Influencing factors and repair advancements in rodent models of peripheral nerve regeneration

ABSTRACT

Peripheral nerve injuries lead to severe functional impairments, with rodent models essential for studying regeneration. This review examines key factors affecting outcomes. Age-related declines, like reduced nerve fiber density and impaired axonal transport of vesicles, hinder recovery. Hormonal differences influence regeneration, with BDNF/trkB critical for testosterone and nerve growth factor for estrogen signaling pathways. Species and strain selection impact outcomes, with C57BL/6 mice and Sprague–Dawley rats exhibiting varying regenerative capacities. Injury models – crush for early regeneration, chronic constriction for neuropathic pain, stretch for traumatic elongation and transection for severe lacerations – provide insights into clinically relevant scenarios. Repair techniques, such as nerve grafts and conduits, show that autografts are the gold standard for gaps over 3 cm, with success influenced by graft type and diameter. Time course analysis highlights crucial early degeneration and regeneration phases within the first month, with functional recovery stabilizing by three to six months. Early intervention optimizes regeneration by reducing scar tissue formation, while later interventions focus on remyelination. Understanding these factors is vital for designing robust preclinical studies and translating research into effective clinical treatments for peripheral nerve injuries.

Plain language summary

Article highlights.

Aim

• To comprehensively review the critical factors influencing peripheral nerve regeneration outcomes in rodent models.

• To examine various injury models and repair techniques used in peripheral nerve research.

• To provide insights for designing robust and reliable studies that translate effectively to clinical applications.

Influencing factors

• Age: Age-related changes in neural tissue, such as decreased nerve fiber density and impaired axonal transport, impact regeneration outcomes.

• Sex: Sex differences influence nerve regeneration capabilities due to the hormonal activity of testosterone and estrogen, which play roles through pathways like BDNF/trkB and nerve growth factor signaling.

• Species and strains: Different strains exhibit varying baseline nerve densities and regeneration speeds, impacting study outcomes. Commonly used species include Sprague–Dawley, Lewis, Fischer, Wistar rats and C57BL/6 mice.

• Time course: Early intervention optimizes regeneration by reducing scar tissue formation, while later interventions focus on remyelination. Circadian rhythms modulate regenerative capacity and should, therefore, be considered.

Injury models

• Crush injury: The standard method to study axonotmesis involves mechanical compression, preserving nerve continuity.

• Chronic constriction injury: Models partial, persistent compression through loose ligatures around the nerve, often used to study neuropathic pain.

• Stretch injury: Simulates nerve elongation without severing axons, relevant for traumatic injuries.

• Nerve transection: Complete or partial severing of the nerve, requiring direct intervention for regeneration.

Repair strategies

• Direct nerve repair: End-to-end or end-to-side reattachment of severed nerves using sutures or collagen glues.

• Nerve grafts: The use of autologous grafts to bridge nerve gaps remains the gold standard for gaps >3 cm.

• nerve conduits: Biological and synthetic conduits provide a scaffold for regeneration and are evaluated for their effectiveness in bridging nerve gaps.

Future perspective

• Development of more sophisticated rodent models that better reflect human nerve injuries.

• Standardization of study variables to enhance comparability and generalizability.

• Advancements in biomaterials and scaffolds, focusing on biocompatibility, mechanical properties and functional integration.

• Integration of age, sex and species-specific data and temporal dynamics to optimize therapeutic strategies.

1. Introduction

Peripheral nerves possess a unique regenerative potential distinct from the central nervous system, allowing them to undergo repair via Wallerian degeneration and Schwann cell support. Understanding the molecular basis underlying these regenerative processes is essential for developing effective therapeutic options for nerve injuries. The complexity of peripheral nerve pathophysiology arises from the diverse physiological processes and etiologies associated with peripheral neuropathies, including those related to diabetes mellitus [1], corneal disease [2], chemotherapy [3] and trauma [4]. This complexity necessitates a thorough examination of various influencing factors that affect peripheral nerve regeneration outcomes in experimental models.

Using in vivo rodent models has been instrumental in advancing our understanding of peripheral nerve regeneration. These models are favored for their ability to provide comprehensive data related to sensorimotor function, electrophysiology, immunohistochemistry and radiographic imaging. Additionally, rodent models are adequate for direct nerve intervention and simulating various nerve injuries, offering a robust framework for evaluating novel therapeutic applications. However, the outcomes of these studies depend not only on the experimental interventions but are also significantly influenced by factors such as age, sex, species of rodent models and the specific injury models and repair techniques employed.

