Authors: Fatima Aldali, Li Tang, Yujie Yang, Yunjie Huang, Yajie Li, Chunchu Deng, Hong Chen
Categories: Review, Peripheral nerve injury, Wallerian degeneration, Mesenchymal stem cell, Growth factors, Nerve guidance conduit, Biophysical stimulation, Regenerative rehabilitation
Source: Journal of Translational Medicine
Authors: Fatima Aldali, Li Tang, Yujie Yang, Yunjie Huang, Yajie Li, Chunchu Deng, Hong Chen
Peripheral nerve injuries (PNIs) remain a major clinical and socioeconomic challenge, frequently resulting in motor weakness, sensory loss, and chronic neuropathic pain that cause long-term disability and restrict daily function. Functional recovery is limited by slow axonal regrowth, Wallerian degeneration, interstitial fibrosis, and progressive denervation-induced muscle atrophy. Although microsurgical epineurial repair and autologous nerve grafting are standard treatments, clinical outcomes remain inconsistent, especially in long-gap or delayed repairs. These limitations underscore the need for more effective regenerative strategies that address both the structural and biological barriers to nerve recovery.
Contemporary research on PNIs focuses on four interconnected structural reconstruction, biological acceleration, functional remodelling, and anatomical restoration. Advanced nerve-guidance conduits offer biomimetic, aligned pathways that reduce axonal misdirection and complement microsuture or autograft repair. Biological approaches, including localized delivery of neurotrophic factors, mesenchymal stem cells, induced-pluripotent stem cell derivatives, and their exosomes, enhance Schwann cell reprogramming, angiogenesis, and pro-regenerative immune polarization while reducing risks associated with live cell transplantation. Non-invasive biophysical stimulation modalities, such as electrical stimulation, magnetic fields, photobiomodulation, low-intensity pulsed ultrasound, and piezoelectric scaffolds, further promote axonal growth and neurotrophic signaling. Emerging integrated strategies that combine stem cell–derived exosomes with physical cues demonstrate synergistic regeneration in preclinical models, representing promising avenues for treating critical-sized nerve gaps. Multi-omics technologies, including transcriptomics, proteomics, metabolomics, and spatial profiling, have deepened mechanistic understanding of Schwann cell plasticity, axon–glia communication, and injury-induced inflammatory dynamics. However, clinical translation remains constrained by heterogeneity in study design, biomaterial manufacturing, regulatory requirements, and the lack of validated biomarkers for monitoring nerve regeneration. Overcoming these obstacles will require coordinated efforts across surgery, biomaterials engineering, stem cell biology, pharmacology, neuromodulation, and rehabilitation medicine.
Recent progress in biomaterial conduits, cell-free biologics, and biophysical stimulation is transforming PNI treatment and providing options that surpass conventional microsurgical repair. Continued advancement will require reliable biomarkers, standardized production and evaluation methods, and well-designed randomized controlled trials. Coordinated collaboration across research, clinical practice, industry, and regulatory agencies is essential to develop safe, effective, and widely applicable neuroregenerative therapies that restore meaningful function after peripheral nerve injury.
Peripheral nerve injuries (PNIs) are a significant clinical challenge, repeatedly causing chronic pain, long-term sensory and motor deficits that greatly lead to long-term disability and ipact patients’ quality of life [1]. While the basic mechanisms of nerve degeneration and regeneration have recently been well understood, effective treatments for complete nerve function are limited. Following injury, peripheral nerves undergo a well-ordered process of degeneration that involves axonal damage, demyelination, and Wallerian degeneration (WD), followed by a regeneration phase that includes activity of Schwann cells (SCs), immune cell activity, and axonal sprouting [1]. Despite advances in microsurgical techniques, the outcomes of nerve repair remain suboptimal, particularly for complex or large-gap injuries.
In recent years, the field has experienced a significant surge of innovative advancements, including biomaterial-based nerve conduits, bioengineered scaffolds, cell-based therapies, electrical stimulation techniques, gene editing technologies, and tissue-engineered constructs [2–5]. Furthermore, cutting-edge advancements in imaging modalities and artificial intelligence are fundamentally altering both diagnostic and therapeutic frameworks [6, 7]. Given these developments, a thorough review is essential to integrate recent breakthroughs across various disciplines, identify translational barriers and opportunities that could enhance clinical implementation, propose a strategic roadmap for future research directions, and foster cross-disciplinary collaboration. Through comprehensive searching in PubMed, Web of Science and ClinicalTrials.gov using combinations of the keywords “peripheral nerve injury” with “surgical treatment”, “pharmacological treatment”, “growth factor”, “stem cell”, “exosome”, “exercise”, “electrical stimulation”, “extracorporeal shock wave therapy”, “low-intensity pulsed ultrasound”, “physical therapy modality”, and “physical stimulation”, we begin by classifying PNIs according to the type and extent of structural damage. Current therapeutic strategies are then examined, encompassing surgical repair techniques, pharmacological interventions, and emerging regenerative approaches such as stem cell and exosome-based therapies. Finally, we highlight rehabilitative modalities—including physical and neuromodulatory interventions—that hold promise for enhancing nerve regeneration and functional recovery.
The peripheral nervous system (PNS) is a complex network of nerves that functionally integrates various body parts with the central nervous system (CNS). It is formed by the fusion of the ventral and dorsal roots, originating from the spinal cord [8]. The ventral root contains motor neurons, whereas the dorsal root houses sensory neurons and the cell bodies are located in the dorsal root ganglion. The motor neuron cell bodies are located in the ventral horn of the spinal cord and specific nuclei of the brainstem. The PNS has a superior regenerating capability than the CNS [9]. The regenerating capability of PNS is affected by its functional environment, the integrity of the injured nerve, and the patient’s age, type, and severity of the injured nerve [10]. A nerve fiber (Fig. 1) is the nerve’s conducting unit, composed of a central core, axons, and SCs. Large nerve fibers (Fig. 1) are wrapped by a myelin component (myelinated nerve fibers), while small ones are with or without a myelin sheath [11]. Axons harbor various organelles, including mitochondria, neurofilaments, endoplasmic reticulum, microtubules, and electron-dense particles [11]. They extend from neuronal cell bodies located in the spinal cord, dorsal root ganglia, or autonomic ganglia, depending on their origin [11]. SCs are PNS glial cells that are located along the axon’s longitudinal expansion. Nerve fibers of various diameters, both large and small, can be found in healthy peripheral nerves. Only large-diameter axons ( > 1.5 μm) are ensheathed in segmental lipoprotein layers, forming myelin. In these cases, adjacent SC membranes wrap concentrically around portions of the axon [12]. The small gap between successive SCs is known as the node of Ranvier [12]. The Ranvier node allows for ionic exchanges between a nerve fiber’s axoplasm and the intercellular space, as well as saltatory conduction of a nerve action potential impulse, which jumps from one node to the next and serves as the foundation for rapid signal conduction.Fig. 1Peripheral nerve structure. This figure illustrates the detailed organization of a peripheral nerve in cross-section. Multiple fascicles—bundles of nerve fibres—are encased by the epineurium, a protective connective tissue sheath. Within each fascicle, individual axons are enveloped by myelinating Schwann cells or supported by non-myelinating counterparts. Each axon is surrounded by the endoneurium, while the perineurium encloses the entire fascicle. The nerve architecture also includes blood vessels, resident macrophages, and fibroblasts, which collectively contribute to homeostasis and facilitate repair processes following injury
Peripheral nerve fibers, the most vulnerable structure in the body, are easily injured by crushing, compression, or trauma. Trauma to the peripheral nerve can cause insufficient interaction between the brain, organs, and muscles [13]. PNIs are a significant public health concern due to their high incidence. PNIs can impair motor activity and sensory loss in certain areas of the body and negatively impact brain function and communication with target organs or muscles. These injuries can have long-term effects on behavior, movement, perception, consciousness, and skin/joint sensations [10]. Treating nerve injuries presents significant challenges due to several factors, including location, severity, and type of damage. These characteristics influence treatment options and the possibility of recovery, and understanding these complexities is critical to determining effective therapy approaches [14].
Trauma of the nerves is the primary cause of peripheral neuropathy. In 1942, Seddon published a classification of nerve injury severity based on his observations of the trauma of the affected nerve (Fig. 2). The classification of PNIs has been developed to provide clinicians and researchers with a systematic framework for understanding the extent of neural damage and guiding treatment options. The foundational classification was categorized nerve injuries into three primary types based on clinical and anatomical Grade I (neurapraxia), a transient conduction block without axonal disruption; Grade II (axonotmesis), where the axon is disrupted but the surrounding connective tissue layers remain intact; and Grade III which represent increasing severity of structural damage. Specifically, in Grade III, both the axons and the endoneurium are injured, while the perineurium is preserved [15]. Sunderland (1951) developed a more detailed classification that distinguishes five degrees of injury severity, correlating with distinct histopathological changes and recovery potential. While grades I and II correspond to Seddon’s neurapraxia and axonotmesis, respectively. The subsequent grades in Sunderland’s system align with progressive damage to the nerve’s internal structures, including the endoneurium, perineurium, and epineurium, offering a more detailed framework to guide prognosis and intervention strategies [16]. The epineurium is a dense, fibrous connective tissue sheath that encloses the entire nerve trunk, providing mechanical protection, structural support, and flexibility to the nerve [16].Fig. 2Classification of peripheral nerve injuries. This figure illustrates progressive structural damage from neuropraxia to neurotmesis across axon and connective tissues
Grade I nerve injuries typically result from compression of peripheral nerves, often due to direct pressure applied to a nerve trunk or nerve root. Nerve compression injuries commonly affect nerves that pass through rigid anatomical structures or traverse bony prominences, and also occur due to prolonged positioning of limbs. Examples include “Saturday night palsy,” characterized by injury to the upper arm following prolonged compression from sleeping with the arm draped over the back of a chair [17]. Comparable injuries can also arise postoperatively when limbs remain immobile in one position for extended durations [18]. These conditions are known as compression neuropathy or nerve compression syndrome.
The pathological mechanisms underlying compression neuropathy typically involve both mechanical compression and subsequent ischemia. Mechanical compression increases intra-epineural pressure, which, when exceeding intra-arterial pressure, leads to ischemic conditions and varying degrees of axonal damage [19]. Clinically, Grade I injuries generally cause transient impairment of motor functions and may also result in neuropathic pain. In addition, complete loss of motor or sensory functions rarely occurs because nerve continuity remains intact.
Grades II, III, and IV in Sunderland’s classification represent more severe forms of PNIs compared to Grade I. Grade II includes axonal disruption with preserved connective tissue structures, while Grade III includes damage to the endoneurium, and Grade IV extends to perineurial disruption, resulting in substantial structural disorganization and impaired regeneration potential.
These injury grades are usually caused by mechanical forces such as crushing or stretching. A crush injury results from sudden compression of the nerve, often due to blunt trauma, surgical clamping, or traffic-related trauma [20]. Although variability in severity and axonal damage is a consistent feature, functional recovery is possible. Pathologically, crush injuries induce WD, a process first described by August Waller and characterized by axonal and myelin sheath degeneration distal to the injury site, followed by clearance of debris by SCs and macrophages [13]. Unlike transection injuries, the connective tissue scaffolds remain intact, thereby facilitating axonal regrowth and enhancing the potential for functional recovery [13]. However, regeneration can be delayed and inconsistent, and the formation of a neuroma—an irregular mass of regenerating nerve fibers and connective tissue at the injury site—may further hinder proper nerve regeneration [21].
The sciatic nerve crush model is widely used in preclinical studies to investigate nerve regeneration. This model is typically induced in rodents using serrated or non-serrated forceps. Due to varying instruments and techniques, there is heterogeneity among studies. Beer et al. [22] introduced a non-serrated clamp to standardize compression, offering reproducible results across rodent species [22]. A more recent instrument, FST toothless forceps—fine surgical forceps manufactured by Fine Science Tools with smooth, non-serrated tips commonly used for delicate tissue handling without causing mechanical damage—is widely employed in experimental research to simulate PNIs in rats due to its reproducibility and relevance to clinical trauma. The limited variability in nerve regeneration between individuals makes this model particularly effective for evaluating new therapies [23]. However, in humans, fibrosis at the lesion site often prevents spontaneous regeneration, necessitating surgical repair or grafting.
Grade V, the most severe form of nerve injury in Sunderland’s classification, comprises a complete nerve transection, resulting in total loss of continuity of the nerve trunk [24]. This type of injury is typically caused by sharp cutting such as knife wounds, broken glass, metal fragments, chainsaws, wood splinters, or animal bites [20]. Accidental transection of spinal nerve branches during surgical procedures is also a potential cause.
In these nerve injury cases, spontaneous functional recovery is often not achievable, although limited partial recovery may occur. The sharpness of the injured object generally influences the clarity of the cut and subsequently the potential for surgical repair. Additionally, the chance of a favorable outcome is higher when the distance between the proximal and distal nerve stumps is minimal.
In preclinical studies, complete sciatic nerve transection is commonly used to investigate severe nerve injuries. While axonal regeneration can occur in laboratory animals following end-to-end neurorrhaphy—the surgical suturing of a severed nerve to restore continuity and promote axonal regrowth—this model replicates human clinical scenarios. In rodents, transection leads to significant loss of motor and sensory function in the affected limb and neuropathic pain. This model is particularly valuable for investigating the consequences of distal denervation, muscle atrophy, and the effectiveness of repair strategies, including suture repair, nerve glues, and anti-fibrotic interventions [25].
Traumatic PNIs are often marked by axonal interruption, resulting in disconnection from the neuronal cell body. This axonal injury leads to a complete loss of nerve conduction, manifesting clinically as muscle weakness, sensory loss, neuropathic pain, and maladaptive responses [26]. The consequences of nerve injury are significant, as functional recovery is frequently incomplete. Fibrotic scarring at the injury site can impede the restoration of limb function. Moreover, severe or proximally located injuries may trigger neuronal death. In contrast, if the injury, such as a transection or crush, is sufficiently distal, regeneration of nerve fibers is possible [27]. Traumatic PNIs stimulate the array of cellular and molecular events of WD in the nerve segments sited distal to the injury site (Fig. 3).Fig. 3Actions following peripheral nerve injury. Illustrations of peripheral nerve injury actions show Wallerian degeneration, immune activation, Schwann cell roles, axon regeneration, or scar tissue formation
Following the injury, the connective tissue components surrounding SCs also influence the regenerative potential after injury. The connective tissue layers of the nerve epineurium and perineurium comprise primarily fibroblasts, wrapping SCs, and their nerve fibers. The integrity of these layers influences the outcome of nerve repair. In crush injuries, the epineurium remains intact, facilitating relatively rapid regeneration. For example, in rodent models of facial nerve crush, significant functional recovery can be observed within days to two weeks [28, 29]. In contrast, nerve transections disrupt all nerve layers, including the axons, myelin, and connective tissue sheaths, resulting in more severe damage. These injuries leave a proximal and distal stump that must be surgically reconnected to permit regeneration, and even then, functional recovery is typically slow and incomplete.
After a PNI, axonal outgrowth is regulated by a variety of factors, including SC transformation, immune cell infiltration, and neurovascular regeneration. Immune cells, such as macrophages, neutrophils, and endothelial cells, along with SCs, play pivotal roles at different stages of nerve repair. In intact peripheral nerves, mature SCs differentiate into either myelinating or non-myelinating forms. However, after PNI Both types contribute to the structural, and functional integrity of the PNS, playing essential roles in neuronal protection, maintenance, and regeneration [30].
