Surgical technique is explained with attention to anatomical landmarks, instrument requirements, and technical considerations that improve accuracy and consistency. Postoperative monitoring and welfare measures are outlined to ensure adherence to ethical standards and reproducibility across laboratories. This model is reproducible, technically accessible, and translationally relevant, as it mimics the clinical reality of sensory neuroma formation. It provides a platform for evaluating surgical strategies and preclinical neuroma research.

Introduction

Neuromas are painful nerve tumors formed after nerve injury1,2. They represent a significant clinical problem with far-reaching societal consequences and increased medical costs3,4,5,6,7. Treatment options such as pharmacological therapy are often symptomatic, addressing pain but failing to resolve its underlying causes8. Surgical interventions have been shown to have variable success rates (56%-82%), and the mechanisms driving painful neuroma formation remain poorly understood9,10,11.

Pain from neuromas is thought to arise from sensory fascicles in the damaged nerve, underscoring the importance of modeling purely sensory contributions to neuroma-associated pain12. Animal models remain an essential platform for exploring the complex mechanisms underlying neuroma pain and also for developing effective surgical therapies13. However, most available animal models use mixed motor sensory nerves, which limits their translational applicability, since approximately 74% of painful neuromas in the clinical setting originate from purely sensory nerves14,15.

Several animal models have been developed to simulate neuropathic pain and evaluate potential interventions16. Firstly, the chronic constriction injury (CCI) model has been described17. This model induces partial nerve damage through loosely tied ligatures around the sciatic nerve, mimicking neuropathic pain conditions. It provides insight into peripheral sensitization, axonal degeneration, inflammatory responses, and central changes in the spinal cord, and is widely used to evaluate potential analgesic therapies17,18. Secondly, the Spared Nerve Injury (SNI) induces injury to two of the three terminal branches of the sciatic nerve (tibial and common peroneal nerves), leaving the remaining sural nerve intact19. Thirdly, the tibial neuroma transposition (TNT) model, developed by Dorsi et al. The tibial nerve is surgically transected and transposed subcutaneously to facilitate the evaluation of neuroma formation and associated pain16,20. This model allows for primary hypersensitivity assessment by Von Frey stimulation of the skin overlying the transposed neuroma at the pretibial site, and secondary hypersensitivity assessment in the adjacent sural or residual tibial nerve dermatomes. The TNT model was surgically designed to replicate neuroma development, without incorporating any surgical or non-surgical intervention aimed at preventing or treating neuroma-related pain20. As such, it does not allow evaluation of therapeutic strategies, like targeted muscle reinnervation (TMR), in the primary setting (Figure 1).

TMR is a technique that connects a transected (sensory) nerve to an expendable motor branch of a target after resecting the neuroma. This concept uses the principle of give the nerve something to do, whereby the nerve will regenerate into a nerve connected to a muscle target. The philosophy aims to prevent the formation of a neuroma per se by allowing the nerve to regenerate into a denervated muscle11.

In the present study, we describe a rodent model involving the transection of the saphenous nerve. This purely sensory nerve is transected, transposed to the gastrocnemius muscle, and either left untreated (saphenous neuroma transposition, SNT) or coapted to a motor branch (TMR). This model enables investigation of the effects of TMR applied to the saphenous nerve on secondary mechanical hypersensitivity in adjacent dermatomes of anatomically intact nerves, specifically within the tibial territory on the central plantar surface of the hind paw (Figure 1)21,22. This anatomical context also highlights the clinical relevance in patients. Due to its superficial anatomical course, the saphenous nerve is particularly vulnerable to iatrogenic or traumatic injury, especially following knee surgery, often resulting in neuropathic pain; notably, 55.2% of patients report persistent symptoms despite revision surgery such as neurolysis or neurectomy23,24.

This study aims to establish a reproducible rodent model in order to replicate the clinical setting of painful neuroma formation. This model is particularly suited for investigating pure sensory neuroma pathophysiology and the effects of TMR, while its primary outcome measure focuses on secondary mechanical hypersensitivity.

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Protocol

This protocol aligns with ethical animal research guidelines and emphasizes its translational potential. Project number: AVD1150020198824.