Given the wide range of methodologies available for peripheral nerve regeneration research [5], it is crucial to understand the influence of these factors to ensure the validity and reliability of experimental results. Age-related changes in neural tissues, sex-related differences in regenerative capabilities, and species-specific variations in nerve regeneration highlight the need for a nuanced approach to experimental design and interpretation. This review aims to provide a comprehensive examination of these critical factors and their impact on the outcomes of in vivo peripheral nerve regeneration studies, thereby offering insights that can guide future research and improve therapeutic strategies for peripheral nerve injuries.

2. General considerations

2.1. Age

During the aging process, a multitude of changes occur within rodent neural tissue [6–8]. As such, the age of the rodent model should be considered in the design and interpretation of peripheral nerve regeneration studies. For instance, age-related morphological changes can be observed in murine corneal nerves with decreasing nerve fiber densities occurring in C57BL/6J mice aged from six months to two years of age [9]. Furthermore, ultrastructural changes in rodent peripheral nerves can be observed in 24-month-old rodents compared with their younger 3-month-old counterparts. In a study on the oculomotor nerves of 9 rodents, older rats showed decreased collagen type I and IV immunoreactivity and NGF secretion, leading to a diminished capacity for regeneration and myelination [10]. This reduction in NGF secretion impacts the ability of nerves to repair and regenerate, ultimately affecting the success of therapeutic interventions aimed at promoting nerve healing. For example, estrogen has been found to enhance nerve regeneration in aged female rats by increasing NGF expression and promoting axonal growth and myelination [11]. Conversely, in younger male mice, the regenerative benefits on peripheral nerves by therapeutic intervention with testosterone appear more pronounced than in aged mice [12]. With that said, strength training paired with testosterone demonstrates protective capabilities against age-related declines in nerve function and sensitivity to the regenerative properties of testosterone in aged models [13].

Research on axonal transport in rodent sciatic nerve axons has found that the number of nicotinamide mononucleotide adenylyltransferase 2 (NMNAT2) vesicles decreases significantly between three and six months of age and again between 18 and 24 months old [14]. Similarly, significant reductions in the average and maximal transport velocities of NMNAT2 vesicles occur at 24 months [14]. These declines impact anterograde and retrograde transport, contributing to age-related axonal disorders. Impaired axonal transport may hinder the delivery of essential growth factors and cellular components necessary for nerve repair, thereby reducing the efficacy of regenerative therapies.

Other studies on rodent sciatic nerves have found the regression of sciatic nerve myelination to contribute to sarcopenia and balance dysfunction, observing significant regression across all fiber sizes and capillary architecture in the distal sciatic nerve to significantly occur with aging [15]. Interestingly, antioxidant treatments such as N-acetylcysteine (NAC) have been shown to improve nerve regeneration in aged rodents by reducing oxidative damage [16]. These findings may relate to how damaged peripheral nerves in aged mice clear debris more slowly than in younger mice [17]. Specifically, the greater the number of obstructions regenerating axons encounter, the slower neural regenerative potential becomes [17]. These findings demonstrate that neurophysiological reactions to injury depend upon the animal model’s age, with more mature animals demonstrating increased vulnerability to damage and impaired regeneration.

Some researchers have utilized aged rodents to observe the effect of senescence on the optic nerve, the development of glaucoma, and the regulation of neuropathic pain [18,19]. Additional studies evaluating the impact of age on peripheral nerves are limited, yet age can still be considered in the design of rodent experiments. Furthermore, a more unified approach in the age of rodent models utilized may allow for better comparability of results across different studies. Indeed, in a 2017 survey from Jackson et al., 297 researchers reported using rodents ranging from 6 to 20 weeks old, with additional findings showing that respondents believe the age range for adults ranges from 6–20 weeks [20]. This survey highlighted that the choice of rodent age is variable within studies, which may affect validity and generalizability.

2.2. Sex

Sex-related differences play a significant role in rodents’ outcomes of peripheral nerve regeneration studies. Initial observations have highlighted disparities in nerve regeneration capabilities between male and female rodents across various species and injury models. While the underlying mechanisms of these differences remain complex and multifaceted, the underlying signaling pathways may offer key targets for therapeutic pipelines.

Pham et al. (2019) reported enhanced corneal nerve regeneration in female mice across different strains in a sample of 36 mice [21], while other studies, such as Vacca et al. (2016), have shown male CD1 mice to demonstrate improved regeneration following sciatic nerve ligature injuries in a sample of 20 mice [22]. These findings suggest that sex may interact with species, nerve type, and other variables in ways that are currently difficult to elucidate due to the heterogeneity of study designs. Moreover, contrasting outcomes in axonal growth rates between male and female Wistar rats in response to different types of nerve injuries further complicate the interpretation of sex-related differences.