Repair and regeneration involve complex interactions between resident and infiltrating cells, including SCs, fibroblasts, and endothelial cells, and infiltrating immune cells like macrophages, neutrophils, and lymphocytes, which collectively coordinate debris clearance, modulate inflammation, secrete growth factors (GFs), and reconstruct the extracellular matrix (ECM) to facilitate axonal regeneration [31]. SCs play a pivotal role in coordinating the interplay between resident and infiltrating immune cells by secreting cytokines and chemokines that recruit circulating immune cells, particularly macrophages. Infiltrating macrophages and other immune cells, in turn, secrete signaling molecules that modulate SC activity. For example, activation of nuclear factor-kappa B (NF-κB) in SCs—potentially driven by T-cell-derived signals and tumor necrosis factor-alpha (TNF-α)—can amplify macrophage accumulation [32]. This reciprocal communication is essential for establishing a regenerative microenvironment conducive to nerve repair [32]. Furthermore, SCs are responsible for the formation of the myelin sheath along peripheral nerves and for the creation of bands of Büngner—longitudinally aligned columns formed by proliferating SCs within the basal lamina tubes of degenerated axons following WD—which are essential for guiding regenerating axons toward their targets [33, 34]. Extensively, the SC response is guided by intrinsic cellular reprogramming triggered by external signals from degenerating axons, immune cells, and the extracellular matrix. Dedifferentiated SCs begin to migrate from both proximal and distal nerve stumps to form a cellular bridge that spans the injury gap. This bridge not only facilitates axonal guidance but also supports neovascularization, which supplies essential nutrients and guides SC, and macrophage migration, creating a permissive microenvironment for axonal regrowth. Furthermore, SCs secrete growth-promoting factors and expression of regeneration-associated genes and secretion of regeneration-associated genes [35].
Vascular endothelial growth factor (VEGF) is a key regulator of orchestrating angiogenesis and peripheral nerve regeneration, particularly through its interaction with SCs during WD. Local hypoxia and inflammation-induced tissue necrosis following PNIs impede nerve repair and regeneration [36]. In response to hypoxic conditions, macrophages promote angiogenesis to restore nutrient supply and support SC migration and proliferation, thereby facilitating the clearance of necrotic debris [36]. In addition, VEGF expression after PNI is upregulated and contributes to both neovascularization and SC-guided axonal regeneration [37]. VEGF supports the formation of blood vessels that act as structural tracks for migrating SCs, facilitating the proper alignment of SCs along the degenerating nerve and the formation of Bands of Büngner structures vital for axon guidance and remyelination [35]. VEGF signals through VEGF receptor1-3 (VEGFR-1- 3), and co-receptors neuropilin-1 and neuropilin-2 to regulate key downstream pathways that promote SC survival, migration, and regenerative capacity [36].
Macrophages play a central role in the repair process following PNIs by clearing debris, modulating inflammation, and influencing SC behavior. Macrophages are initially recruited to the injury site by SC-derived signals, including monocyte chemoattractant protein-1 and leukemia inhibitory factor. They then rapidly infiltrate the damaged nerve and play a critical role in facilitating WD [31, 38]. Furthermore, polarization toward an anti-inflammatory M2 phenotype promotes SCs bridge formation and enhances axonal regeneration [31]. Fibroblast growth factor (FGF)9 has been shown to play a pivotal role in SC maturation, myelination, and the regulation of inflammatory responses after nerve injury. Studies indicated that the expression of FGF9 in SCs impacts macrophage behavior; its downregulation post-injury correlates with the upregulation of proinflammatory cytokines and delayed macrophage infiltration [39]. Macrophage-derived microvesicles specifically from M2 macrophages stimulate SC proliferation and migration via to facilitate nerve repair [40]. Additionally, macrophage-secreted Slit-3 signals through SC-expressed Robo-1 to regulate SC migration and coordination with fibroblasts during regeneration [41]. Furthermore, macrophages indirectly influence the expression of trophic factors and neurotrophins such as glial cell line-derived neurotrophic factor (GDNF), nerve growth factor (NGF), and brain-derived neurotrophic factor (BDNF), which support SC and axonal regeneration [42, 43].
Fibroblasts play an essential role in PNIs and regeneration, particularly in association with SCs. Following transection injury, fibroblasts assist in early debris clearance and release ECM components promoting SC migration and regeneration [44, 45]. Their involvement is vital in traumas when nerve sheaths become damaged, allowing fibroblasts to infiltrate and interact with SCs. Fibroblasts upregulate tenascin-C (TNC) following injury, which promotes SCs migration via β1 integrin binding and promotes axonal regeneration [45]. Furthermore, fibroblasts stimulate tissue vascularization by expressing cytokines and trophic factors [46].
The ECM is vital in guiding SC behavior and axonal regeneration during WD following PNI. The ECM is composed of structural proteins such as collagen, elastin, fibronectin, laminin, glycoproteins, proteoglycans, and glycosaminoglycans [47]. ECM provides both mechanical support and signaling cues essential for SC migration, proliferation, differentiation, and morphogenesis [48]. Furthermore, Type VI collagens not only contribute to myelin sheath formation but also modulate macrophage function, thereby promoting regeneration [49]. Fibronectin, primarily secreted by SCs, supports SC migration and axon growth through interactions with α5β1 integrin and is upregulated following nerve injury [50, 51]. Incorporation of fibronectin into alginate hydrogel nerve conduits has been shown to significantly enhance peripheral nerve regeneration by promoting SC viability and enhancing their supportive role in axon growth, highlighting the essential contribution of fibronectin to the regenerative microenvironment during nerve conduit transplantation [52]. Laminin is a key basement membrane glycoprotein composed of α, β, and γ chains that interacts with collagen IV and cell surface receptors to regulate cell activities [53]. In nerve regeneration models, cross-linked laminin enhances angiogenesis and promotes repair by promoting capillary formation around injured peripheral nerves [54]. Thus orchestrates the structural and molecular events essential for effective peripheral nerve regeneration.
Although rodent models have significantly contributed to the interpretation of PNI and nerve regeneration; nevertheless, their anatomical and physiological variations from humans limit their practical applicability. Rodents present shorter axonal pathways and a microenvironment that is more advantageous to regeneration, as follows, enabling prompt restoration across gaps that may range from millimeters to centimeters [55]. Contrariwise, human nerves often necessitate regeneration across considerably greater distances, frequently surpassing 30 cm, which induces the development of fibrotic scars [55, 56]. Furthermore, distal SCs undergo degeneration due to the lack of contact with proximal neurons, which results in reduced expression of neurotrophic GFs, changes in the ECM and loss of SC basal lamina, all of which obstruct axonal extension [55]. Consequently, more studies indicate that larger animal models, such as sheep or guinea pigs, may produce more clinically relevant insights into nerve regeneration due to their anatomical similarities, fascicular organization, and electrophysiological characteristics that closely resemble those of humans [57, 58].
A comprehensive literature search was conducted across PubMed, Web of Science, and ClinicalTrials.gov using a combination of the keyword “peripheral nerve injury” with relevant terms including “surgical treatment”, “pharmacological treatment”, “growth factor”, “stem cell”, “exosome”, “exercise”, “electrical stimulation”, “extracorporeal shock wave therapy”, “low-intensity pulsed ultrasound”, “physical therapy modality”, and “physical stimulation”. Studies were considered for inclusion if they specifically addressed PNI and its associated therapeutic interventions.
A diverse range of therapeutic strategies have been developed in clinical and preclinical settings, including surgical interventions, physical therapies, pharmacological treatments, nanotechnology-based approaches, neurotrophic factors, stem cell therapies, and extracellular vesicles (Fig. 4).Fig. 4Current peripheral nerve injury treatments
Traumatic PNIs often require surgical intervention to restore sensory and motor function and prevent long-term disability. However, effective surgical management depends on factors (i.e., type of injury, the gap between nerve stumps, and the surrounding tissue environment). Here, we outlined the key suturing techniques and grafting strategies used in clinical practice.
Microsurgical nerve suturing remains the main strategy of peripheral nerve repair, particularly for short-gap PNIs ( < 1 cm). This approach entails suturing the proximal and distal ends of the severed nerve to restore continuity, while minimizing tissue trauma and ensuring precise fascicular alignment. A fundamental principle is to use as few sutures as needed to avoid unnecessary foreign body reaction and tension, both of which may impair axonal regeneration [59]. High-quality suture materials and magnification, either through surgical loupes or a microscope, are essential to achieve precision [60]
Three main suturing techniques are epineurial, fascicular, and epi-perineurial suturing.
Epineurial suturing is the most commonly used method, involving end-to-end approximation of the nerve stumps by passing sutures through the outer epineurium. This technique minimizes intrusion into the nerve fascicles, requires fewer sutures, and is technically less demanding. However, a potential drawback is a misdirection of regenerating nerve fiber due to a lack of fascicular nerve repair [60].
Fascicular Although this method improves specificity and reduces the chance of misdirection of nerve fibers, it is more technically challenging, increases surgical time, and may result in greater tissue reaction at the repair site [60]. A variation, group fascicular repair, is particularly useful in nerves with many fascicles, such as the median nerve, as it reduces the complexity of the full fascicular matching [61].
Epi-perineurial suturing is a hybrid approach in rare cases, where needles pass through both the epineurium and the perineurium of adjacent fascicular groups. This less commonly used method provides enhanced fascicular alignment and may improve regenerative outcomes in severe injuries [62].
For repairing extensive nerve gaps ( > 3 cm), severe nerve injuries, and more proximal sites, the autograft remains the gold standard since it provides both structural scaffolding and trophic support for axonal growth. This approach involves harvesting a segment of nerve tissue from functionally less critical nerves, including the sural, superficial cutaneous, or lateral femoral cutaneous nerves [63]. The selection of the optimal donor nerve relies on various factors, including the length of the nerve gap, the site of repair, and the potential morbidity at the donor site. Autologous nerve grafts provide an immunologically compatible scaffold that facilitates nerve regeneration by offering adhesion molecules, neurotrophic factors, and a supportive microenvironment [64]. However, this technique has several limitations. These include donor site morbidity, the requirement for additional surgery, functional loss at the harvest site, scarring, neuroma formation, and the limited availability of suitable grafts. Furthermore, challenges such as fascicle mismatch and the requirement for a second incision can complicate outcomes [65].
To overcome the limitations related to nerve autografts, several alternative strategies have been investigated, including nerve allografts, biological conduits, and synthetic conduits. This is especially in cases of nerve gap exceeding 3 cm in humans, and when the available autologous donor nerve is insufficient, allografts offer a viable clinical alternative. Nerve allografts provide structural guidance and may preserve viable SCs, which support axonal regeneration and facilitate target tissue reinnervation [10]. However, one of the disadvantages of this technique is the requirement for long-term immunosuppression, which lasts 18 to 24 months post-implantation to prevent immune rejection and allows adequate integration of the graft. This prolonged immunosuppression increases the patient’s susceptibility to the risk of infections and systemic complications, thereby limiting the clinical application of nerve allografts [64]. However, concerns related to immunogenicity and inconsistent regenerative outcomes have driven the development of artificial nerve guidance conduits (NGCs). These engineered constructs aim to promote nerve regeneration while eliminating the necessity for immunosuppressive therapy.
NGCs have increasingly become integral to the field of peripheral nerve repair, particularly for injuries where direct suturing is not feasible and autografts establish many limitations. These tubular scaffolds are designed to bridge nerve gaps by providing a physical and biochemical environment that supports axonal regrowth and SC migration. Over the years, multiple materials and configurations have been explored, some of which have been translated into clinical use, and others remain in experimental or preclinical development. Biomaterials are utilized to support the regeneration of peripheral nerves. A variety of biomaterials have been used to achieve this [66]. Among the conduits approved for clinical use, collagen-based and polyglycolic acid-based materials are the most prevalent [67]. Collagen, a primary component of ECM, offers excellent biocompatibility, biodegradability, and the ability to support SC adhesion and axonal guidance [68]. Currently, five commercially available FDA-approved collagen type I nerve conduits are in clinical NeuraGen®, NeuroMatrix®, NeuroFlex®, NeuraWrap®, and NeuroMend®. These conduits are primarily composed of type I collagen. The degradation time varies among products, with most conduits resorbing within 4–8 months (NeuroMatrix®, NeuroFlex®), while some, such as NeuraWrap®, may persist up to 36–48 months. The degradation time of NeuraGen® could be 3 to 4 years. This variability raises concerns that degradation may occur before completion of the nerve regeneration process, particularly in the context of larger nerve defects [69]. In contrast, non-biodegradable conduits—such as those made from expanded polytetrafluoroethylene (ePTFE, e.g., Gore-Tex®) or silicone tubes—remain permanently in the body, offering long-term structural support but with the potential for chronic foreign-body reactions, fibrous encapsulation, and limited integration with regenerating tissue [70]. The choice between bioabsorbable and non-biodegradable conduits depends on the nerve gap length, required mechanical support, and desired long-term tissue remodeling outcomes.
Current pharmacological practice for PNIs can be divided into three groups. The first group comprises established analgesics that target nociceptive and neuropathic pain without directly influencing axonal repair, which are used immediately after trauma or surgery, non-steroidal anti-inflammatory drugs, or acetaminophen as first-line; these both reduce prostaglandin-driven peripheral sensitisation and reduce postsurgical discomfort [71]. The corticosteroid mimicking adrenal hormones like cortisol, alleviates PNI-related inflammation and pain. Their pharmacokinetics are complex, typically requiring local injections to reduce systemic risks while efficiently treating PNIs, while lidocaine blocks ion channelsin peripheral nerves, permanent focal anaesthetic is treated topically with lidocaine patch, which has strong analgesic and anti-inflammatory characteristics, whereas widespread burning or electric shock pain is treated with gabapentin or pregabalin, both of which are now considered first-line treatments and can relieve neuropathic pain by binding to voltage-gated Ca^2+^ channels [71]. Intravenous (IV) administration of ultra-high-dose methylcobalamin (MeCbl), an analogue of vitamin B12, was evaluated in a small open clinical trial involving fourteen patients with immune-mediated or hereditary neuropathy in the chronic progressive or stable phase. In this trial (UMIN000009359), patients received 25 mg/day MeCbl for 10 days followed by monthly infusions for five months. The treatment was well tolerated in most participants, with no serious adverse effects reported, and over half of the evaluated patients exhibited improved muscle strength in the Medical Research Council sum score in at least two of the 20 assessed muscles. These findings indicate that MeCbl therapy is both safe and potentially effective in enhancing motor function in chronic peripheral neuropathy, highlighting its promise as a disease-modifying approach to support axonal regeneration in clinical settings [72, 73]. Local delivery of tacrolimus (FK506), which binds FK506-binding protein 12 (FKBP12) to modulate calcineurin and other downstream pathways, via biodegradable wraps consistently increases myelinated-axon counts and myelin thickness in rodents [74, 75]. However, this regeneration candidate is still in the experimental stage.
NGFs) are endogenous molecules released during the injury response that actively support nerve regeneration. Consequently, replicating their release presents a promising strategy to enhance neuronal survival, differentiation, and axonal growth [76]. Therefore, incorporating these growth factors (GFs) represents an additional therapeutic strategy to optimize the microenvironment within guidance conduits, thereby enhancing their capacity to support axonal regeneration [76, 77]. Various types of GFs play important roles in peripheral nerve regeneration, each contributing through distinct mechanisms such as promoting neuronal survival, guiding axonal elongation, and supporting SC function (Fig. 5). These include classical neurotrophins like NGF, BDNF, neurotrophin-3 (NT-3), and neurotrophin-4, as well as neuropoietic cytokines such as ciliary neurotrophic factor (CNTF), and GDNF, which are critical for motor neuron survival and axon guidance [76, 78].Fig. 5Key neurotrophic and angiogenic factors that contribute to peripheral nerve regeneration following injury. Each factor exerts specific Glial cell line-derived neurotrophic factor (GDNF); Brain-derived neurotrophic factor (BDNF); Insulin-like growth factor-1 (IGF-1); Ciliary neurotrophic factor (CNTF); Fibroblast growth factor-2 (FGF-2); Vascular endothelial growth factor (VEGF) improves vascularization and axonal growth; and Nerve growth factor (NGF)
In the section that follows, we review the most commonly studied GFs in the context of PNIs and highlight their therapeutic relevance and limitations. NGF is the most extensively studied among these GFs, as it is naturally expressed in healthy nerves, upregulated following nerve injury, and plays a crucial role in neuronal growth and survival [78]. The physiological effects exerted by GFs are mediated through their interaction with specific receptors expressed on the surface of neuronal cells. Their principal application lies in promoting axonal elongation and facilitating SC migration [78].