1. Von Frey measurements

  1. Perform Von Frey testing of the plantar surface of the hind paw according to the adapted protocol of Brakkee et al. to assess secondary mechanical hypersensitivity20.
    1. Determine the baseline sensitivity of the innervation areas of the tibial and sural nerves on the plantar surface of the hind paw. Perform three baseline measurements on separate days using Von Frey filaments to obtain reliable and reproducible data20,25,26,27,28,29,30.
    2. Acclimate the rats to mesh-bottom cages for at least 5-7 days before baseline measurements. Handle the animals daily to reduce stress responses. Baseline Von Frey data are provided in Supplementary Table 1. Assess surgery-induced hypersensitivity in the tibial and sural innervation areas using Von Frey testing starting 7 days after surgery.
  2. Position the animal calmly in the testing cage and confirm that spontaneous movements are minimal before starting.
  3. Apply a series of calibrated monofilaments (0.6 g, 1 g, 1.4 g, 2 g, 4 g, 6 g, 8 g, 10 g, 15 g) perpendicular to the skin until the filament bends slightly. Hold contact for approximately 3 s.
    NOTE: Do not perform testing when the animal is grooming, urinating, standing, or defecating. Conduct testing only when the weight is evenly distributed on all four paws.
  4. Vary the order of dermatomes across animals to avoid systematic bias as described below.
    Sural nerve territory: lateral plantar surface, close to the hairy-glabrous border.
    ​Tibial nerve territory: central plantar surface, avoiding the pads.
  5. Use the up-down method: start with the 4 g filament. If a withdrawal occurs (sudden paw lift, flinch, licking, or vocalization), apply the next lower force; if no response occurs, apply the next higher force. Continue until 5-6 responses after the first change in behavior are obtained. Record a positive response as any brisk withdrawal, flinch, licking, or vocalization. Record the absence of these behaviors as a negative response.
    1. Calculate the 50% withdrawal threshold using the Dixon up-down method, which determines the force at which there is a 50% probability of a withdrawal response based on a sequence of increasing and decreasing von Frey filament applications31.
  6. For assessment of secondary hyperalgesia, compare post-operative thresholds in the tibial and sural zones to baseline values and to the contralateral paw.
    NOTE: Conditioned place preference (CPP) is a validated behavioral paradigm for assessing ongoing pain and treatment effect in animal models of peripheral nerve injury, particularly when mechanical testing is not feasible due to the anatomical location of neuroma formation, such as deep in the thigh32,33,34. It is mentioned here as background information only and was not applied in the present study.

2. Preoperative preparation

  1. Book an animal surgery room and prepare/order sterile instruments (See Table of Materials).
  2. Analgesia, anesthesia, and preparation
    NOTE: This study was conducted on 12 (6 female and 6 male) Sprague Dawley rats that were 12 weeks old.
    1. Inject 5.0 mg/kg of analgesic carprofen subcutaneously in the abdominal region 1 h prior to surgery.
    2. Anesthetize the animal using 5% isoflurane for induction, maintaining anesthesia with 2%-3% isoflurane.
    3. Place the animal on its back with its head either left or right to be operated on, opposite to the surgeon's position.
    4. Place the animal on a heating pad set to 37 °C to maintain its body temperature during the procedure. Distribute the heating surface evenly and prevent overheating. Monitor temperature with a rectal thermal probe, secured to the tail using paper tape. Adjust the heating pad as necessary to maintain normothermia.
      NOTE: This step is crucial for preventing hypothermia induced by anesthesia and ensuring physiological stability throughout the surgery.
    5. Apply a sterile ophthalmic ointment to the eyes to prevent dryness caused by gas anesthesia.
      NOTE: This step ensures that the corneal surface remains moist and protected during the procedure, minimizing the risk of ocular irritation or damage.
    6. Shave the surgical site from thigh to ankle using an electric razor. After shaving, use the adhesive side of a piece of tape to remove any remaining loose hairs for optimal preparation of the skin.
    7. Disinfect the shaved area with three alternating rounds of 0.5% chlorhexidine digluconate using a sterile gauze pad. Allow each application to dry completely before proceeding to the next round.
      NOTE: Three alternating rounds provide thorough sterilization of the surgical site and minimize infection risk.
    8. Verify anesthesia by gently pinching the webspace of the animal's foot with forceps. Absence of withdrawal indicates suppressed and adequate anesthesia.
      NOTE: Proceed only when reflexes are absent to confirm effective anesthesia and compliance with ethical standards for minimizing animal distress during surgery. Proceed only when you can confirm that reflexes are absent.