Recent studies have expanded our understanding by examining behavioral recovery, pain perception following nerve injury, and the role of sex hormones, providing a more nuanced view of sex influences in nerve regeneration. For instance, Michailidis et al. (2021) and others have demonstrated sex differences in depressive-like behavior and neuroimmune responses following peripheral nerve injury, suggesting that sex-specific neurobiological pathways influence behavioral recovery and pain perception [23].

Studies evaluating the effect of sex in rats are also challenging to interpret due to study design variation. The elongation rate of nociceptive axons in female Wistar rats demonstrates greater growth rates than that of male rats following sural nerve crush injury [24]. However, male Wistar rats have been shown to display better axonal growth than female Wistar rats with transection of the sciatic nerve [25]. Notably, the injury type in these two experiments is different. Other variables, such as animal age, should also be acknowledged and explored in the context of animal sex. Supporting this, aged female rats have been shown to remyelinate more efficiently than aged males but not in younger rats [26]. As testosterone levels naturally decline with age, this reduction is likely a contributing factor to the diminished regenerative potential observed in the nervous system of older mammals [27–30]. Thus, there may be a complex interplay among sex, species, and other variables potentially influencing nerve regeneration.

One significant factor contributing to sex-related differences in nerve regeneration is the activity of sex hormones [11]. Testosterone has been shown to enhance nerve regeneration in males, [12,31,32] with studies indicating that this effect is mediated through neural androgen receptors (AR) [33–35]. Androgen signaling has been linked to the regulation of BDNF and its receptor, trkB, in motoneurons [36]. BDNF/trkB signaling promotes local protein synthesis necessary for axonal regeneration through pathways like PI3K/Akt and MAPK/ERK [37,38]. The activation of these pathways leads to the enhancement of cytoskeletal proteins such as actin and tubulin, which are critical for growth cone formation and axonal elongation [39]. Additionally, BDNF/trkB signaling increases cAMP levels through the inhibition of phosphodiesterases, further promoting axonal growth and regeneration [40,41].

Similarly, estrogen has demonstrated beneficial effects on nerve regeneration, particularly in females. Estrogen promotes nerve regeneration by upregulating NGF expression, which in turn facilitates axonal growth and myelination through the ERK1/2 and JNK signaling pathways. These pathways are critical for promoting neuronal survival, axonal elongation, and remyelination after injury [11]. The administration of 17β-estradiol in female mice has been shown to improve neural recovery following sciatic nerve ligature injury, highlighting the potential of targeting estrogenic pathways as a therapeutic strategy [42].

Moreover, estrogen’s effects are mediated through its receptors, particularly estrogen receptor-β (ERβ), which has been shown to activate the PI3K/Akt pathway, leading to enhanced neuroprotection and repair [43]. Additionally, estrogen influences mitochondrial function and reduces oxidative stress, further contributing to its neuroprotective effects [11,22].

However, the role of sex hormones in neural repair may be subject to additional regulatory mechanisms [22,24]. Indeed, in ovariectomized female mice, Kovacic et al. found that female rats still demonstrated greater elongation distances of regenerating nociceptive axons – a finding that the authors attributed to more effective cell support in the distal nerve segment of females [24]. This suggests that while sex hormones like estrogen and testosterone are critical modulators, other factors, including cellular and molecular interactions within the nerve tissue, also play a vital role in the regenerative process.

Moreover, the extensive body of literature on sex-based differences in pain following nerve injury, as highlighted by landmark studies, emphasizes the necessity of considering these differences in the context of peripheral nerve regeneration research. The differential neuroimmune pathways contributing to neuropathic pain in males and females underscore the importance of incorporating a sex-based perspective in both the analysis of regenerative outcomes and the development of therapeutic strategies [44–47].

In summary, the influence of sex on peripheral nerve regeneration encompasses not only physical regeneration capabilities but also extends to behavioral recovery, pain perception and the modulatory role of sex hormones. The signaling pathways of testosterone (via AR, PI3K/Akt, MAPK/ERK) and estrogen (via ERβ, ERK1/2, JNK) offer promising therapeutic targets, particularly through their regulation of NGF expression and androgen receptor-mediated effects. Additionally, the BDNF/trkB pathway, influenced by androgen signaling, presents a potential avenue for enhancing nerve regeneration by promoting actin polymerization and growth cone dynamics. These factors underscore the necessity of a nuanced consideration of sex as a biological variable in the design, analysis, and interpretation of peripheral nerve regeneration studies. Future investigations are needed to fully flesh out the influence of sex hormones on nerve regeneration and their potential for therapeutic intervention.