NGF promotes neuronal survival, axonal elongation, and SC migration by binding to its high-affinity receptor Tropomyosin receptor kinase A (TrkA) and low-affinity receptor p75^NTR [78]. These interactions activate intracellular signaling cascades such as PI3K/Akt and MAPK/ERK, which are essential for nerve repair [79]. While NGF enhances functional recovery, its overexpression may lead to neuropathic pain, emphasizing the need for precise therapeutic regulation [80].
BDNF, a key neurotrophin, plays critical roles in learning and memory, hippocampal neurogenesis, synaptic plasticity, and nerve regeneration following injury. In peripheral nerves, BDNF is produced by SCs, motor neurons, and a subset of dorsal root ganglion (DRG) neurons. Notably, after nerve compression or complete transection, BDNF mRNA levels peak in all three cell types [81]. However, BDNF’s effects are primarily restricted to specific neuronal subpopulations, including sensory DRG neurons, where it promotes neurite outgrowth. BDNF stimulates the production of pro-regenerative cytokines via the activation of the Janus kinase/signal transducer and activator of transcription (JAK/STAT) pathway in SCs rather than neurons, as previously thought [82]. Furthermore, GDNF has also been utilized to target sensory neurons to alleviate pain in cases of chronic denervation [83]. BDNF-based therapies for PNIs face challenges such as short half-life, uncontrolled diffusion, dose-dependent pain sensitization, and receptor downregulation, limiting therapeutic efficacy and clinical translation Controlled, targeted delivery systems are essential to use BDNF’s regenerative potential without adverse effects [84].
GDNF has also been employed to target sensory neurons in order to alleviate pain associated with chronic denervation [85]. Given the challenges in achieving optimal dosing of GDNF, recent strategies for its application in PNI have primarily focused on cell-based or gene therapy approaches [86]. In this context, Shakhbazau, Mohanty et al. [86] conducted a proof-of-concept study utilizing SCs genetically engineered via lentiviral transduction to express GDNF under the control of a doxycycline-inducible system [86]. Upon transplantation of these GDNF-modified SCs into transected peripheral nerves and subsequent time-restricted doxycycline administration, the study demonstrated that GDNF expression could be precisely regulated and monitored within the transplanted cells [86]. Furthermore, GDNF supports nerve repair by promoting motor neuron survival, enhancing axonal regeneration, and activating SC. Its regenerative effects were demonstrated in sensory nerve repair models, where exogenous GDNF improved corneal nerve regeneration and functional recovery [87]. Subconjunctival injection of NGF or GDNF neutralizing antibodies significantly impaired corneal repair. Specifically, blocking NGF or GDNF delayed corneal epithelial wound healing and reduced corneal nerve regeneration, with subbasal nerve fiber density decreased by 59% and 51%, respectively [87].
Beyond classical neurotrophins, VEGF is crucial for peripheral nerve repair. Given the particular structural and functional relationship between vascular and nervous systems [88]. VEGF not only promotes angiogenesis but also exhibits neurotrophic in the survival and the physiology of motoneurons [89]. VEGF enhances sciatic nerve recovery through improved vascularization and contributes to axonal growth [90]. Notably, VEGF also shows benefits and holds promise as a therapeutic agent that could promote the proliferation, migration, and differentiation of SC [91]. Hepatocyte growth factor (HGF) is another promising factor in nerve regeneration. Following PNIs, HGF is upregulated at both proximal and distal sites [92]. Exogenous HGF enhances axon diameter and myelin thickness, whereas blocking c-Met reverses these effects. Gene therapy using HGF-bearing plasmids significantly improved axon density and functional recovery, rivaling autografts [93]. At 14 days post-injury, HGF gene therapy significantly reduced the compound action potential (CAP) latency compared to the untreated control [93]. The CAP latency for the pC4W-hHGF200 group was 880.0 ± 141.84 μs and for the pC4W-hHGF100 group was 1032.0 ± 115.53 μs [93]. Both HGF-treated groups demonstrated a significantly shorter latency than the Control group, which had a value of 1511.43 ± 125.27 μs [93]. Both the HGF100 and HGF200 groups showed a statistically significant increase in CAP amplitude compared to the Control group [93]. The CAP amplitude was 0.10 ± 0.01 mV for both HGF100 and HGF200 groups [93]. The Peak-To-Peak amplitude was 0.22 ± 0.03 mV for the HGF100 group and 0.21 ± 0.03 mV for the HGF200 group [93]. Despite this improvement, the CAP characteristics of the HGF-treated nerves did not fully recover to the level of the Intact group [93]. Beyond its regenerative effects, HGF exhibits anti-inflammatory, anti-apoptotic, and angiogenic properties, beneficial to multiple neural populations, including motor sensory neurons and parasympathetic neurons [94].
Insulin-like Growth Factor 1 (IGF-1) is vital for peripheral nerve repair by preventing SC apoptosis, promoting neurite outgrowth, and preventing muscle atrophy after denervation. It also supports SC proliferation and myelination, thereby facilitating axonal regeneration. Studies have shown that local delivery of IGF-1 accelerates functional recovery in sciatic nerve injury models, making it a strong candidate for therapeutic intervention in PNIs [95]. Based on the study of sciatic nerve crush in neonatal rats, the major findings are that heparin treatment significantly enhanced muscle reinnervation and recovery compared to saline-treated controls [96]. Specifically, heparin increased the extent of reinnervation from 61.1% to 89.7%, restored nerve-evoked muscle twitch tension, and increased extensor digitorum longus muscle weight from 12.0 ± 0.8 mg in untreated denervated muscles to 20.8 ± 0.6 mg, a value comparable to the 21.6 ± 0.4 mg observed in undenervated controls [96]. This recovery was associated with a marked increase in muscle IGF-1 levels [96]. The concomitant administration of anti-growth hormone releasing hormone partially abolished these beneficial effects, reducing the extent of reinnervation to 77.8% and the muscle weight to 16.1 ± 0.7 mg, indicating that the mechanism of heparin’s action is partially dependent on the growth hormone releasing hormone pathway [96].
FGF-2 is essential for promoting peripheral nerve regeneration. It stimulates proliferation, supports angiogenesis, and enhances axonal elongation [97]. FGF-2 may exert neuroprotective effects in part by suppressing autophagy, a process potentially mediated through activation of the PI3K/Akt signaling pathway [97]. FGF-2 delivery has been shown to significantly improve both morphological recovery and restoration of function after nerve injury, especially when used in combination with biomaterials or other GFs [98].
CNTF supports axonal regeneration and facilitates remyelination following nerve injury [99]. It enhances neuronal survival, facilitates SC proliferation, and contributes to the formation of a regenerative environment [100]. Despite its potent effects, CNTF has a short half-life in vivo, making its clinical translation dependent on sustained-release systems or gene-based approaches [101]. However, owing to their broad biological activity, the therapeutic application of GFs remains challenging, necessitating the delivery of highly controlled, low-dose regimens. Additional limitations include their short biological half-life and pleiotropic effects, which can complicate targeted treatment outcomes [76]. These limitations underscore the need for advanced therapies. In this context, stem cell-based strategies emerge as a promising solution, offering sustained neurotrophic support and structural integration to enhance nerve regeneration.
As growth factor-based therapies continue to show promise, Mesenchymal stem cells (MSCs) therapy emerges as the next transformative approach in peripheral nerve regeneration. Stem cells not only act as biological factories producing neurotrophic factors but also provide a structural and cellular platform that integrates with host tissue to guide axonal growth, support remyelination, and enhance long-term functional recovery. This dual function positions stem cells as a powerful complement to molecular therapies in nerve repair.
In addition to recombinant GFs, small-molecule agonists that target neurotrophic-factor receptors have emerged as promising pro-regenerative agents in both the peripheral and CNS. Several receptor-directed classes—including tropomyosin receptor kinase B (TrkB), TrkA, and RET/GDNF-pathway agonists—have demonstrated the ability to promote neurite outgrowth, enhance axon regeneration, and improve functional recovery in preclinical models [102–106]. Small-molecule TrkB agonists such as 7,8-dihydroxyflavone (7,8-DHF) and deoxygedunin induce TrkB phosphorylation, activate pro-survival signaling cascades, and robustly enhance axon regeneration and reinnervation in PNI models [103]. Similarly, LM22A-4 and related BDNF mimetics have been widely used to probe TrkB-mediated repair, showing activity across multiple neuronal models where they promote neuronal survival and differentiation [102]. TrkA-directed agonists have also been described. Gambogic amide, identified by Jang and colleagues, selectively binds to the cytoplasmic domain of TrkA, inducing receptor dimerization and activation of downstream Akt/MAPK signaling [104]. Functionally, it mimics NGF by driving neurite outgrowth and protecting neurons from cell death in vitro, while in vivo administration reduces neuronal loss and tissue damage in models of stroke and excitotoxicity [104]. These findings highlight its utility both as a research probe for NGF/TrkA biology and as a potential therapeutic candidate for neurodegenerative and ischemic injury contexts. Beyond Trk-family receptors, RET agonists such as BT13 and BT18 activate GDNF-family signaling, promoting neurite growth in sensory neurons and attenuating experimental neuropathy in rodents—indicating translational potential for sensory nerve regeneration and neuropathic pain management [105]. These small molecules offer key pharmacokinetic and practical advantages over recombinant proteins, including improved tissue penetration, oral bioavailability, and reduced cost, while avoiding the intrinsic limitations of GFs such as short half-life and manufacturing complexity [102–106]. Nonetheless, significant caveats partial or biased agonism, off-target effects, and receptor-specific adverse outcomes (e.g., pain syndromes linked to NGF/TrkA activation), as well as unresolved challenges in translational dosing and delivery [102, 107108]. Together, these insights underscore both the promise and the complexity of small-molecule neurotrophic mimetics as next-generation therapeutics for nerve regeneration.
MSCs are multipotent cells sourced from various tissues, including adipose tissue, bone marrow, umbilical cord, placenta, dental pulp, and amniotic fluid [109]. Due to their ability to differentiate into diverse cell types, such as neurons and glial cells, MSCs have gained significant attention in regenerative medicine and nerve repair [110] (Fig. 6). Additionally, MSCs possess immunomodulatory properties that create a favorable microenvironment for tissue regeneration [111]. MSC therapies for PNI utilize various delivery methods, each with distinct advantages and limitations. IV injection allows for systemic distribution, potentially reaching distant injury sites and minimizing surgical risks. However, the low concentration of MSCs at the injury site may limit therapeutic efficacy [110]. Local injections, such as epineural or subepineural delivery, provide targeted MSC administration directly at the injury site, enhancing localized therapeutic effects [112]. Yet, they carry risks of surgical complications and may not be feasible for extensive nerve injuries [113]. Intramuscular injection of MSCs provides a minimally invasive method for delivering cells to muscle tissue adjacent to nerve injuries. This approach can support nerve regeneration through paracrine signaling, in which MSCs secrete bioactive factors that promote tissue repair and reduce inflammation [114]. However, the distance from the nerve injury site may limit the direct impact on nerve repair, as the efficacy of paracrine signaling diminishes with increased distance from the source [114]. This limitation underscores the need for optimizing delivery methods to enhance the therapeutic potential of MSCs in peripheral nerve injuries [114]. Conduit-based delivery using biodegradable NGCs provides a scaffold for MSCs and SCs, promoting axonal growth across nerve gaps, especially for long defects [115–117]. NGCs support cell survival and axonal extension, but fabrication requires careful selection of biomaterials and manufacturing techniques to ensure biocompatibility, and surgical implantation carries risks such as infection or tissue injury [115–117]. Despite these challenges, NGCs are a promising strategy for peripheral nerve repair, with ongoing studies optimizing design and functionality [115–117]. ach delivery method should be selected based on the specific clinical scenario, considering factors like injury type, location, and the desired therapeutic outcomes.Fig. 6The sources of Mesenchymal stem cells (MSCs) and their role in PNI
Several methods have been explored for MSC delivery in PNI, including IV, intramuscular (IM), epineural or sub-epineural injections, and cell-loaded nerve conduits [118]. IV administration, while effective, faces limitations due to cell entrapment in capillaries, reducing the number of cells reaching the injury site [119]. Studies demonstrate that IM injections of bone marrow-derived MSCs (BMSCs) significantly enhance functional recovery and nerve conduction compared to IV methods [120]. Combining MSCs with immunomodulatory treatments further enhances functional outcomes in facial nerve injury models [121]. These immunomodulatory factors, such as cytokine inhibitors or tumor necrosis factor, further enhance the efficacy of MSCs; these effects mediate the local inflammatory response after PNI and would reduce loss of muscle mass and change the environment towards a pro-regenerative rather than profibrotic phenotype, leading to better functional recovery [122].
Considering the limitations of autologous nerve grafts, MSC-loaded conduits have emerged as promising alternatives. Chitosan nerve conduits loaded with BMSCs-derived SCs exhibit regenerative capabilities comparable to sciatic nerve-derived SCs [123]. Based on a study bridging a 12 mm sciatic nerve defect in rats, nerves repaired with chitosan conduits seeded with BMSC-derived SCs achieved a conduction velocity recovery (%CVR) of approximately 40%, which was significantly higher than the 30% CVR in PBS-filled conduits (p < 0.05) and statistically similar to conduits seeded with sciatic nerve-derived SCs, though inferior to autografts, which achieved near 60% CVR [123]. Furthermore, muscles receiving input from nerves bridged with SC-seeded conduits showed a muscle girth recovery (% MR) of approximately 75%, a result similar to the ~80% MR achieved with autografts and significantly higher than the ~55% MR observed with PBS-bridged nerves (p < 0.001) [123]. These results, consistent across functional (%CVR) and morphological measures, confirm that BMSC-derived SCs in chitosan conduits significantly enhance nerve regeneration and functional recovery, supporting their use as an effective alternative to autografts [123]. Furthermore, chitosan-based nerve grafts incorporating both human-derived BMSC-ECM (hBMSC-ECM) and microchannels significantly enhance peripheral nerve regeneration, achieving repair effects comparable to autologous nerve grafts [124]. Similarly, adipose-derived stem cells (ADSCs) embedded in conduits effectively promote nerve regeneration and functional recovery, as shown by Shen et al. using undifferentiated human ADSCs in polycaprolactone conduits to repair 6 mm sciatic nerve defects in rats, showing efficacy in preventing muscle atrophy and regaining innervation [125]. Recent advances include using three-dimensional (3D) bioengineered conduits containing human-induced pluripotent stem cell-derived MSCs (iMSCs) for bridging 5-mm sciatic nerve gaps in immunodeficient rats, showing superior nerve regeneration compared to silicone tubes, with improved morphology, function, and gene expression of neurotrophic and angiogenic factors, along with induced angiogenesis compared to conventional methods [126]. In detail, electrophysiologically, the Bio 3D group exhibited a significantly greater compound muscle action potential (CMAP) amplitude (19.07 ± 7.88% vs. 3.15 ± 0.66%; p < 0.05) and a higher, though not statistically significant, mean motor nerve conduction velocity (58.52 ± 12.96% vs. 39.23 ± 17.77%) [126]. Structurally, the Bio 3D group demonstrated superior axonal regeneration with a significantly larger number of myelinated axons (14,837 ± 4,094 vs. 5824 ± 1,255; p < 0.01), larger axon diameter (4.241 ± 0.502 μm vs. 2.064 ± 0.173 μm; p < 0.01), and thicker myelin sheaths (0.861 ± 0.114 μm vs. 0.367 ± 0.036 μm; p < 0.01), resulting in a significantly healthier G-ratio (0.591 ± 0.017 vs. 0.640 ± 0.020; p < 0.01) [126]. Furthermore, the prevention of muscle atrophy was significantly greater in the Bio 3D group, as evidenced by a higher wet weight of the tibialis anterior muscle (58.77 ± 4.44% vs. 31.65 ± 4.87%; p < 0.01) [126].