3. Surgical technique

  1. Place the animal in a supine position and rotate the operative leg laterally to expose the surgical site.
  2. Under 6x microscope magnification, mark the incision line using a sterile skin marker. Inject 0.05 mL of lidocaine 1% subcutaneously using a 1 mL syringe with a 25G needle along the marked line to provide local anesthesia and reduce surgical site pain. Distribute the injection evenly and confirm the absence of any immediate adverse reactions before proceeding. This step minimizes discomfort and improves surgical precision.
  3. Create an 8-10 mm longitudinal dermal incision over the medial hind limb using a scalpel 15 blade (Figure 2A). Only the dermis should be incised.
    NOTE: Extra care should be taken regarding the depth of the incision. If possible, initiate the excision and continue along the marked line using scissors to open the skin. As the saphenous nerve lies directly underneath the skin, meticulous attention is given to avoid a deep incision (Figure 2B).
  4. Identify the superficial neurovascular bundle, including the saphenous artery, vein, and nerve. Carefully pass and isolate the saphenous nerve by needle dissection to prevent accidental damage. Use electrocautery if necessary to control minor bleeding from surrounding vessels.
    1. Perform needle dissection by orienting the bevel of a fine needle towards the operator and gently sliding the tip along the tissue plane. This technique creates a sharp edge at the underside of the needle tip, allowing precise separation of delicate connective tissue without exerting excessive traction or pressure on adjacent structures.
  5. Increase isoflurane anesthesia to 5% prior to nerve transection. After completing the transection, reduce the isoflurane anesthesia back to 2%-3% to maintain adequate but controlled anesthesia levels.
  6. Change the microscope magnification to 16x.
  7. When approaching the adductor muscles medially and performing careful separation, a perpendicularly crossing vein, the vena caudalis femoris, is encountered centrally within the dissection area, accompanying the ramus nervi obturatorii. This vein becomes visible when dissecting the adductors longitudinally deeper towards the medial motor branch of the tibial nerve, which innervates the gastrocnemius muscle. Perform careful dissection either medially or laterally to preserve the venous integrity before establishing the coaptation field.
    NOTE: The vena caudalis femoris crosses perpendicular to the fiber direction of the adductor muscles; take care to avoid placing excessive traction on this vessel to prevent damage and unnecessary blood loss.
  8. Apply 1 mA stimulation with the nerve stimulator to assess muscle activity to the motor branch of the tibial nerve and confirm a response in the gastrocnemius muscle.
  9. During the preparation of the recipient nerve, dissect precisely along its trajectory and transect it with microsurgical scissors less than 3 mm from the muscle surface. Place the coaptation site close to the motor branch's muscle entry point to facilitate optimal axonal regeneration. Increase isoflurane to 5% briefly when transecting the nerve.
    NOTE: This technique ensures that the coaptation site remains near the insertion of the motor branch into the muscle, facilitating optimal axonal regeneration and functional recovery in TMR procedures. An increased distance between the coaptation site and the entry point of the motor branch may lead to axonal dispersion and impaired signal conduction.
  10. Tunnel the saphenous nerve through the adductor muscles and coapt it with the medial motor branch of the tibial nerve using 11.0 nylon sutures (Figure 3). Make two sutures and ensure the sutures do not obstruct the interface to allow for effective neural coaptation. For untreated controls, after tunneling, do not perform coaptation; instead, transpose the distal stump of the saphenous nerve and secure it to the gastrocnemius muscle fascia.
    NOTE: Two sutures should be sufficient to establish an optimal contact surface between the two nerves. A single suture is not suitable since it may cause one of the nerves to rotate.
  11. Adjust the microscope magnification to 6x before proceeding with skin closure. Use 8-0 sutures to approximate the epidermal edges with an intradermal technique. Maintain minimal tension on the wound. After closure, clean the skin with 0.9% NaCl using a sterile cotton swab to remove residual debris and maintain an optimal healing environment.

4. Postoperative care

  1. Place the animal in a clean, temperature-controlled cage with a heating pad partially positioned beneath the cage to maintain optimal warmth. Provide access to food and water. Adjust the lighting to a dim setting and shield the cage from the bright lights of the operating room.
    NOTE: These measures minimize stress and support smooth recovery after surgery.
  2. After waking up, the animal can be returned to its shared cage after approximately 40 min. Confirm full alertness and normal behavior before reintroducing to cage companions. This minimizes the risk of injury or stress during reintegration.
  3. Administer a dose of carprofen (5.0 mg/kg) in the abdominal region subcutaneously at 24 h and 48 h after surgery.
  4. Monitor for weight loss no greater than 15%. Provide adequate hydration and food intake.

5. Waste disposal

  1. Dispose of sharps immediately in an approved sharps container. Collect anesthesia waste in accordance with institutional biosafety guidelines. Handle animal remains according to institutional and national regulations for biological waste disposal.