2.3. Species

The selection of rodent strains is pivotal in determining the outcomes of peripheral nerve regeneration studies. Among commonly used strains, Sprague-Dawley, Lewis, Fischer, Wistar and Long Evans rats, along with C57BL/6 or BL6 mice, each exhibit unique regenerative characteristics. For example, BALB/c mice, known for having the highest baseline corneal nerve density, demonstrate faster nerve regeneration than CFW and C57BL/6 strains [21]. Similarly, Murphy Roths Large (MRL/MpJ) mice are recognized for their exceptional wound healing capabilities, which translate into significantly improved cutaneous nerve regeneration, a trait less pronounced in other strains [48,49]. Our investigations further corroborate these findings, showing enhanced remyelination in MRL/MpJ mice during sciatic nerve regeneration relative to C57BL/6 despite similar functional recovery [50].

Extending this inquiry to rat models, studies such as those conducted by Strasberg et al. (1999) and Steiner et al. (2019) have assessed regeneration across various rat strains with mixed findings, suggesting that the impact of strain on nerve regeneration may not be as pronounced in rats as in mice [51,52]. The inherent genetic diversity within strains like Sprague-Dawley suggests underexplored potential for regeneration studies [53]. For instance, Sprague–Dawley rats are often employed for studying general peripheral nerve injury due to their robust and reproducible responses, making them suitable for broad-spectrum PNI studies. Fischer rats, however, are particularly noted for their unique characteristics in morphology and response to neuropathic pain, making them more applicable in studies focusing on pain-related nerve regeneration mechanisms.

The importance of strain selection is further highlighted by studies from Webb and Muir in 2003 that used five strains with eleven rats each and Lovell et al. in 2000 that used four strains with six rats each, which demonstrated significant strain-dependent differences in morphology, sensory, motor, and neuropathic pain responses, particularly in Fischer rats [54,55]. These findings, coupled with the varying incidence of autotomy – a behavior linked to neuropathic pain – across different rat strains, emphasize the complexity of post injury pain mechanisms and the critical role of strain selection in the experimental design [56]. Studies have shown that different rat strains, such as Sprague–Dawley and Lewis, display divergent tendencies of autotomy behavior. This variation not only reflects genetic factors influencing neuropathic pain but also underscores the complexity of pain mechanisms following nerve injury [55–57].

To address the variability in regenerative outcomes and improve comparability across studies, Sprague–Dawley rats and C57BL/6 mice are recommended as standardized strains for future studies. These strains offer consistent and reproducible results in a variety of peripheral nerve injury models, making them ideal candidates for use in research aimed at advancing our understanding of peripheral nerve regeneration. As the field evolves, further large-scale, comparative studies are necessary to fully elucidate and leverage the influence of strain differences on regenerative outcomes.

2.4. Time course

The temporal dynamics of nerve injury and interventions significantly influence peripheral nerve regeneration outcomes. Circadian rhythms modulate biological processes in rodents, including nerve regeneration [58]. The timing of surgical interventions and treatments impacts regenerative efficacy due to diurnal fluctuations in hormone levels and metabolic activity [59,60]. Seasonal variations in temperature, light exposure and breeding cycles can also affect immune responses and tissue repair mechanisms, necessitating careful planning for long-term studies [60].

Recent research underscores the critical role of circadian rhythms in axonal regeneration. De Virgiliis et al. (2023) demonstrated that sensory neurons possess an intrinsic circadian clock, exhibiting diurnal oscillations in regenerative capacity in a murine sciatic nerve injury model [61]. Transcriptomic analyses revealed time-of-day-dependent enrichment in axonal regeneration and circadian clock pathways. Gene knockout experiments showed that Bmal1 is essential for intrinsic circadian regeneration and target reinnervation [58]. Specifically, Bmal1 deletion enhances axon regeneration by increasing diurnal Tet3-5hmC oscillations post-axotomy, elevating Tet3 expression, and resulting in global 5hmC gains, thereby impacting the DNA methylome and transcriptional output [46].

Lithium was included in these experiments due to its known influence on circadian rhythms and neuroprotective properties. Prior studies have demonstrated that lithium can modulate circadian gene expression and stabilize mood disorders, which are often linked to circadian disruptions [62,63]. In this scenario, lithium facilitated nerve regeneration in wild-type mice, but the effect was absent in mice lacking functional circadian clocks [61]. This suggests that pharmacological modulation of circadian pathways, such as through lithium, could offer novel therapeutic strategies for nerve repair.

The timing of post injury surgical procedures is also crucial. Immediate interventions can prevent inhibitory scar tissue formation and promote regeneration, while delayed repairs may require different therapeutic approaches [64]. Study duration is vital as well. Short-term studies focus on initial regenerative responses and inflammation, while long-term studies assess functional recovery, remyelination and nerve integration [65,66]. Standardizing follow-up durations enhances comparability across studies.