MSCs contribute to nerve repair primarily through differentiation into SC-like cells [127], which play critical roles in nerve regeneration by producing GFs, clearing cellular debris, and supporting axonal growth. Differentiated MSCs display SC characteristics, including positive staining for SC markers glial fibrillary acidic protein (GFAP), S100 calcium-binding protein (S100), and nerve growth factor receptor (NGFR) [127]. Research indicates that when MSCs are directed toward SC lineage through specific differentiation protocols, they mimic the functional and molecular properties of SCs, including the ability to promote nerve repair and regeneration in various animal models of PNIs [127, 128]. MSC differentiation into SC–like phenotypes has been achieved from multiple tissue sources, including bone marrow, adipose tissue, and amniotic membranes, using stepwise induction protocols that mimic peripheral glial signaling environments. Typically, MSCs are first exposed to β-mercaptoethanol or all-trans retinoic acid to induce a neural progenitor-like state, followed by treatment with glial growth factors such as bFGF, PDGF, forskolin, and heregulin-β1 (neuregulin-1) to promote Schwann cell differentiation [129, 130]. BMSCs treated with this sequential induction acquire a bipolar morphology and express canonical SC markers including S100, p75^NTR^, and GFAP, exhibiting functional properties such as myelin protein expression and neurotrophic factor secretion [129, 130]. ADSCs respond similarly, showing efficient differentiation and higher proliferation potential, making them an accessible source for autologous applications [131]. Amniotic MSCs can also be guided toward Schwann-like cells via exposure to forskolin and neuregulin-1, supporting peripheral nerve regeneration in vivo [132]. Despite these advances, the stability of the Schwann-like phenotype and in vivo integration efficiency remain under investigation. Additionally, molecular mechanisms underlying MSCs involve microRNAs (miRNAs), which are crucial in regulating SC behavior. Human amniotic mesenchymal stem cells (hAMSCs) differentiated into SC-like cells, which, when transplanted into a sciatic nerve injury model, promoted functional recovery by secreting neurotrophic factors. Furthermore, SC-like cells combined with a NT-3 chitosan conduit enhanced nerve recovery via the Sex-determining Region Y-Box 2/fibronectin 1 (SOX2/FN1) axis, which boosted SC-like cell proliferation and migration. SOX2 overexpression increased SC viability and FN1 expression, promoting nerve regeneration [133].
MSCs also exert beneficial effects through paracrine secretion, modulating inflammation and immune responses at injury sites. They lower the levels of pro-inflammatory cytokines including interferon gamma (IFN-γ), Interleukin 1 (IL-1), TNF-α, and IL-6, while enhancing the production of anti-inflammatory cytokines like IL-10, IL-4, and transforming growth factor beta (TGF-β) [122]. MSC-mediated immunomodulation promotes type 2 helper T cell responses and facilitates regulatory T-cell activity, crucial for tissue repair [134]. Furthermore, MSCs influence macrophage polarization toward an M2 anti-inflammatory phenotype, promoting regeneration and reducing scar formation [135]. MSCs also secrete neurotrophic factors, including NGF, BDNF, and GDNF, directly stimulating axonal regeneration and neuronal survival [136]. Recently, researchers have underscored the importance of miRNAs in MSC-mediated nerve regeneration by regulating gene expression, which enhances neuroprotection, myelination, and functional recovery. For example, miR-202-3p, modulated by BMSCs, suppresses neuronal apoptosis after spinal cord injury by activating autophagy through the AMPK, MAPK, and PI3K/AKT/mTOR pathways [137]. Additionally, the downregulation of miR-206 in human umbilical cord MSCs enhances anti-aging effects, suggesting its potential for neuroprotection [138]. Furthermore, SC-like cell derived from hAMSCs enhance sciatic nerve repair by promoting myelination and functional recovery. miR-214 targets c-Jun, a negative regulator of myelination, to improve therapeutic outcomes. The results underscore the efficacy of miR-214 in promoting nerve regeneration and facilitating myelination processes, offering a promising strategy for PNI treatment [139]. These findings underscore the significant role of miRNAs in MSC-based therapies for nerve regeneration, underlining their potential to improve outcomes in nerve regeneration and other neurological disorders. miRNAs not only modulate cellular processes like apoptosis and autophagy but also support neuronal differentiation, synaptic integration, and tissue repair, making them key players in enhancing the therapeutic potential of MSCs for nerve regeneration.
Biomaterial scaffolds play a vital role in supporting MSC therapy for peripheral nerve regeneration with a protected microenvironment, protecting them from surrounding pathological changes and bridge spinal cord stumps, offering physical and directional support for axonal regeneration [140]. Furthermore, biomaterial scaffolds mimic the ECM, helping to suppress immune responses and promote tissue healing. These scaffolds also prevent host-cell infiltration at injury sites, promoting a favorable environment for nerve repair [140]. A diverse range of natural and synthetic scaffolds has been investigated for peripheral nerve and tissue repair, each with unique chemical properties, biodegradation profiles, and translational potential (Table 1). Chitin is a natural β-(1→4)-linked N-acetyl-d-glucosamine polysaccharide with low immunogenicity but limited solubility and slow degradation in vivo [141, 154]. Modified chitin conduits have shown promise in rodent models, particularly when combined with autogenous nerve grafts or growth factor-releasing hydrogels [143, 145]. Chitosan, its partially deacetylated derivative, is cationic, more soluble, and degradable via lysozyme and hydrolysis, with degradation rate dependent on molecular weight and crosslinking [153, 154]. Preclinical studies consistently demonstrate enhanced axonal regeneration and functional recovery with chitosan conduits, especially when combined with SCs, stem cells, or growth factor delivery [148, 197, 198]. Clinically, chitosan-based conduits (e.g., Reaxon®) are CE-marked for bridging sensory nerve gaps and perform comparably to autografts in hand nerve lesions [146]. Additional clinical uses include hemostatic dressings and wound care [149, 152]. Alginate, derived from brown algae, forms hydrogels upon ionic crosslinking and is biocompatible but lacks intrinsic cell adhesion motifs, often requiring functionalization with Arginylglycylaspartic acid (RGD) peptides or ECM proteins [156, 158]. Degradation occurs via ion exchange or hydrolysis, tunable by crosslink density and oxidation [199]. Preclinically, alginate hydrogels have served as carriers for stem cells, GFs, and drugs in cardiac, tendon, and neural repair models [155, 157]. Clinically, alginate is widely used in wound dressings and hemostatic agents, supported by meta-analyses demonstrating efficacy in exudative and burn wounds [200]. Poloxamers are amphiphilic block copolymers of poly(ethylene oxide)–poly(propylene oxide)–poly(ethylene oxide) with thermosensitive sol–gel transitions near body temperature [160]. They are excreted renally and generally biocompatible [161]. Preclinically, poloxamer hydrogels have delivered neurotrophic factors and stem cells in PNI, improving axonal regeneration and functional recovery [159] Li, Li et al. 2018). Clinical translation remains limited due to mechanical stability concerns [201]. Gelatin, derived from collagen hydrolysis, contains RGD motifs that promote cell adhesion and neurite extension [167]. It is enzymatically degradable, primarily by matrix metalloproteinases, with rates modulated by crosslinking [165]. Preclinically, gelatin hydrogels have improved conduction and myelination in rodent nerve repair and have also been applied in bone regeneration, vascular grafts, and 3D cell culture [164, 168, 172]. Clinically, gelatin is used as absorbable hemostatic sponges (Gelfoam) and as carriers in wound healing and drug delivery [166]. Safety concerns persist regarding some chemical crosslinkers. Expanded polytetrafluoroethylene (ePTFE) is a permanent, nonresorbable, microporous polymer with high mechanical durability and resistance to enzymatic degradation [174]. Preclinically, Gore-Tex tubes were tested in peripheral nerve repair and vascular grafts [173, 175]. Clinically, Gore-Tex was approved as a nerve conduit for peripheral nerve gaps but its use has declined due to chronic inflammation and encapsulation, with current applications mainly in vascular grafts and hernia repair [176, 177]. Matrigel, a basement membrane extract from mouse sarcoma, is rich in laminin, collagen IV, and GFs [202]. While biodegradable, its undefined composition, batch variability, and immunogenicity preclude clinical use [183, 186]. It remains widely used preclinically for 3D cell culture, stem cell transplantation, angiogenesis assays, and tumor xenograft models [181, 184]. Composite scaffolds combining synthetic and natural polymers with bioactive ceramics have advanced bone and potentially nerve repair. Poly(lactic-co-glycolic acid) (PLGA)@Type I collagen (Col)/hydroxyapatite (HA) integrates the controlled degradation and mechanical strength of PLGA with collagen bioactivity and hydroxyapatite osteoconductivity [187, 188]. PLGA degrades into lactic and glycolic acids, which are naturally metabolized, while collagen and HA promote adhesion, osteogenic differentiation, and mineralization [191]. Preclinically, PLGA@Col/HA supports osteogenesis, angiogenesis, and neural ingrowth in bone defect models [189, 195]. Clinically, related HA–collagen and PLGA composites are approved as bone graft substitutes and guided bone regeneration materials in dentistry and spinal fusion, with outcomes comparable to autologous grafts [192, 193, 196].Table 1Properties, biodegradation, and clinical translation of scaffold biomaterials for nerve and tissue regenerationMaterialNatureBio-absorption/DegradationPreclinical applicationsClinical applicationsKey referencesChitinNatural polysaccharide, β-(1→4)-linked N-acetyl-d-glucosaminePoorly soluble, minimal degradation in vivo; enzymatic by chitinasesNerve conduits; combined with autologous nerves, platelet-rich plasma, or growth factors for enhanced regenerationLimited; experimental only[141] [142] [143], [144], [145]ChitosanPartially deacetylated chitin, cationic, forms hydrogels/fibersBiodegradable by lysozyme/hydrolysis; tunable by degree of deacetylationNerve conduits, growth factor/drug delivery, MSC-seeded scaffoldsCE-approved nerve guides (Reaxon®), wound dressings, hemostatic pads[68, 146–151], [152], [153, 154]AlginateLinear copolymer of guluronate and mannuronate from brown algaeIon exchange, slow hydrolysis, tunable by crosslinking and oxidationHydrogels for cell/drug delivery, stem cell encapsulation, tendon/cardiac repairWound dressings, hemostatic agents, controlled-release drug systems[52, 155–158]poloxamerSynthetic PEO–PPO–PEO block copolymer, thermosensitiveNot enzymatically degraded; cleared renally; reversible sol–gelInjectable hydrogels for controlled release, neurotrophic factor/stem cell delivery in nerve injuryLimited clinical use; investigated in musculoskeletal repair[159–162]GelatinHydrolyzed collagen, contains RGD motifsEnzymatically degraded (MMPs, proteases)Nerve conduits, bone regeneration, vascular grafts, 3D cell cultureHemostatic sponges (e.g., Gelfoam), wound healing carriers[163–172]ePTFE (Gore-Tex)Non-resorbable fluoropolymer, microporousBiologically inert; non-degradable, risk of fibrous encapsulationNerve conduits, vascular grafts, hernia repairFDA-approved nerve conduit (historic), vascular grafts, hernia meshes[173–180]MatrigelMurine basement membrane extract (laminin, collagen IV, growth factors)Biodegradable but undefined composition, immunogenic in humans3D culture, stem cell transplantation, angiogenesis assays, xenograftsNot clinically approved due to tumor origin, variability, immunogenicity[181–186]3D PLGA@Col/HAComposite: PLGA (synthetic), collagen (natural), hydroxyapatite (inorganic)PLGA hydrolyzes to lactic/glycolic acid; collagen/HA enhance bioactivityBone defect repair, spinal fusion, osteochondral models, angiogenesisBone graft substitutes in dental & orthopedic surgery, spinal fusion[187–196]PEO: poly(ethylene oxide); PPO: poly(propylene oxide); 3D: three-dimensional; FDA: U.S. Food and Drug Administration; ePTFE: expanded polytetrafluoroethylene; PLGA: poly(lactic-co-glycolic acid); HA: hydroxyapatite
For example, studies using chitosan conduits loaded with BMSCs-derived SC-like cells have demonstrated effective repair of sciatic nerve defects [123]. A study functionalizing chitosan nerve guides to repair a 15-mm sciatic nerve defect in rats demonstrates that the alignment of pro-regenerative cells within the conduit is critical for successful regeneration [203]. Functional analysis revealed that scaffolds aligned with SCs resulted in a 100% regeneration success rate [203]. This was superior to constructs aligned with MSCs, which had a 90% success rate, and significantly better than acellular bridges, which had only a 75% success rate [203]. These results indicate that the combination of chitosan conduits with ECM-enriched cellular gels, particularly those aligned with SCs, represents a promising alternative to autografts for repairing long, critical-size nerve gaps [203]. Alginate hydrogels, when combined with ADSCs and Gore-Tex tubes, significantly improve nerve conduction velocity and compound muscle action potential in facial nerve repair. Similarly, poloxamer hydrogels containing ADSCs have demonstrated favorable outcomes in sciatic nerve repair [159]. Gelatin hydrogels delivering SCs and transforming growth factor-β1 enhance axon regeneration and myelination comparably to those treated with autograft [168].
Furthermore, combining decellularized allografts with differentiated MSCs or undifferentiated MSCs nerve allografts significantly improves nerve regeneration, particularly neoangiogenesis, with outcomes comparable to autografts [204]. Challenges in using biomaterial scaffolds for MSC therapy include limited long-term stability, long-term efficacy, and safety of biomaterial scaffolds in vivo should be examined, including the degradation cycle, as well, the interaction between he tissues and degradation products [205].
Although the promising results of MSCs in preclinical studies, the tissue‑derived MSCs still face major drawbacks in the way for clinical use in peripheral nerve repair. This, including heterogeneity, is donor age, tissue source, and culture passage generate batches with widely differing transcriptomes, secretomes, and immunomodulatory strength, producing variable outcomes complicating regulatory standardisation and large-scale expansion of MSCs [206, 207]. After IV infusion, most MSCs become trapped in the lung’s small blood which limits the number reaching the injured site and raises the risk of micro‑emboli. Even when the cells are injected directly around the nerve, the inflamed, oxygen‑poor environment causes most of them to die within days, cutting short the paracrine signals needed for long‑term healing [208]. Finally, obtaining MSCs from bone marrow or fat involves an invasive procedure and produces only a limited number of cells, driving up production costs and complicating large‑scale manufacturing [209, 210].
Induced pluripotent stem‑cell (iPSC) technology directly overcomes these challenges. Reprogramming resets telomere length and epigenetic age, so iMSCs maintain proliferative capacity and immunomodulatory potency over extended passages, avoiding the senescence‑linked functional decline of primary MSCs [211, 212]. Because a single iPSC clone can be expanded indefinitely under xeno‑free GMP conditions and then differentiated en masse along a neural‑crest lineage, iMSC preparations are molecularly uniform and virtually limitless, resolving supply and batch‑to‑batch variability [213]. While in preclinical studies, iMSCs formed into scaffold‑free Bio‑3D conduits successfully bridged 5‑mm sciatic nerve gaps in rats, yielding superior axonal growth, electrophysiological recovery, and robust neovascularisation compared with inert silicone [126]. Furhermore, human iPSC‑derived neural precursors transplanted one week after rat spinal cord injury survived, migrated, and filled the lesion cavity. Grafts released neurotrophins, preserved white and grey matter, and significantly accelerated locomotor recovery on the Basso-Beattie-Bresnahan locomotor scale, beam‑walk, rotarod, and plantar tests. By 17 weeks, transplanted cells matured into multiple neuronal phenotypes, supporting structural and functional restoration [126].