6. Specimen collection and euthanasia

  1. At 6 weeks after surgery, euthanize the rats after collection of tissue samples for optimal differentiation between nerve and surrounding tissue.
  2. Induce anesthesia and administer analgesia as described in steps 2.2.1 to 2.2.5.
  3. Reopen the previous incision line on the medial thigh with a scalpel. Extend the incision by 2 cm if required. Perform blunt dissection through the subcutaneous layer.
  4. Retract or divide the adductor muscles to expose the saphenous nerve.For reconstructed nerves: identify the regenerated segment and any neuroma. Excise the coaptation site together with 2 mm of proximal and distal nerve.For untreated controls: identify the distal stump of the saphenous nerve and excise a 4 mm segment.
  5. Transfer the specimen directly into fixative. Embed longitudinally in paraffin or resin for morphological analysis.
  6. Euthanize the animal under terminal anesthesia (5% isoflurane) by cardiac puncture. Handle all tissue samples according to standard laboratory protocols to preserve structural and biochemical integrity for subsequent analysis (Figure 4).

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Results

Assessment of secondary hypersensitivity (log-transformed 50% withdrawal thresholds) showed group differences in the tibial dermatome (Figure 5A). Transection of the saphenous nerve without treatment (SNT) resulted in the highest sensitivity across the 6-week follow-up. In contrast, animals treated with TMR displayed a consistent trend toward reduced hypersensitivity in the tibial territory. The divergence between groups was most evident at week 2, when the mean 50% withdrawal threshold in t...

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Discussion

This approach has several strengths. The stepwise description allows reproducibility across laboratories, the procedure can be performed efficiently with microsurgical skills, and the use of a purely sensory nerve enhances its translational relevance. Together, these aspects make the model practical for comparative studies and useful for testing new interventions. Critical steps, such as maintaining aseptic conditions, precise dissection of the saphenous nerve, and careful handling of the vascular structures like the ven...

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Disclosures

Research performed by a PhD student (CB) supported by an Axogen grant (no product relationship). All the other authors have no conflicts of interest to disclose.

Acknowledgements

The authors gratefully acknowledge Sabine Versteeg for her contribution to behavioral pain assessments. Anja van der Sar and Trudy Oosterveld-Romijn (Common Animal Laboratory, Gemeenschappelijk Dieren Laboratorium) are thanked for their technical support in preparing surgical and microscopy facilities and for their dedicated animal care. The authors also thank Wim van Hecke for his expert evaluation of the collected nerve specimens.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
CarprofenLocal Veterinary Pharmacyn/aThe local veterinary pharmacy prepares the carprofen dilution
Cotton swabsNobamed974255—
Electric razorAesculapGT421
ElectrocauteryFine Science Tools18010-00—
Ethanol 70%Interchema BV400406—
Ethilon 10.0Johnson & JohnsonBV130-3: code 2820G
Ethilon 11.0Johnson & JohnsonBV50-3L code 2881G
Ethilon 4.0Johnson & Johnson1854G: code 1854G-
Ethilon 6.0Johnson & JohnsonBV130-5: code 2809G
Ethilon 8.0Johnson & JohnsonBV130-5: code 2808G—
Ethilon 9.0Johnson & JohnsonBV130-3: code 2819G
Examination gloves nitrileSempermed GmbH1065920
Eye protection3M™ Health Care71512-00001M
Glass Bead SterilisatorVanPel laboratory equipmentGBS300H
GraphPad PrismGraphPad Software, San Diego, California USA version 10
Heating padConduct ScienceCS-HP-2005
Isoflo (isoflurane)Zoetis / Dechra Veterinary ProductsB506—
Lidocaine 1%Henry Schein809755
Mesh bottom cagesStoelting Co.57816, 57824—
Micro forcepsFine Science Tools11251-35—
Micro needle holderFine Science Tools12076-12—
Micro scissorsFine Science Tools15019-10—
Micro tweezersFine Science Tools11254-20—
NaCl 0.9%TrademedH7 1000-FRE—
Needle 25G × 1"Becton, dickinson & company300400
Needle holderFine Science Tools12004-16—
Nerve stimulator (VARI-STIM Nerve locator)REF 856200
RandoMice softwarevan Eineige et al.  LeidenVersion 1.1.7
Rodent rectal probeWorld Precision InstrumentsNC9506514
ScalpelFine Science Tools10003-12—
ScissorsFine Science Tools14001-12—
Sensory evaluatorLinton Instrumentation514007–5140150.6 g to 15 g monofilaments
Stereo surgical microscopeLeicaA60 F—
Sterile gauze padsNobamed740113
Sterile ocular lubricant ointmentAlcon ‎B00BRLSC8S
Sterile sheet with holeEvercare / OneMed1555-01—
Surgical blade nr. 15Fine Science Tools10015-00—
Surgical face maskMedline Industries, Inc.MSK1002
Syringe luer lock 2 mlBecton, dickinson & company300613
TweezersFine Science Tools11617-12—

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