Finally, postoperative care can significantly influence outcomes, including analgesics, antibiotics and supportive measures. Consistent care protocols minimize variability and ensure animal well-being, directly impacting study results [67,68]. By considering these temporal factors, researchers can design more robust experiments to advance our understanding of peripheral nerve regeneration and develop effective therapies.

Key findings from studies on age-related changes, sex differences, species selection and time course effects are summarized in Table 2.

Table 2.

Summary of key findings for general considerations, injury models, and repair options for in vivo murine models of nerve regeneration.

Section	Key Findings	Ref.
2. General considerations
2.1. Age	Age-related changes in rodent neural tissue impact nerve regeneration. Morphological changes corneal nerves occur in C57BL/6J mice aged 6 months and 2 years. Two distinct phases of reduced axonal transport occur in rat sciatic nerves at 3 months and 18–24 months of age. Decreased collagen type I and IV immunoreactivity and NGF secretion in older rats. Estrogen enhances regeneration in aged female rats, while testosterone benefits younger mice. NAC improves regeneration in aged rodents by reducing oxidative damage. Damaged peripheral nerves in aged mice clear debris more slowly than in younger mice.	[6–20]
2.2. Sex	Significant sex-related differences in nerve regeneration. Female mice show enhanced corneal nerve regeneration. Male CD1 mice show improved regeneration following sciatic nerve ligature injuries. Sex hormones like estrogen and testosterone play crucial roles. BDNF/trkB critical for testosterone and NGF for estrogen signaling pathways. Behavioral recovery and pain perception also differ between sexes.	[21–47]
2.3. Species	Rodent strain affects peripheral nerve regeneration. BALB/c mice have the highest nerve density and regeneration speed, followed by CFW and C57BL/6. C57BL/6 Mice: Known for their robust regenerative responses, making them a standard choice for many studies. BALB/c Mice: Exhibit the highest baseline corneal nerve density and faster nerve regeneration compared with other strains like CFW and C57BL/6. Murphy Roths Large (MRL/MpJ) Mice: Renowned for exceptional wound healing capabilities, translating into significantly improved cutaneous nerve regeneration, a trait less pronounced in other strains. Sprague-Dawley Rats: Often used for general peripheral nerve injury studies due to their robust and reproducible regenerative responses. Fischer Rats: Noted for their unique characteristics in morphology and response to neuropathic pain, making them applicable in pain-related nerve regeneration studies. Different rat strains, such as Sprague-Dawley and Lewis, exhibit varying tendencies of autotomy behavior, a factor linked to neuropathic pain. Sprague-Dawley rats and C57BL/6 mice are recommended as standardized strains for future research. Mixed findings in rat strains; further investigation needed.	[21,48–57]
2.4 Time course	Sensory neurons possess an intrinsic circadian clock that influences their regenerative capacity. Diurnal oscillations in axonal regeneration are observed, with genes like Bmal1 playing a critical role. Bmal1 deletion enhances axon regeneration by modulating Tet3-5hmC oscillations, impacting the DNA methylome and transcriptional output. Immediate surgical interventions post-injury can prevent inhibitory scar tissue formation and promote optimal regeneration. Delayed repairs may require different therapeutic approaches, often focusing on remyelination rather than initial regenerative processes. Lithium enhances nerve regeneration in wild-type mice but not in clock-deficient counterparts. Pharmacological modulation of circadian pathways, like with lithium, could be a novel therapeutic strategy for nerve repair. The first month post-injury is crucial for degeneration and regeneration processes. Functional recovery typically stabilizes by three to six months post-injury.	[58–68]

3. Injury models
3.1. Crush injury	Mechanical compression of nerves studied via standardized “crush” methods. Extent of injury influences recovery; mild crushes recover faster. Standardized tools like non-serrated clamps and digital pressure devices improve reproducibility.	[69–82]
3.2. Chronic constriction injury	Reliable method to induce CCIs involves loose ligatures around the sciatic nerve. Ensures Wallerian degeneration and epineural inflammation, aiding in studying peripheral neuropathic pain.	[83,84]
3.3. Stretch injury	Stretch injuries characterized by rapid or slow stretches, affecting epineural and perineurial integrity. Reproducible methods developed for consistent damage.	[85,86]
3.4. Nerve transection	Involves complete or partial severing of nerves. Models reflect severe clinical outcomes like chronic denervation. Techniques secure proximal nerve stump to adjacent tissues to prevent spontaneous regeneration.	[87–93]