Taken all together, iPSC technology supplies a limitless, rejuvenated, and more predictable cell source that avoids heterogeneity, aging, delivery, and supply challenges of traditional MSC therapy, contributing toward safer and more reliable treatments for peripheral nerve injuries.
Extracellular vesicles (EVs) are lipid bilayer-enclosed entities secreted by cells into the extracellular milieu. Although evidence of their existence dates back almost 80 years, only in the last two decades have the pathways of their generation, roles, and prospective uses started to emerge [214]. EV-related research has expanded significantly in recent years, and various pre-clinical experiments have been conducted in rodent animals, summarized in Table 2. “Extracellular vesicle” is a broad term that covers exosomes, microvesicles (often called ectosomes), and apoptotic bodies. Classification has traditionally relied on diameter and exosomes ranging from ~30–150 nm in size, ectosomes, and larger microvesicles (100–500 nm and up to 1000 nm) [228, 229]. Apoptotic bodies and the predominant subset of microvesicles are formed through direct outward budding from the plasma membrane of dying or activated cells [228]. Ectosome production relies on specialised lipid‑protein microdomains that assemble at the cell surface [229]. These vesicles generally lack the endosomal signature that defines exosomes and tend to be larger, although size ranges overlap [230]. By strict definition, the word “exosome” refers only to vesicles generated inside multivesicular bodies (MVBs) and released after MVB fusion with the plasma membrane. Because EVs vary widely in size, buoyant density, and cellular origin, and the absence of a universal molecular marker can unambiguously separate subtypes, nomenclature is challenging. The International Society for Extracellular Vesicles therefore issued the Minimal Information for Studies of Extracellular Vesicles MISEV guidelines to harmonise terminology and reporting standards [231].Table 2A summary of pre-clinical investigations on MSC-derived exosome therapies for PNIsCell sourceExosome dose & deliveryInjury/disease modelMechanism and functional outcomeReferencerADMSCInjected to the crushed nerve sites (dose not specified)Wistar rat, sciaticcrushMechanism unknown; vesicles notably boosted neurite extension in vitro and accelerated axon regrowth in vivo relative to PBS[215]rdADSCIn vitroNG10815 cellsTransfer of Gap43, Tau mRNAs and miRNAs; enhanced neurite elongation, matching SCexosome effect[216]rBMSC0.4 × 10^10^, 0.9 × 10^10^, 3.7 × 10^10^, and 7.4 × 10^10^/mlSD rat, sciaticcrushmiRNAdriven increased levels of VEGFA; 0.9 × 10^1^ ^0^ per ml exhibited highest axon diameter, LTP, SFI & motor recovery compared to the group treated with BMSCs.[217]rADSC6.25, 12.5, 25.0, and 50.0 μg/mLPrimary SC cultureAntiapoptotic shift (increase in Bcl2, decrease in Bax); reduced SC death and increased proliferation[218]hGMSC1 μg/μL in chitin conduitSD rat, SNIExos promoted SC division and DRG axon growth in vitro; in vivo conduit containing exos improved myelin, CMAP and motor scores vs the group with only chitin conduit treatment[142]rADSCInjected into crushed nerves (dose not specified)Wistar rat, SNImiR26b mediated reduce in Kpna2; reduced SC autophagy leads to improved remyelination[219]hBMSC3 × 10^9^/mL; intravitrealSD rat, opticnerve crushBroad miRNA cargos protected retinal cultures; in vivo preserved RGCs, promoted axon regrowth, and maintained function[220]hUCMSCChitosan–collagen conduits containing 400 µg/mL chitosanSD rat, segmental sciatic defectElevated BDNF expression enhances OEC survival under hypoxic conditions and leads to improved motor and sensory recovery compared to treatment with conduits alone.[221]hADSC1 mg/mL incorporated into MatrigelSD rat, sciatic defectIn vitro, upregulation of BDNF, CNTF, NGF; SC proliferation and migration; In vivo, more myelinated fibres, higher muscle mass[182]hADSC/hBMSC100 μg/ml/injected in lateral aspect of the penisSD rat, BCNIRestored nNOS, neurofilament, vWF, αSMA and erectile function[222]BMSC2 μg/μL administered to both proximal and distal nerve stumps.SD rat, sciaticcrushTSG6, NFκB/NLRP3 that enhances M2 Macrophage polarization; better axon regrowth, myelination, locomotor recovery than normal-conditioned BMSC-exosomes[223]rADSC5, 10, and 20 μg/μL,DRG and SC culturesmiR223p decreases PTEN, AKT/mTOR increase leads to longer DRG neurites and more SC proliferation/migration vs PBS treated group[224]hADSCiSCSD rat, sciaticdefectSOX2/FN1 modulation → ↑ SC viability; conduit + exos boosted nerve repair vs conduit alone[133]hADMSCs, dhADMSCSC3 × 10^9^/ml/in NT3 chitosan conduit (in vitro)SC, sensory neuronsExos rich in miR1323p, miR199b5p, which have superior protective effects on neurons, and anti-inflammation. Compared to uhADMSC exosomes, dhADMSC-SCs secreted higher levels of neurotrophic and angiogenic growth factors, promoting axonal growth and regeneration.[144]hUCMSC400 μg/ml in soft hydrogelSD rat, sciaticcrushloaded into soft hydrogels exhibited rapid exorelease; reduced proinflammatory factors and improved functional recovery vs stiff gel[225]rADMSC1.9 × 10^8^ /µL + BCM scaffoldSD rat, sciaticdefectBCM + exos improved fibre number/diameter and thermal latency, Similar to autograft[226]hA MSC40 μg/ml in prefabricated graftSD rat, SNISCLCs-exosomes ↑ SOX10, Egr2, OCTN6 and myelin proteins → superior SC proliferation, axon regrowth, vs hAMSCexos[227]hUCMSC10 µl ml; Loaded into 3D PLGA@Col/HASD rat, SNIPLGA@Col/HA loaded with hUC-MSC-exosome restored SFI, CMAP, muscle mass, angiogenic effect comparable to autograft[194]MSCs: mesenchymal stem cells; ADMSCs: adipose-derived mesenchymal stem cells; rADMSCs: rat adipose-derived stem cells; PBS: phosphate buffered saline; ADSCs: adipose-derived stem cells; dADSCs: differentiated adipose-derived stem cells; rdADSCs: rat differentiated adipose-derived stem cells; SCs: Schwann cells; Gap43: Growth associated protein 43; DRG: dorsal root ganglion; SD Sprague Dawley rats; BMSCs: bone marrow mesenchymal stem cells; rBMSCs: rat bone marrow mesenchymal stem cells; VEGFA: vascular endothelial growth factor A; LTP: latency of thermal pain; SFI: sciatic function index; hGMSCs: Human-gingival mesenchymal stem cells; SNI: sciatic nerve injury; vs :versus; ↑:increasee; ↓:decrease; →:lead to CAMP: compound muscle action potential; Kpna2: karyopherin subunit alpha 2; rADSCs: rat adipose-derived stem cells; hBMSCs: human bone marrow mesenchymal stem cells; ONC: optic nerve crush; RGCs: retinal ganglion cells; hUC-MSCs: human umbilical cord mesenchymal stem cells; BDNF: brain-derived neurotrophic factor; OECs: olfactory ensheathing cells; CNTF: ciliary neurotrophic factor; NGF: nerve growth factor; BCNI: bilateral cavernous nerve crush injury; nNOS: neuronal nitric oxide synthase; α-SMA: alpha-smooth muscle actin; LPS: lipopolysaccharide; PTEN: Phosphatase and Tensin Homolog deleted on Chromosome 10; hADSCs: human adipose-derived stem cells; iSCs; induced SClike cells; hADMSC -SCs: human adipose-derived mesenchymal stem cell-Schwann like cells; FN1: fibronectin 1; NT-3: neurotrophin 3; ADMSCs: adipose-derived mesenchymal stem cells; hADMSCs: human adipose-derived mesenchymal stem cells; dhADMSCs: differentiated human adipose-derived mesenchymal stem cells; uhADMSCs: undifferentiated human adipose-derived mesenchymal stem cells; BCM: Biosynthetic cellulose membrane; SCLCs: Schwann cell-like cells; hAMSCs; human amniotic MSCs: 3D PLGA@Col/HA: a three-dimensional composite conduit featuring a collagen/hyaluronic acid inner sponge encapsulated within an electrospun hollow poly(lactic-co-glycolic acid) scaffold
Direct MSC transplantation has several drawbacks. MSC‑derived exosomes (MSC-Exos) are now being investigated as a cell‑free strategy to enhance peripheral nerve regeneration [232]. Exosomes, lipid‑bilayered vesicles typically 30–150 nm in diameter, are produced by nearly every mammalian cell type, including immune cells, neurons, tumour cells, epithelial cells, osteocytes, myocytes, and MSCs [233] (Fig. 7). Initially identified as a cellular disposal route for unwanted biomolecules, exosomes are now recognized as key mediators of intercellular communication. Their biogenesis is orchestrated through the endocytic pathway. Briefly, the plasma membrane’s inner budding produces tiny vesicles. Subsequently, these components fuse to form early endosomes. During their maturation, intraluminal vesicles (ILVs) emerge through the inward budding of the limiting endosomal membrane, an essential step in exosome biogenesis [234]. ILVs formation is orchestrated by the endosomal sorting complex required for transport. During this maturation process, cargo molecules such as RNAs are selectively encapsulated within ILVs. ILVs accumulate within late endosomes to produce MVBs [234]. MVBs either fuse with lysosomes for degradation or merge with the plasma membrane, releasing their ILVs as exosomes into the extracellular space. Under transmission electron microscopy (TEM), exosomes display a characteristic cup‑shaped morphology and float at densities between 1.13 and 1.19 g/ml [235]. Isolation of exosomes from culture supernatants or biological fluids can be achieved by ultracentrifugation, density‑gradient separation, immunoaffinity capture, paper‑based affinity platforms, or size‑exclusion chromatography [236]. Ultracentrifugation and gradient protocols are most common, however, the limitations include being time‑consuming, requiring large volumes, and occasionally compromising vesicle integrity [236]. Cargo profiles mirror the physiological state and lineage of the parent proteins (Rab GTPases, flotillin, Alix, TSG101, Hsp70, Hsp90, tetraspanins CD9, CD63, CD81), lipids, metabolites, and nucleic acids (mRNA, miRNA, rRNA, circRNA, lncRNA) [237]. Evidence from mammalian culture systems shows that oxidative or thermal stress triggers a calcium‑dependent activation of the endosomal sorting complex required for transport machinery, boosting exosome biogenesis by 8‑ to 10‑fold within 7 h [238]. Exosomes specifically modify their cargo to promote tissue repair. For instance, SC-derived exosomes enriched in Milk Fat Globule–Epidermal Growth Factor–Factor 8 (MFG-E8), a secreted glycoprotein that bridges phosphatidylserine on apoptotic cells with integrins (αvβ3/αvβ5) on phagocytes, promote apoptotic debris clearance and anti-inflammatory signaling [239, 240]. MFG-E8 plays a pivotal role in maintaining tissue homeostasis and facilitating nerve regeneration by inducing macrophage and microglial polarization toward an anti-inflammatory M2 phenotype via SOCS3/STAT3 signaling, thereby suppressing secondary inflammation and accelerating axonal regrowth following spinal cord or sciatic nerve injury [239, 240]. Exosome release can increase significantly under microbial attack, oxidative stress, or other challenges, underscoring their role in cell stress responses [240]. MSC-Exos carry bioactive proteins that promote immunomodulation and tissue regeneration. Proteomic surveys show that adipose and umbilical cord MSC-exos are noticeably enriched in basic FGF, VEGF, and HGF [241]. When delivered to ischemic or injured tissue, these vesicular GFs amplify endothelial proliferation and neovascular sprouting, thereby accelerating granulation tissue maturation, angiogenesis, and matrix remodelling during tissue repair [241]. MSC-derived exosomes promote tissue repair through their RNA cargo. Engineered exosomes carrying IFN-γ mRNA can traverse restrictive biological interfaces such as the blood–brain barrier and the dense glioblastoma microenvironment. Once internalized, the translated IFN-γ protein reprograms tumor-associated and injury-site immune cells, highlighting exosomal mRNA as a promising and safe vector for targeted protein replacement therapy [242]. MSC-derived exosomes enriched with miRNAs such as miR-182-5p or miR-21 suppress pyro-inflammatory genes, including Gasdermin D, thereby inhibiting apoptosis and reducing infarct size in models of myocardial ischemia–reperfusion injury [243]. Gasdermin D is a pore-forming protein that mediates pyroptosis, a form of inflammatory programmed cell death [243]. Adipose mesenchymal stem cell-derived exosomes (ADMSC-Exos) deliver MALAT1 and other lncRNAs, which sponge miR-124, activate Wnt/β-catenin, and promote angiogenic, scar-free skin healing in diabetic wounds [244]. Furthermore, MSC‑Exos carry anti‑inflammatory ILs and their upstream vesicles derived from cytokine‑primed adipose MSCs transport IL‑10 as well as TGF‑β and concurrently repress transcripts for IL‑1β, IL‑6, and TNF‑α in recipient macrophages, which shift phagocytes toward an M2 reparative phenotype and decrease chronic inflammation in vivo [245]. Through the concerted delivery of growth‑factor and IL payloads, MSC‑derived exosomes create a paracrine niche that both mollifies immune responses and promotes trophic signals indispensable for tissue regeneration.Fig. 7Biogenesis, release, and molecular composition of exosomes. Plasma-membrane invagination forms intraluminal vesicles inside multivesicular bodies; fusion with the plasma membrane releases
As central mediators of intercellular communication, exosomes are significantly involved in regenerative they can deliver neurotrophic factors, regulate immune responses, modulate vascular growth, and influence cellular metabolism, thereby aiding tissue repair [246]. Numerous studies indicate that exosomes are favoured over both cell transplants and synthetic vesicles as precision vectors for delivering pro‑regenerative signals in peripheral nerve repair. Their bi‑layered membrane and innate compatibility minimise immunogenicity and shield their therapeutic cargo from degradation [247]. Consequently, MSC‑derived exosomes (MSC‑Exos) have emerged as promising, low‑risk therapeutics for PNI. Exosome treatment accelerated functional recovery, reduced fibrosis, enhanced remyelination, and attenuated muscle atrophy following exosome administration in rodent sciatic nerve models. Collectively, these findings support continued development of MSC‑exosomes as a scalable, cell‑free platform for peripheral nerve regeneration. For this aim, we investigated typical examples of up-to-date pre-clinical studies.
Exosomes released by differentiated adipose-derived MSCs notably enhanced the neurite elongation of NG108-15 neurons, a mouse neuroblastoma–rat glioma hybrid cell line commonly used as a neuronal model [216]. Exosomes from SCs and differentiated ADMSCs contained mRNAs reflecting transfer of pro-regenerative mRNAs and miRNAs (miR-1, miR-18a, miR-21, miR-182, miR-222) [216]. Another study suggests that exosomes derived from ADSC, which contain multiple neurotrophic factors such as BDNF, FGF-1, GDNF, IGF-1, and NGF that promote neural survival and axonal growth, play a vital role in peripheral nerve regeneration and represent a promising therapeutic approach for tissue-engineered nerve repair [215]. While in vivo, weekly administration of ADSC-Exos injected proximally and distally to the crushed nerve site with a Hamilton syringe after rat sciatic crush improved walking-track analysis, increased axonal diameter, and elevated neurotrophic factor expression [212]. Parallel findings were obtained with BMSCs-Exos. The researcher identified a dose range that resulted in the highest axon counts, myelin thickness, and sciatic functional index, with exosomes at four different doses injected into the gastrocnemius muscles of rats with sciatic nerve crush injury. These findings were attributable to miRNA-mediated regulation of PMP22, VEGFA, NGFR, and S100B [217]. Gingiva-derived MSC (GMSC-Exos) combined with biodegradable chitin conduits further increased fibre number, diameter, and myelination, restoring compound muscle action potentials and gait, compared to the control group without GMSC-Exos [142]. More recently, a three-dimensional PLGA@Col/HA conduit loaded with human umbilical-cord MSC exosomes bridged a 10-mm gap, exosomes promoted angiogenesis, and achieved functional recovery similar to autografting [248].