4. Repair options
4.1. Direct nerve repair	Direct reattachment using ETE or ETS repairs. Modern methods use fibrin glue to minimize gaps and improve regeneration.	[94–98]
4.2. Nerve graft	Autologous nerve grafts remain gold standard for gaps >3 cm. Factors like graft diameter, vascularity, and type influence outcomes. Motor nerves yield better outcomes than sensory nerves.	[99–116]
4.3. Nerve conduits	Biological and synthetic conduits explored to augment autografts. Tissue-engineered grafts combining autologous vein and nerve show promise. FDA-approved conduits available for gaps <3 cm.	[117–139]

3. Injury models

3.1. Crush injury

Mechanical compression of nerves is a standard method to study in vivo peripheral nerve regeneration. Compression aims to exert a uniform and standardized “crush” of the nerve, preserving the continuity of all neighboring axons and connective nerve fibers. Such an intervention results in axonotmesis – where the continuity of the nerve’s connective scaffold, particularly the epineurium, is preserved without disrupting the nerve trunk [69]. As a result, Wallerian degeneration and nerve regeneration may occur and be subsequently observed [70,71].

The extent of the crush injury plays a significant role in determining the nerve's recovery trajectory. Mild crush injuries typically result in quicker functional recovery and less extensive axonal damage, whereas severe crush injuries can cause substantial axonal degeneration, leading to prolonged recovery periods and incomplete functional restoration. For example, Bridge et al. (1994) demonstrated varying outcomes using different crush forceps, showing that more severe crushes resulted in delayed and less complete regeneration [72]. Similarly, Beer et al.’s (2001) development of a non serrated clamp provided insights into how standardized crush pressures influence recovery, with higher pressures correlating with more significant nerve damage and slower regeneration [73].

The historical perspective on the peripheral nerve crush injury (PNCI) reveals an evolution of techniques to induce reproducible and consistent injuries. Initiated in 1986 by De Koning et al., the methodology sought to achieve reproducible PNCIs to the sciatic and tibial nerves in rodents using consistent anatomical landmarks [74]. The introduction of specialized instruments [75–77] and methods further advanced this [78,79]. For instance, in 1994, Bridge et al. utilized #5 jewelers forceps to create a controlled nerve crush injury [72], marking a significant advancement in the field.

Beer et al.’s 2001 development of a non-serrated clamp designed to apply standardized pressure to the nerve represented a breakthrough, ensuring the reproducibility of crush injuries [73]. The non-serrated clamp also offers a reliable model for investigating nerve repair and regeneration across different animal species [73,80,81].

More recent developments have also improved PNCI methodology, such as the novel digital pressure measuring device introduced by Wandling et al. in 2021. This device allows for the real-time measurement of pressure applied to a nerve, significantly enhancing the reproducibility and accuracy of crush injuries [82].

3.2. Chronic constriction injury

A sciatic chronic constriction injury (CCI) is where the nerve is damaged or compressed in a manner where the compression is everlasting or partially remaining after the trauma event. Before CCIs were widely studied, it was difficult to consistently induce damage and conditions characteristic of these injuries. However, Bennet et al. created a standardized, reliable, reproducible method to induce CCIs. Three or four loose ligatures can be loosely tied around the sciatic nerve but tight enough to prevent sliding [83]. This allows for epineural blood flow to occur while persistently applying nerve compression. CCI models have been shown to help research peripheral neuropathic pain in mice [84]. Moreover, the method is reproducible and ensures induction of Wallerian degeneration, nerve compression and epineural inflammation.

3.3. Stretch injury

Another nerve injury often utilized in rodent models is the stretch injury, where the nerve is elastically stretched. This damages the nerve without compromising holistic axon continuity. Stretch injuries can be further characterized as either rapid or slow, resulting in varying degrees of epineural and perineurial ruptures [85]. Mahan et al. demonstrated that rapidly stretched nerves exhibit a zone of elastic recovery–beyond elastic recovery, inelastic rupture results. The authors highlighted that discrepancies in the findings of peripheral nerve studies may stem from differing testing conditions, including the speed at which nerve injuries occur [85]. The main challenge associated with the stretch injury in its early stages was maintaining a reproducible method of injury. However, a reproducible rapid-stretch nerve injury method was recently developed by Yeoh et al. in 2020 [86]. Based on biomechanical principles, using standardized weights, one-directional stretching, and uniform duration of applied stress, the method produces consistent nerve damage.

3.4. Nerve transection

Nerve transection involves a nerve’s partial or complete severing, thus eliminating axon continuity. One early nerve transection model was developed in 1979 by Wall et al. and employed a complete transection [87]. In this study, Wall et al. reported that nerve transection results in a phenomenon called “autonomy,” in which rodents attack the resultant anesthetic limb. To avoid the effects of autonomy, transection injury models should abstain from long-term studies. Following nerve transection, direct interventions are required for regeneration to occur. Various options are discussed below.