After injury, mature myelinating SCs differentiate into “repair” SCs, proliferate, and clear myelin debris in concert with macrophages [249]. Transition from a myelinating to a repair phenotype is regulated by transcription factors including c-Jun and Sox2, and is sustained in part by SC-derived exosomes enriched in miR-21, which downregulates Phosphatase and Tensin Homolog deleted on chromosome Ten (PTEN) and activates neuronal PI3K signalling [250]. MSC exosomes reinforce these processes. ADSC -Exos promote SC proliferation and Ki-67 expression, extend DRG neurites, and upregulate neurotrophic genes [215]. Further study showed that exosomal miR-22-3p suppresses PTEN, activates AKT/mTOR, and simultaneously stimulates DRG axon elongation and SC migration [224]. Innovative bio-hybrid approaches are being developed to sharpen these effects. Induced Schwann-like cell-derived exosomesseeded in NT-3-laden chitosan conduits restored sensory function via the SOX2/FN1 axis and exosome release [133]. Exosomes derived from differentiated human ADMSCs (dExo) contained more neuroprotective miRNAs (miR-132-3p, miR-199b-5p) than those from undifferentiated cells; dExo better attenuated oxidative stress in SCs, enhanced angiogenesis, and promoted sensory-neuron outgrowth [144]. Similarly, exosomes from SC-like cells differentiated from hAMSC -Exos and SCLCs-Exos promoted SC proliferation, migration, and expression of myelin regulators (SOX10, EGR2, MBP, MPZ). In a rat sciatic gap model, a prefabricated nerve graft was used to deliver hAMSCs-exo or SCLCs-exo in an injured sciatic nerve rat model, and SCLC-exo accelerated motor recovery, reduced muscle atrophy, and increased angiogenesis [227].
Peripheral nerve regeneration is deeply linked to revascularization. VEGF and its receptors orchestrate this process [251]. Recent studies indicated that MSC exosomes are potent pro-angiogenic stimuli. For instance, MSC-derived sEVs were loaded onto the heparinized PCL grafts to obtain functional vascular grafts. Followed by implantation to replace a segment of rat abdominal artery (1 cm) for up to 3 months [252]. MSC-EVs markedly enhance the regeneration of vascular smooth muscle and endothelium. This regenerative effect is likely driven by the sEV cargo, particularly VEGF, miR-126, and miR-145 [252]. Furthermore, neural stem cell exosomes enriched in VEGF-A boost spinal-cord microvascular endothelial-cell angiogenesis in vitro and, when delivered via injection into the tail vein after spinal cord traumatic injury, promote microvascular regeneration, reduce lesion cavities, and improve Basso scores [253]. Furthermore, MSCs-Exos transfected with miRNA-126 delivered to injured spinal cord tissue via tail vein injection, suppress SPRED1/PIK3R2, enhance angiogenesis and reduce apoptosis, lesion volume, and significantly improve functional recovery [254]. Human-urine stem cell-derived exosomes embedded in an injectable hydrogel locally delivered into the injured spinal cord, deliver ANGPTL3, activate PI3K/AKT-dependent angiogenesis, and thereby significantly improve restoration of neurological function following spinal-cord injury [255]. In peripheral-nerve models, the 3D PLGA@Col/HA conduit cited above also increased CD31-positive microvessels when loaded with hUCMSC exosomes, coupling angiogenesis with enhanced axonal bridging [190].
PNI triggers Wallerian degeneration, during which SCs and infiltrating macrophages clear myelin debris and secrete cytokines. Early pro-inflammatory mediators (TNF-α, IL-1α/β, IFN-γ) recruit leukocytes, whereas later anti-inflammatory cytokines (IL-4, IL-10, TGF-β) are expressed and suppress the production of all cytokines [256]. An appropriately balanced inflammatory milieu is excess inflammation causes neuropathic pain and impedes regeneration, whereas insufficient activation leaves inhibitory debris unresolved. MSC exosomes provide immunomodulatory properties [257]. Exosomes derived from lipopolysaccharide-preconditioned mesenchymal stem cells (LPS-Exos) markedly promote sciatic nerve regeneration. In vivo, LPS-Exos induce macrophage polarization toward the pro-regenerative M2 phenotype, suppress NF-κB/NLRP3 signaling axis, and enhance axonal elongation and remyelination [223]. In another study, dADSC exosomes (dExos) were more effective than undifferentiated exosomes (uExos) in down-regulating TNF-α, IL-1β, and IL-6 expression in human monocyte–derived macrophages (Mɸ) that were differentiated from peripheral blood mononuclear cells and subsequently stimulated with lipopolysaccharide and interferon-gamma to induce a proinflammatory (M1-like) phenotype, a property related to enriched anti-inflammatory miRNA-132-3p and miRNA-199b-5p [144]. When MSC-Exos were loaded in soft hydrogels, MSC-Exos reduced macrophage infiltration and pro-inflammatory cytokine expression, leading to superior functional recovery in a rat sciatic-crush model [225].
MSC-derived exosomes and EVs can be administered locally at injury sites, incorporated into biomaterial scaffolds such as conduits, hydrogels, or microspheres, or delivered systemically by intravenous (IV) injection [215] [142] [254], [217] [227, 248, 253, 255]. A systematic in vivo study in mice demonstrated that EV fate is strongly shaped by their cellular origin, dose, route of administration, and the presence of targeting ligands [258]. While most EVs accumulate in the liver and spleen, subsets can be directed to specific tissues, such as tumors, highlighting their potential as drug delivery vehicles [258]. Recent positron emission tomography-imaging studies confirmed that biodistribution is not universal but depends on EV origin, host immune status, and labeling method [259]. For example, macrophage-derived EVs show dominant hepatic uptake, while MSC-derived EVs exhibit extended circulation and relatively low liver, spleen, and lung retention. Notably, biodistribution patterns are significantly altered by immune competence, with splenic uptake of MSC-EVs markedly increased in immunodeficient mice [259]. These results underscore that both EV source and host physiology critically influence translational outcomes. Meta-analyses further confirm that EVs consistently distribute to liver, lungs, kidneys, and spleen across species and cell sources, although kinetics vary by size and time [260]. Small EVs preferentially accumulate in liver early after dosing, whereas larger EVs initially lodge in the lungs before redistribution [260]. Methodological heterogeneity across studies has complicated comparisons, reinforcing calls for standardized guidelines for EV biodistribution research [260].
Extravasation of EVs into parenchymal tissues depends on vascular permeability, inflammatory status, and EV surface properties [261, 262]. In blood–brain barrier models, HEK293T- or erythrocyte-derived EVs crossed only under inflammatory conditions, with LPS treatment increasing permeability by ~300% [261]. In LPS-induced lung injury, EV accumulation in inflamed tissue was significantly higher than in healthy controls, peaking at maximal vascular leak (24–48 h) [262]. Nonetheless, only a small fraction of circulating EVs typically reaches a remote lesion after systemic dosing, with reports suggesting < 1% of injected EVs access the brain [260, 263]. Local injection or incorporation into biomaterial scaffolds improves retention at the target site by orders of magnitude compared to systemic administration [262, 264]. Route, dose, and local pathology remain the dominant determinants of EV uptake in vivo [260].
Engineering approaches are being developed to enhance targeting, including genetic editing of EV membrane proteins to display ligands or homing peptides that increase tropism and therapeutic efficacy [263]. Such strategies may mitigate the challenge of non-specific biodistribution, as EVs carry proteins, lipids, and RNAs capable of modulating immune function, metabolism, fibrosis, angiogenesis, and tumor biology [118, 265, 266]. Because liver and immune organs encounter the majority of systemically delivered EVs, MSC-EVs often exert immunomodulatory or trophic effects in these compartments, which may be beneficial in inflammatory disease but undesirable in other contexts [267]. Moreover, tumor-derived EVs illustrate risks of non-specific they can reprogram fibroblasts into cancer-associated fibroblasts, enhance angiogenesis, and drive metastasis [268, 269]. Consequently, immune modulation, pro-fibrotic or angiogenic signaling, and altered metabolism represent important safety considerations [267, 270, 271].
Non-surgical, based therapies like exercise, electrical stimulation (ES), extracorporeal shock wave therapy (ESWT), and low-intensity pulsed ultrasound (LIPUS) have been employed as adjuncts to the microsurgical repair of peripheral nerve injuries.
Exercise is the most accessible of these interventions. In rodent models, moderate treadmill, ladder-climb, or swimming protocols consistently increase axon regeneration, thicken myelin, muscle mass, reinnervation, and accelerate functional recovery after sciatic-nerve crush injury [272]. Mechanisms include enhanced SC proliferation, elevation of GDNF, BDNF, and IGF-1, neurotrophin expression, and a macrophage shift from M1 to pro-regenerative M2 phenotypes [273]. Further, applying a combination of exercises with fibrin glue in transected nerve markedly boosts bridge vascularisation, extends axon length, enhances neuromuscular-junction re-innervation, and accelerates functional recovery [274]. A review about the role of exercise in clinical application revealed that exercise programmes consistently reduce pain and enhance physical performance, while also improving deep sensation, thermal perception, and static balance. By limiting axonal degeneration and supporting peripheral neuronal function, limb-specific rehabilitation can strengthen muscles, restore balance, and enhance quality of life
in chemotherapy-induced peripheral neuropathy [275]. Comparable benefits are documented in diabetic neuropathy, where endurance, balance, and overall physical function are significantly improved through endurance, balance, and multimodal training protocols [275].
The effectiveness of ES applied to the proximal stump of injured nerves in promoting nerve regeneration has been confirmed in multiple animal studies [276]. Recently, bioresorbable electronic stimulators have emerged as a promising technology in bioelectronic medicines, offering a novel therapeutic platform for modulating disease processes, accelerating wound repair, and eliminating infections to improve clinical outcomes [277]. In a 10 mm sciatic nerve gap injury model in SD rats continuous ES using a biodegradable, self-electrified conduit constructed from dissolvable galvanic cells on a nerve guiding scaffold, improved axonal regeneration, SC, and neurotrophic factors [221]. Furthermore, in a sciatic nerve injury model using dynamic covalent polymer–based bioresorbable stimulators that deliver stable wireless ES, these devices maintain muscle excitability, enhance conduction, and accelerate functional recovery [277]. ES protocols and outcome assessments varied across clinical trials in which electrodes were implanted at the repair site (one study also using a fully implanted pulse generator). Sessions lasted either 20 minutes or 60 minutes at a median frequency of 20 Hz, with intensities ranging from 3 V to 30 V and pulse widths of 0.1 ms to 1.007 ms. ES were received either perioperative, intraoperatively, or immediately afterwards. The recovery was quantified by electrophysiological measures alongside motor, sensory, and functional criteria. Across all studies, those receiving adjunctive electrical stimulation demonstrated consistently superior nerve regeneration and functional improvement compared with control animals that underwent identical nerve repair but without ES [278, 279].
There are typically two types of ESWT: focused shock wave therapy (FESWT) and radial shock wave therapy (RESWT), which stimulate tissue regeneration via mechanotransduction, making it an effective noninvasive method for peripheral nerve regeneration [280]. Preclinical rodent studies consistently demonstrate that low-energy ESWT accelerates morphological regeneration and improves electrophysiological outcomes after sciatic nerve injury [281]. Low-energy ESWT promotes peripheral nerve regeneration through multiple mechanisms. Activating ERK/MAPK signaling and the p75 neurotrophin receptor [282]. ESWT stimulates SC proliferation, which leads to an increase in secretion of key neurotrophic factors. Among these, BDNF is upregulated via SC-dependent Janus kinase (JAK)/signal transducer and activator of transcription (JAK/STAT). In addition to direct trophic effects, ESWT boosts the expression of VEGF, angiopoietin-1, essential to angiogenesis [283, 284]. The therapy also modulates the inflammatory it attenuates M1 macrophage activation and promotes an M2, anti-inflammatory phenotype, thereby limiting pro-inflammatory cytokine release and promoting a regenerative environment [285].
Clinical data supporting ESWT in peripheral neuropathy are arising. In a systematic study, employing ESWT in peripheral neuropathy showed electrophysiological improvements compared to control treatment [286]. ESWT may reduce short-term motor nerve distal latency, improve short- and mid-term sensory nerve conduction velocity. Further subgroup analyses revealed that ESWT was more successful than the Control group in enhancing sensory nerve conduction velocity (short-term, MD, 4.36; 95% CI: 1.23, 7.49) and reducing mid-term sensory nerve action potential distal latency (MD, −0.39; 95% CI: −0.52, −0.26) (Yang, Li et al. 2024). For motor nerves, it reduced short-term motor nerve distal latency (MD, −0.61; 95% CI: −0.91, −0.30) [286]. Combining ESWT and physical therapy (PT) was more effective than PT alone in lowering sensory nerve action potential amplitude and distal latency, enhancing sensory nerve conduction velocity, and decreasing motor nerve distal latency [286].
LIPUS is a type of ultrasound that is usually utilized for therapeutic purposes in rehabilitation medicine. It is provided at a much lower intensity ( < 3 W/cm2) than conventional ultrasound is emitted in a pulsed wave mode [287]. Because of its pulsed output mode and low intensity, LIPUS has minimal thermal impacts, making it a potential noninvasive technique for tissue regeneration [288]. By converting acoustic pressure into biological signals, LIPUS activates mechanotransductive pathways in neurons and SCs, enhances trophic support, reduces inflammation, and increases angiogenesis, all essential for efficient nerve regeneration [289]. LIPUS has attracted significant interest in peripheral nerve regeneration due to its noninvasive delivery and outstanding safety profile. Early rat investigations showed that daily LIPUS sessions for 12 days after sciatic nerve axotomy accelerated morphological regeneration, increased fibre counts, and improved conduction velocities compared to sham controls [290]. In a 10 mm reversed autograft model, LIPUS at intensities of 250, 500, or 750 mW/cm^2^ improved the sciatic functional index and CMAP amplitude [287]. Following sciatic nerve transplantation, treatment with 250, 500, or 750 mW/cm^2^ LIPUS revealed that the lowest intensity of 250 mW/cm^2^ significantly accelerated axonal regeneration. This was confirmed by regenerated myelinated nerve fiber the 250 mW/cm^2^ group had the highest density (19725.6 ± 913.9 n/mm^2^), followed by 500 mW/cm^2^ (16089 ± 1154.4 n/mm^2^), 750 mW/cm^2^ (14172.4 ± 813.8 n/mm^2^), and the control (12910.8 ± 1764.1 n/mm^2^). Statistical analysis showed significant differences (p = 0.000) between all groups except between the control and the 750 mW/cm^2^ group (p = 0.121), demonstrating that lower LIPUS intensity is more effective [287]. Histology demonstrated more brittle axonal regrowth, likely through neovascularization and increased neurotrophic factor release [287]. In an in-nerve compression model, LIPUS decreased pro-inflammatory cytokines (TNF-α, IL-6) and growth inhibitors (SEMA3A, GSK3β), demonstrating its anti-inflammatory and regenerative actions [291]. Mechanistically, LIPUS delivers mechanical signals to the plasma membrane ECM interface, triggering, activating integrin focal adhesion kinase signaling and downstream ERK1/2 and mTOR pathways key regulators of neuronal survival, proliferation and differentiation. LIPUS also engages the TrkB/Akt/CREB pathway in SC, promoting proliferation, angiogenesis, and remyelination [292]. Enhanced SC metabolism under LIPUS drives secretion of β-NGF, BDNF, and GDNF, creating a trophic microenvironment for axonal extension [293]. In addition, LIPUS at 5 MHz, 500 mW/cm^2^ significantly enhanced SC neurotrophic factor secretion (β-NGF, BDNF, GDNF), suppressed reactive oxygen species (ROS) and TNF-α via NF-κB pathway, induced antioxidant effects, and promoted M2 macrophage polarization [293].
Engineered NGCs are now the principal biomaterial alternative to autografts. Despite advances in micro-architectural alignment, controlled growth factor release, and incorporation of conductive elements in commercially approved NGCs, their clinical application remains limited to the repair of minor injuries, typically short digital or facial nerve gaps ( < 3 cm). Their capacity to achieve functional reinnervation across critical-sized defects remains inadequate, necessitating further development.
In recent years, an increasing body of research has explored the effects of combining these, ES, ESWT, or LIPUS, with bioengineered components, resulting in outcomes superior to those obtained by any single modality [294]. Recent studies show that appropriately adjusted mechanical and electrical stimuli can significantly reduce the expression of pro-inflammatory cytokines [295] and reprogram the post-PNI environment; nonetheless, long-term functional recovery following large-gap or chronic injuries would probably require more than biophysical stimulation alone. A recent study using electrospun piezoelectric PVDF-TrFE nanofiber conduits, periodically shockwave-activated in vivo, delivered mechano-electrical stimulation across 15 mm rat sciatic gaps. This non-invasive stimulus enhances SC maturation, axonal regrowth, myelination, and superior functional recovery compared to the control group, indicating piezoelectric scaffold-based physical cues as a promising strategy for peripheral nerve repair [296]. Furthermore, tissue engineering strategies have employed synthesized electroconductive polypyrrole nanoparticles with enhanced hydrophilicity to facilitate uniform dispersion with collagen hydrogel matrices. Subsequently, cell-laden collagen polypyrrole hybrid hydrogel microfibers with highly aligned microstructures were engineered using a microfluidic chip tailored to replicate the architecture of native ECM.
The scaffold guided neuronal alignment, enhanced neurite outgrowth, and, when combined with electrical stimulation, up-regulated neurogenic genes through L-type calcium-channel activation, increasing intracellular Ca^2^ ^+^ and accelerating neurogenesis, offering a promising platform for nerve-tissue engineering applications [297].
In streptozocin-diabetic rats, intracavernosal ADSCs injection combined with LIPUS at 200 mW/cm^2^ achieved significantly higher recovery of erectile function [298]. Erectile function improved to different extents with intracavernosal ADSCs (0.32 ± 0.03, 0.23 ± 0.01), LIPUS treatment (0.46 ± 0.01, 0.32 ± 0.02), and the combination of ADSCs plus LIPUS (0.61 ± 0.02, 0.46 ± 0.03), as shown by notable increases in maximal and total ICP/MAP ratios [298]. LIPUS accelerated ADSC proliferation and enhanced CXCL12, FGF-2, and VEGF release via Piezo-activated ERK signalling, indicating that LIPUS potentiates ADSC therapy for diabetic erectile dysfunction through a Piezo-ERK-VEGF mechanism (Liu, Jiang et al. 2022). Furthermore, recent evidence indicates that exosomes, especially those from MSCs, are highly sensitive to physical factors. These exosomes, which are critical for intercellular communication, can undergo beneficial modifications when exposed to certain physical stimuli [299]. By modifying their properties through exposure to well-defined physical interventions, it is possible to enhance the effectiveness of MSC-Exo and improve their therapeutic impact on the cellular microenvironment [300]. Building on this idea, SC exosomes produced under LIPUS stimulation were superior to untreated they extended major pelvic ganglion/cavernous nerve neurites in vitro, enhanced cavernous-nerve regeneration, SC proliferation, and restored erectile function in vivo [301]. High-throughput sequencing was performed to profile miRNA expression in LIPUS-SCs-Exo and SCs-Exo, with 68 differentially expressed miRNAs identified, several of which are known to regulate processes such as axonal growth and neurological diseases. Validation by qRT-PCR confirmed changes in 10 miRNAs, and computational prediction (miRWalk) revealed 1,689 potential target genes. KEGG pathway analysis indicated enrichment in multiple signaling cascades, including PI3K–Akt, AMPK, FoxO, MAPK, axon guidance, and neurotrophin pathways. Results of western blot of the PI3K–Akt–FoxO axis and confirmed increased phosphorylation of PI3K, Akt, and FoxO proteins in major pelvic ganglion neurons following LIPUS-SCs-Exo treatment, demonstrating the supplementary function of exosome cargo and LIPUS [301]. Taken together by adding physical factors, it is possible to enhance the regenerative potential of MSC-Exo, providing a more targeted and effective approach to treating a wide range of diseases and injuries.
Before going to clinical trials, based on previous pre-clinical studies, biopharmacological approaches (exogenous GFs, neurotrophic peptides, small-molecule modulators) provide targeted molecular cues that accelerate axonal sprouting and remyelination and can be formulated for local or sustained delivery, but they face delivery, dose-stability and off-target-effect challenges in clinical translation [78]. Cellular therapies—transplantation of SCs or mesenchymal/stem cells—supply living trophic support, ECM guidance and immunomodulation and are among the most potent strategies for bridging gaps and improving histological regeneration in animal models; however, they require complex manufacturing, carry immunologic/oncologic/regulatory hurdles and human evidence is still limited [302]. Exosome therapies are a rising, cell-free alternative that deliver many of the paracrine benefits of cells (miRNA, proteins) with lower immunogenicity and easier storage/delivery; preclinical data show improved axon counts, reduced atrophy and functional gains, but clinical translation is nascent [303]. Physical “shock” modalities (low-intensity extracorporeal shockwave therapy and targeted electrical stimulation) act by enhancing local blood flow, modulating inflammation and activating regeneration-associated signalling to speed recovery without implants; they are minimally invasive and already used clinically for some neuropathies, though parameter optimization remains under study [281]. Finally, exercise and rehabilitation produce activity-dependent they upregulate endogenous neurotrophic factors, preserve muscle, reduce neuropathic pain and improve functional reinnervation. Exercise is low-cost, low-risk and broadly applicable, but its regenerative effect is generally slower and smaller in magnitude than the structural repair produced by targeted biological or cellular interventions in preclinical models [304].
When comparing exercise with more invasive therapies, including cell therapy, conduits, grafts, and ES, in PNIs, most are preclinical in animals. There remains a paucity of human RCTs doing direct head-to-head comparisons of exercise and invasive biologics, but animal data give us useful insights. Cobianchi, Casals-Diaz et al. investigated the effects of acute ES and early treadmill running (TR) on nerve regeneration and neuropathic pain in rats after sciatic nerve injury [305]. The key findings were that TR primarily reduced mechanical allodynia, significantly increasing withdrawal thresholds versus controls at 60 days post-injury (dpi) (TR: 20.9 ± 1.5 g vs. C: 17.1 ± 1.1 g; p < 0.05). In contrast, ES accelerated sensory and motor reinnervation, leading to earlier recovery of thermal sensitivity (e.g., 12.5 ± 0.6 s vs. C: 16.4 ± 0.7 s at 31 dpi; p < 0.05) and improved mechanical thresholds later in recovery (ES: 20.7 ± 1.3 g vs. C: 12.8 ± 1.3 g at 60 dpi; p < 0.001). The combination therapy (ES+TR) was most effective, showing strong, synergistic hypoalgesia and the highest final mechanical threshold (23.4 ± 1.7 g at 60 dpi; p < 0.01). These functional improvements were linked to distinct, treatment-specific modulations of neurotrophic factor mRNA levels in sensory and motor neurons [305]. This study also demonstrates that ES significantly accelerates motor nerve reinnervation. In the proximal tibialis anterior (TA) muscle, the ES group showed a significantly higher CMAP amplitude from 21 to 61 days post-injury, culminating at 52 ± 7% of normal versus 31 ± 3% in controls (p < 0.01). This benefit extended to the distal plantar (PL) muscle, where the ES group’s CMAP reached 19 ± 4% versus 10 ± 1% in controls (p < 0.01). TR provided a more modest improvement, increasing TA CMAP to 39 ± 3% at 61 dpi (p < 0.05), while the combination therapy (ES+TR) yielded results similar to ES alone in the TA muscle [305]. Histologically, ES groups showed a 12% lower mean myelinated fiber count, suggesting more mature, refined regeneration [305]. Activity treatments differentially regulate neurotrophic factor expression after nerve injury [305]. ES induced an early, transient upregulation, significantly increasing BDNF in the ventral horn at 1 dpi (p < 0.01) and 3 dpi (p < 0.05), and GDNF in the DRG at 1 dpi (p < 0.05). Conversely, ES significantly reduced the injury-induced elevation of BDNF (p < 0.05) and NGF (p < 0.01) in the DRG by 3 dpi. ES also uniquely increased NT3 mRNA in the ventral horn at 1 dpi (p < 0.001) [305]. These findings indicate that ES accelerates the neurotrophic response, creating an early peak in motor neurons while subsequently downregulating pro-nociceptive factors in sensory neurons, which correlates with its enhanced regenerative and analgesic effects [305]. Goulart, Jürgensen et al. tested a combined therapy of SC transplantation (3 × 10^5^ cells) and treadmill training (TMT) in mice after sciatic nerve transection [306]. The combination (TMT + SC) yielded superior results, significantly increasing the number of myelinated fibers (2060 ± 92.10 vs. DMEM 1324 ± 279.9) and myelin area (8195 ± 1090 vs. 4628 ± 781). This group also showed accelerated motor recovery and the highest levels of neurotrophic factors (BDNF, NGF, neurotrophin-4) in the nerve and DRG. Furthermore, the TMT + SC and SC groups better protected spinal motor neurons (15.80 ± 2.69 and 14.20 ± 1.80 vs. DMEM 3.2 ± 0.91), while TMT alone was most effective for sensory neuron survival in the DRG (102.6 ± 4.73 vs. DMEM 61.0 ± 4.14) [301]. The TMT and SC transplantation strategies demonstrated distinct strengths [306]. The SC group was more effective at protecting spinal motor neurons from death, with a significantly higher neuron count (15.80 ± 2.69) compared to the DMEM control, a benefit not seen in the TMT group [306]. Conversely, the TMT group excelled at preserving sensory neurons in the dorsal root ganglia (102.6 ± 4.73), significantly outperforming the DMEM group, while the SC group’s effect (97.80 ± 10.96) was not statistically significant. Morphologically, both groups increased the number of blood vessels compared to controls, but the SC group, particularly when combined with TMT, led to superior nerve structure with higher numbers of myelinated fibers and myelin area [306]. The combined TMT+SC group ultimately achieved the best functional recovery, indicating that the therapies have complementary rather than redundant mechanisms [306]. In studies where exercise is added to invasive modalities (grafts, cell grafts, conduits, surgical repair), the combination almost always outperforms either alone in terms of functional recovery (motor/sensory), morphologic metrics (axon counts, myelin thickness), neurotrophic factor upregulation, and in some cases pain or neuropathic pain markers. E.g., the SC graft + exercise study shows accelerated motor recovery and greater histologic regeneration versus SC graft alone or exercise alone [306]. However, there is heterogeneity in how “exercise” is defined (timing of initiation, intensity, duration, modality), making comparisons hard. Some exercise protocols are more effective than others (e.g. intermittent compared to continuous treadmill running) in promoting both morphological and functional recovery [307]. Long-term outcomes (beyond early regeneration) are less well studied, especially in combination settings, and for large/gap injuries in larger animals or in human patients.