Transection lesions present significant challenges for translational research, primarily because most surgically relevant nerve lesions in humans involve at least partial transection or laceration, differing from experimental crush injuries in laboratory settings. In humans, spontaneous axon regeneration observed in laboratory animals is less common, often hindered by extensive fibrosis at the lesion site, necessitating surgical intervention to remove damaged tissue and facilitate repair through conduits [88].

An adequate experimental model of nerve transection that reflects human clinical scenarios is vital to mimic clinically relevant conditions. Chronic denervation represents one of the most severe clinical outcomes, where complete transection without subsequent reconstruction leads to a permanent disconnect between neurons and their distal targets, culminating in the loss of sensory and/or motor function. Such conditions often arise from severe traumas like brachial plexus injuries, resulting in multifaceted challenges, including the loss of motor function in the thigh’s posterior muscles and all muscles below the knee, alongside significant sensory function loss in the hindlimb and foot areas. This loss of sensory function can lead to progressive auto-mutilation in postoperative conditions [89–93].

To effectively study the impacts of chronic denervation and prevent spontaneous axonal regeneration in laboratory settings, special attention must be paid to securing the proximal nerve stump and suturing it to adjacent tissues, such as a muscle. This model is invaluable for investigating the denervation of the distal nerve trunk and skeletal muscles and identifying strategies to mitigate these effects.

Early post denervation changes and immediate and delayed effects on the nerve and muscle are pivotal areas of study within this model. Typically, a 1-month delay post denervation offers a substantial window to observe degeneration and atrophy within the affected nerve and muscle tissues. Long-term denervation studies often span three to 6 months post-injury, providing insights into the progression of degeneration and potential therapeutic interventions.

3.5. Comparison of models

The injury models discussed – crush, chronic constriction, stretch and transection – each offer unique advantages and limitations in peripheral nerve regeneration research. Crush injuries are highly reproducible and useful for studying early repair but may not capture the complexity of severe human injuries. Chronic constriction models are ideal for investigating chronic pain mechanisms but less relevant for acute regeneration. Stretch injury models, while simulating traumatic elongation seen in humans, can yield variable outcomes due to inconsistent stretching degrees. Transection models closely mimic severe clinical lacerations, essential for refining surgical techniques, though they can be challenging to standardize. Selecting the appropriate model based on specific research goals improves the relevance and translational value of findings.

A summary of the key findings from these injury models is provided in Table 2.

4. Repair options

4.1. Direct nerve repair

Nerve injuries may result in the laceration or complete separation of the nerve. The resulting neurotmesis can be treated by direct reattachment of the two severed nerves using end-to-end (ETE) nerve repair via sutures, collagen glues or grafts (Figure 1). In the event that the proximal axon stump is not available, end-to-side (ETS) nerve repair is a plausible strategy in the direct reconnection of severed nerves. Both methods involve direct suture repairs, where the nerves are directly reattached to one another. However, the traditional direct repair creates tension within the nerve, limiting optimal nerve regeneration [94,95]. As a result, early research sought to investigate methods to maximize regeneration in direct nerve repairs [96,97]. Instead of using sutures to reattach the nerves, a small epineural window can be opened directly and filled with a healthy nerve graft donor.

Figure 1.

Surgical exposure and direct repair of the sciatic nerve. (A) An incision is made over the postero-lateral aspect of the thigh. The tissue plane between the biceps femoris (*) and gluteus maximus (#) muscles is developed to expose the sciatic nerve (blue chevron). (B) The sciatic nerve is sharply divided. (C) A direct nerve repair is completed using two 10-0 nylon interrupted sutures (enlargement, black arrows).

More modern research seeks to improve upon direct repair methods. In 2020, researchers developed a novel nerve transection and repair method using fibrin glue that is consistent and reproducible without any further nerve manipulations or excessive axon loss [98]. Augmenting the transection with fibrin glue can achieve a minimal nerve gap, thereby improving subsequent nerve regeneration. The authors postulate that fibrin glue can act as a more standardized alternative to suture repair.

4.2. Nerve graft

Nerve grafting is typically employed when a nerve injury results in a gap that cannot be addressed through direct or tension-free repair. Research has demonstrated that attempts to repair nerves under tension often lead to suboptimal outcomes [99]. This is largely due to tissue ischemia caused by increased tension, which can block blood flow within the nerve, leading to necrosis and scar tissue formation that impairs axonal regeneration [100,101]. Using nerve grafts helps mitigate these challenges by providing a bridge across the gap, although it requires regenerating axons to traverse two neurorrhaphy sites [102].