A number of representative published and notable ongoing clinical trials have been documented (Table 3). For example, Levy et al. recently initiated a clinical trial (NCT03011541) investigating the combined effects of autologous bone marrow-derived stem cells and surgical intervention for peripheral vision loss. Their earlier open-label, non-randomized Stem Cell Ophthalmology Treatment Study demonstrated visual acuity improvement in five of six patients with dominant optic atrophy following intravitreal, subretinal, retrobulbar, or subtenon BMSC administration, followed by IV infusion [309]. The therapeutic effects were hypothesized to involve mitochondrial transfer and exosome-mediated neuroprotection. Similarly, a 2015 case study (NCT01920867) involving a patient with relapsing optic neuritis and bilateral visual field deficits reported marked visual acuity gains 15 months after BMSC administration via optic nerve, retrobulbar, subtenon, and intravitreal routes [308]. Several additional stem cell-based trials for nerve repair are ongoing but unpublished, including VEGF165 plasmid gene therapy for PNI (Neovasculgen; NCT02352649), intraoperative autologous adipose-derived MSC therapy for revision nerve reconstruction (NCT04346680), bone marrow aspirate concentrate for segmental nerve defects up to 7 cm (NCT03964129), and a combination of allogeneic adipose-derived MSCs with human amniotic membrane to augment nerve transfer in traumatic brachial plexus injury (NCT04654286). In parallel, regenerative strategies are being explored for compression neuropathies such as carpal tunnel syndrome. While corticosteroid injections offer short-term symptom relief, an ongoing trial (NCT03722303) at Columbia University is evaluating autologous fat grafting as an alternative. The investigators hypothesize that the regenerative and angiogenic properties of adipose tissue may reduce fibrosis and enhance nerve gliding more effectively than corticosteroids. In addition to clinical trials, a case report described a 24-year-old male with traumatic radial nerve transection caused by a knife injury who underwent microsurgical repair using a sural nerve autograft supplemented with 1 mL of MSC-derived exosomes containing 5 billion microvesicles [113]. The exosomal suspension was partitioned into four 0.25 mL aliquots and administered subepineurially to both proximal and distal nerve stumps [113]. Electrophysiological evaluation at 6 months post-intervention revealed marked improvement in electromyographic activity, consistent with ongoing axonal regeneration [113].Table 3A summary of clinical trials on peripheral nerve injuriesTreatment methodsDiseasesDelivery methodEffects/aimsReference/ClinicalTrials.gov IDMSC-exosomes treatment after autograft repairA patient sustained a left radial nerve injury following a knife assault to the left armMicrosurgical intervention was performed on both the proximal and distal nerve stumps.Electrophysiological assessments at the 6-month follow-up revealed a marked enhancement in electromyographic signals, indicative of sustained neural regeneration。[113]Autologous bone marrow-derived stem cells (BMSC)A patient with relapsing optic neuritisBMSCs were administered via optic nerve injection in the right eye, and through retrobulbar, subtenon, and intravitreal routes in the left eye.The patient exhibited a marked improvement in visual acuity across both visual fields.NCT 01920867 [308];Physical exercise and sensory trainingNerve injuries in the arm and handPhysical exercise on a stationary bike, performed for 30 minutes twice weekly over six weeks, at a moderate intensity level (40–59% of VO2max).To investigate whether physical exercise and sensory training enhance sensory function and alleviate pain through upregulation of BDNF levels.NCT06846788Autologous bone marrow derived stem cells (BMSC)Neurologic disease and injuryBMSCs were administered intravenously, accompanied by topical application to the lower third of the nasal passages.To facilitate improvement in neurological function in individuals with specific neurological conditions.NCT02795052Autologous bone marrow-derived stem cells (BMSC) and surgical practicesRetinal and optic nerve damage or diseaseBMSCs were administered via retrobulbar, subtenon, and intravenous injections targeting one or both eyes.To evaluate the therapeutic effects of combining autologous BMSCs with surgical intervention in patients with peripheral vision loss.NCT03011541 [309];Autologous fatCarpal tunnel syndromeLipografting versus corticosteroid injectionsTo evaluate whether fat transfer more effectively prevents scar formation and facilitates nerve excursion within the canal—potentially through neoangiogenic and regenerative growth factors—compared to the standard steroid injection treatment.NCT03722303Electrical stimulationComplete digital nerve lacerationAt a fixed continuous frequency of 20 Hz.To determine the impact of pre-operative electrical stimulation on sensory nerve regeneration following surgical repair of traumatically transected digital nerves.NCT03205124Electrical stimulationCarpal tunnel syndrome surgeryStainless steel electrode wires were positioned adjacent to the median nerve, proximal to the surgical decompression site, to deliver immediate bipolar electrical stimulation at 20 Hz for 1 hour.Brief post-surgical electrical stimulation enhances axonal regeneration and restoration of muscle innervation, with no change in functional outcomes.[310]Electrical stimulationTraumatic PNIs with axon damage and clinical impairment of two musclesNot availableNo significant improvements were observed in clinical or neurophysiological assessments compared to the sham group.[311]External non-invasive peripheral nerve stimulationLongstanding moderate to severe neuropathic pain after peripheral nerve injuryAt a low frequency of 2 Hz.To assess the therapeutic efficacy and cost-effectiveness of external non-invasive peripheral nerve stimulation compared to control treatment for neuropathic pain.[312]Brief electrical stimulation (BES)Adult participants with a newly diagnosed head and neck cancer undergoing operative neck dissection.BES was applied continuously for 60 minutes at 20 Hz, 3–5 V, with 100-millisecond pulses following completion of neck dissection.To investigate whether brief electrical stimulation enhances postoperative clinical and objective shoulder function, as well as pain relief, following oncologic neck dissection.[313]Brief postsurgical electrical stimulationPatients with complete digital nerve transection received epineurial repair surgery.One hour of continuous electrical stimulation at 20 Hz.Compared to sham stimulation, postsurgical electrical stimulation consistently enhanced recovery across all sensory modalities at 5 to 6 months postoperatively.[314]Extracorporeal shock wave therapy (ESWT)Patients with nerve injury in hands and hypertrophic scarESWT was delivered with an energy flux density ranging from 0.05 to 0.30 mJ/mm^2^ at 4 Hz frequency, administering 1000–3000 impulses per treatment. A total of 12 weekly sessions were performed over 12 weeks.Beneficial effects on enhancing hand performance, facilitating scar recovery, and reducing scar-associated pain.[315]Low-intensity pulsed ultrasound (LIPUS)Carpal tunnel syndromeUltrasound was applied at an intensity of 0.3 W/cm^2^ and a frequency of 1 MHz for 20 minutes in pulsed mode with a 20% duty cycle, alongside a conventional physical therapy program.To examine the effects of LIPUS on pain levels, pinch grip strength, sensory and motor distal latency of the median nerve, and overall hand functionNCT05637684Low-intensity pulsed ultrasoundCarpal tunnel syndromeLow-, medium-, and high-dose groupsTo determine the effective therapeutic ultrasound dose for treating carpal tunnel syndrome by evaluating pain levels, functional ability, nerve conduction parameters, and pinch strength.NCT05863546Low-intensity pulsed ultrasoundCarpal tunnel syndromeTherapeutic ultrasound was applied over the carpal tunnel region using a handheld 5 cm^2^ transducer (BTL 5000 Combi, UK) at 1 MHz frequency, 1 W/cm^2^ intensity, and pulsed mode with a 4 duty cycle. Each session lasted 6 minutes with Aquasonic gel as a couplant. A total of 10 sessions were performed over a two-week period, with five sessions per week.To compare the efficacy of physical therapy modalities for carpal tunnel syndrome, including low-level laser therapy and ultrasound treatment.NCT03061149Recombinant Fibroblast Growth Factor-1 (ES135)Diagnosis of mild-to-moderate carpal tunnel syndromeSono-guided injection between carpal tunnel and median nerveTo investigate the efficacy and safety of ES135 as a treatment for carpal tunnel syndromeNCT06328166Placental growth factor or platelet-rich plasmaTraumatic nerve injuriesPerineural injectionTo investigate whether placental growth factor and platelet-rich plasma injections can promote recovery from peripheral nerve injury in adultsNCT06638008
ES has been more extensively investigated in clinical trials, with several studies reporting positive outcomes. A 2008 randomized controlled trial studied the impact of brief electrical stimulation on axonal regeneration after carpal tunnel release surgery. Researchers administered a single 1-hour session of 20 Hz bipolar stimulation using stainless steel electrodes placed next to the median nerve immediately after surgery. Axonal regeneration was measured by motor unit number estimation and nerve conduction studies. After 6–8 months, the stimulation group had increased motor unit number estimation values from 150 ± 62 to 290 ± 140 MU (p < 0.05); the control group showed no significant change (p > 0.2). Terminal motor latency improved in the stimulation group but not in controls, and sensory conduction recovered sooner in those who received stimulation [310]. Similarly, in eighteen patients with complete digital nerve transection treated with epineurial repair, postsurgical ES (20 Hz, 1 hour) led to significantly greater improvements in sensory function at 5–6 months postoperatively compared to sham stimulation [314]. For example, At 6 months, ES subjects reached a near-normal Cold Detection Threshold mean threshold of 14.33 ± 0.46 Just Noticeable Difference, compared with 17.22 ± 0.44 Just Noticeable Difference in controls [314]. Large fiber function was evaluated using static 2-point discrimination and Semmes–Weinstein monofilament testing. At the late stage, controls recovered to 8.69 ± 1.05 mm in static 2-point discrimination, whereas ES patients achieved near-normal 4.71 ± 0.90 mm. Similarly, Semmes–Weinstein monofilament testing thresholds approached normal in ES subjects (3.38 ± 0.12) compared with controls (3.91 ± 0.11) [314]. Functional disability was assessed with the DASH Questionnaire. At 6 months, controls averaged 19.42 ± 6.05, while ES subjects improved to 3.33 ± 1.21 (0 = normal). Two-way ANOVA showed significant treatment and follow-up effects. Late-stage recovery favored ES, but differences (p = 0.014, p = 0.027) lacked statistical significance, likely due to small sample size. [314]. However, outcomes across studies are not uniformly positive. A multicenter, double-blind, randomized clinical trial by Piccinini and Cuccagna et al. found that ES did not significantly improve clinical or neurophysiological outcomes in 38 patients with traumatic PNIs affecting at least two muscles and showing axonal damage [311]. More nuanced neuromodulation approaches, such as low-frequency transcutaneous magnetic stimulation (LFTMS) combined with transcutaneous electrical nerve stimulation (TENS), have shown complex physiological effects. Sham LFTMS plus TENS significantly reduced sensory conduction velocity and increased sensory onset latency and motor peak latency. In contrast, real LFTMS plus TENS reversed these effects and significantly improved sensory latency parameters (p = 0.036). LFTMS alone selectively modulated slow-conducting afferent fibers while reversing TENS-induced slowing in fast-conducting fibers, suggesting a potential peripheral modulatory role [316]. LFTMS alone significantly (p < 0.05) elevates sensory peak latency and onset-to-peak latency [316]. A single-site, blinded, randomized controlled trial (ISRCTN53432663), launched in 2016, evaluated the efficacy and cost-effectiveness of non-invasive peripheral nerve stimulation versus standard treatment for longstanding moderate-to-severe neuropathic pain in 76 patients with PNIs [312]. In another trial (NCT02268344), patients diagnosed with head and neck cancer undergoing oncologic neck dissection received intraoperative brief ES (BES) for 60 minutes at 20 Hz, 3–5 V, with 100 ms pulses. This study aims to assess whether intraoperative BES can improve postoperative shoulder function and reduce pain [313].
A double-blind, randomized controlled trial involving 120 patients demonstrated that ESWT significantly improved hand function, reduced hypertrophic scarring, and alleviated scar-related pain in individuals with hand nerve injury [315]. Treatment parameters included an energy flux density of 0.05–0.30 mJ/mm^2^, a frequency of 4 Hz, 1000–3000 impulses per session, administered once weekly over 12 weeks [315]. In comparison with the sham group (n = 51), the ESWT group (n = 52) demonstrated greater improvements across multiple outcomes. The ESWT group experienced a notable decrease in Visual Analogue Scale pain scores (5.50 ± 1.16) compared to the sham group (6.43 ± 1.42; p = 0.004). Extension range of motion in hand joints improved to −28.08 ± 57.74 versus −46.67 ± 81.81 (p = 0.02). On the Jebsen–Taylor Hand Function Test, ESWT patients showed superior performance in writing (11.31 ± 3.50 vs. 10.49 ± 5.29; p < 0.001), small object manipulation (10.79 ± 3.90 vs. 6.96 ± 4.34; p < 0.001), and light object handling (9.10 ± 2.61 vs. 8.80 ± 4.00; p = 0.002). Significant benefits were also observed in skin assessments, with reduced melanin content (163.42 ± 71.74 vs. 194.55 ± 134.49; p = 0.004), increased distensibility (0.54 ± 0.64 vs. 0.18 ± 0.15; p < 0.001), and enhanced biological skin elasticity (53.42 ± 21.76 vs. 37.07 ± 34.10; p < 0.001) [315]. Three completed clinical trials investigating LIPUS for carpal tunnel syndrome (CTS) were identified on ClinicalTrials.gov (NCT05637684, NCT05863546, NCT03061149). The first trial (NCT05637684) evaluated the effects of LIPUS on pain intensity, pinch grip strength, and sensory and motor distal latency of the median nerve in patients with chronic CTS. The second trial (NCT05863546) aimed to determine the optimal therapeutic ultrasound dosage by assessing its impact on pain, functional ability, nerve conduction, and pinch strength. The third trial (NCT03061149) compared the efficacy of physical therapy modalities, including low-level laser therapy and ultrasound, in the management of CTS.
Additional physical and neuromodulatory therapies are under investigation for PNI and related pain syndromes. For instance, mirror therapy is being evaluated in CTS patients in a registered clinical trial (NCT04087577), reflecting the growing interest in non-invasive neurorehabilitative interventions. Neuromodulation strategies such as peripheral nerve stimulation, spinal cord stimulation, DRG stimulation, and motor cortex stimulation are also being explored. A multi-site clinical trial (NCT06644807) is currently assessing the efficacy of implantable electrodes targeting these sites to alleviate chronic pain in patients with upper or lower limb amputation. Another trial (NCT06846788) seeks to evaluate the combined effects of physical exercise and sensory training on sensory recovery and pain reduction in patients with upper limb nerve injuries. This study is grounded in evidence that exercise enhances cognitive function and neuroplasticity via increased BDNF levels. However, the synergistic impact of exercise and sensory training on peripheral nerve recovery has not been previously tested.
Growth factor-based interventions are also entering clinical evaluation. A recent trial (NCT06328166) is investigating the safety and efficacy of recombinant human fibroblast growth factor-1 (rhFGF1, ES135) administered via ultrasound-guided perineural injection in CTS patients. Separately, a trial (NCT06638008) aims to compare the therapeutic potential of placental growth factor and platelet-rich plasma injections for PNIs. This study integrates neuromuscular ultrasound and nerve conduction studies to guide administration and monitor outcomes.
Collectively, the reviewed clinical trials underscore both the therapeutic promise and translational challenges of stem cell-based therapies and physical rehabilitation strategies for PNIs. While preclinical studies and early-phase trials have demonstrated encouraging signs of neuroregeneration and functional recovery, several barriers continue to hinder routine clinical implementation. First, axonal regeneration proceeds at a rate of approximately 1 mm/day, insufficient for long-gap or proximal injuries [317], often resulting in irreversible atrophy of target muscles and sensory receptors [111, 317]. Second, the heterogeneity of PNIs—from neurapraxia to complete neurotmesis—requires highly individualized treatment approaches [242]. The mixed sensory, motor, and autonomic composition of peripheral nerves further complicates targeted repair [318]. Third, although autologous nerve grafts remain the clinical gold standard, they are limited by donor site morbidity, restricted availability, and functional sacrifice [319]. Allografts offer an alternative but introduce immunological risks and necessitate immunosuppression [320]. Fourth, despite their potential, stem cell-based therapies yield variable clinical outcomes and face unresolved issues related to cell sourcing, delivery, standardization, and risks of immune rejection or tumorigenesis [118, 321, 322]. Exosome-based interventions are similarly constrained by uncertainties in dosage, timing, administration routes, and manufacturing complexity [321, 118]. Fifth, chronic inflammation and fibrotic scarring at the injury site remain significant biological impediments to axonal regrowth [323, 324]. Sixth, clinical trials are often limited by small cohorts, brief follow-up durations that fail to capture the extended timeline of nerve regeneration, and a lack of standardized outcome measures. Additionally, spontaneous recovery can confound treatment effects. Regulatory challenges—including high costs, long approval timelines, and the logistical burdens of manufacturing and quality control for biologics—further delay translation. Finally, although neuromodulation offers a promising adjunct, its efficacy depends on optimizing stimulation parameters and improving patient stratification.
Addressing the complex and multifactorial nature of PNI requires integrated therapeutic strategies that unify advances in biomaterials, cellular therapy, neuromodulation, and clinical translation [118]. To promote axonal regeneration, next-generation biomaterials are being engineered to provide both structural and biochemical guidance, incorporating aligned nanofibers, spatiotemporally controlled growth factor gradients, and ECM–mimetic scaffolds within bioactive nerve conduits [325]. For long-gap nerve defects, composite grafts combining autologous cells—such as SCs or MSCs—with synthetic or decellularized scaffolds represent a promising alternative to autografts and allografts, potentially mitigating donor site morbidity and immunogenicity [326–328]. Strategies aimed at accelerating axonal outgrowth and enhancing target-specific reinnervation increasingly rely on the sustained, localized delivery of NFs—including NGF, BDNF, and GDNF—as well as exosome-based therapeutics, now under refinement for improved potency and controlled release [329–331]. In parallel, stem cell approaches are advancing through optimized protocols for sourcing, purification, and preconditioning, with integration of gene-editing tools (e.g., CRISPR–Cas9) and real-time cell tracking technologies poised to enhance therapeutic precision, safety, and monitoring [332, 333].
Neuromodulation is emerging as a promising adjunctive strategy, leveraging activity-dependent plasticity to enhance functional recovery. Approaches such as BES, TENS, and DRG stimulation are under active investigation, alongside next-generation technologies including closed-loop systems and optogenetic interfaces that enable feedback-driven modulation of nerve repair [334–340]. To overcome post-injury barriers such as inflammation and fibrosis, immunomodulatory interventions—including anti-fibrotic agents, macrophage phenotype modulation, and engineered exosomes carrying anti-inflammatory cargo—are being actively explored [341–344]. In parallel, early and targeted rehabilitation remains essential. Task-specific motor training, mirror therapy, and sensory re-education aim to guide neuroplasticity and cortical reorganization, thereby maximizing functional outcomes [345–348]. Translational success will depend on rigorously designed, multicenter randomized controlled trials incorporating harmonized protocols, sensitive and standardized outcome measures—including electrophysiology, advanced neuroimaging (e.g., diffusion tensor imaging, magnetoneurography), and patient-reported outcomes—and extended longitudinal follow-up [349–351]. The identification of biomarkers, such as circulating neurofilaments and miRNA signatures, may enable real-time monitoring of regeneration and treatment stratification [352–355]. Ultimately, addressing regulatory and manufacturing challenges will necessitate collaborative efforts among researchers, industry stakeholders, and regulatory authorities. The deployment of GMP-compliant, scalable, and automated cell processing platforms—alongside decentralized manufacturing models—will be critical for ensuring equitable access to biologic therapies. The convergence of computational modeling, machine learning, and multi-omics profiling is poised to facilitate precision-targeted interventions, marking a paradigm shift in the management of peripheral nerve injuries.
Insights from previously published studies highlight the need for integrative, multi-omics approaches in PNI repair. Effective translation will require coordinated efforts across disciplines—including surgical innovation, pharmacological interventions, stem cell-based therapies, physical modalities, neuromodulation techniques, and biomaterial engineering. Collaboration among researchers, industry stakeholders, and regulatory agencies is essential to establish standardized therapeutic strategies and ensure quality control. The development of robust biomarkers for nerve regeneration and patient stratification will be critical. Furthermore, well-designed, double-blind, randomized controlled trials with adequate sample sizes are urgently needed, and their results should be made transparently available to guide clinical practice and inform the public [351–355]