Studies comparing direct nerve repairs performed under moderate tension with those using nerve grafts have shown comparable outcomes, but a lack of precise measurements for the applied tension complicates the interpretation of these results [102,103]. Various materials have been explored for bridging nerve gaps, but autologous nerve grafts remain the preferred method for gaps greater than 3 cm [104]. Since their first use in 1885 [105], advancements in understanding nerve anatomy and the development of microsurgical techniques have demonstrated that autologous grafts yield the most favorable outcomes [106–108]. These grafts provide an ideal environment for nerve regeneration, serving as a scaffold composed of Schwann cell basal laminae, neurotrophic factors and adhesion molecules, all of which play critical roles in promoting axonal growth [108–110].

Two key factors influencing axonal growth through the graft are the graft's diameter and its vascularization [109]. Initially, grafts rely on diffusion from surrounding tissues, but they must undergo neovascularization to maintain viability. Large-diameter grafts, however, may experience central necrosis, which can be replaced with scar tissue over time [111,112]. The type of nerve used in the graft is also important; motor nerves tend to produce better outcomes than sensory ones [113]. In some animal models, sensory grafts have even been shown to inhibit axonal regeneration [114], though in human clinical practice, sensory nerve grafting is commonly used with generally successful results. This discrepancy may be explained by the fact that motor nerves provide a more permissive environment for axonal growth, especially in cases where different types of axons (motor or sensory) are regenerating through the same graft [113,115,116]. If interactions or competition between different regenerating axon populations exist, successful regeneration then depends on the more spacious environment of motor nerve grafts based solely on architecture [113,114]. The larger diameter of human nerves thus supports the clinical practice of using sensory nerve grafts for reconstructing motor nerve defects with good clinical results [115].

4.3. Nerve conduits

While autografts are the standard in nerve graft repair, they can also result in partial loss of function, sensitivity in the donor region, and mismatched nerve diameter. Nerve conduits and scaffolding have been researched to augment autografts; however, they are often the subject of rejection by the host. Biological conduits, such as veins and arteries, have been widely researched for their use in peripheral nerve repair, especially for digital nerve injuries [117]. These vascular tissues and variations of skeletal muscle [118], mesothelial tissue [119] and epineural sheaths [120] have been employed to bridge nerve gaps. However, despite their biological compatibility, these tissues often show limited effectiveness in promoting nerve regeneration when compared with more sophisticated alternatives. Studies assessing the morphological outcomes of these conduits frequently reveal inferior results in terms of axonal regrowth and functional recovery [120].

Synthetic conduits, both degradable and nondegradable, present another approach. Early nondegradable options such as silicone and polytetrafluoroethylene tubes provided structural support but were hindered by inflammation, nerve compression and the necessity for a second surgery to remove the conduit, which increased patient morbidity [121–123]. In contrast, degradable synthetic conduits made from materials like collagen, polyglycolic acid (PGA), polylactic acid (PLA) and poly(lactide-co-glycolide) (PLGA) have gained traction due to their ability to support Schwann cell proliferation in large part by allowing sufficient oxygen and metabolite diffusion via pores while prohibiting the entry of fibroblast cells [124]. These materials have been effective in promoting nerve regeneration over short distances, and the combination with extracellular matrix components or neurotrophic factors can enhance their regenerative potential [125–129]. Recently, Wang et al. constructed a novel tissue-engineered nerve graft that combines autologous vein and nerve [130]. Veins have low immunogenicity and can also be used as nerve regeneration conduits.

Innovations in bioengineering have further refined nerve conduit design, incorporating tissue engineering techniques and support cells like Schwann cells or stem cells. These advancements have led to the development of bioabsorbable and compound conduits that provide structural support and promote chemotactic regeneration of peripheral nerves. For instance, biodegradable chitosan-collagen and collagen tubes have shown efficacy in promoting axonal growth, while more complex conduits incorporating Schwann cells or neurotrophic factors have demonstrated enhanced functional recovery in various models [131].

Several synthetic nerve conduits have received US FDA approval for clinical use, designed specifically for nerve gap repairs of up to three centimeters. These conduits, often used in peripheral nerve surgeries, serve to bridge the gap between damaged nerve endings and promote regeneration by providing a scaffold. FDA approved devices include a range of bioabsorbable and non absorbable conduits, as well as nerve-protectant wraps. Table 1 details some of these devices, offering insights into their material composition, clinical applications and outcomes in human nerve repair cases [132–136]. Nonetheless, recent studies indicate potential for expanding these conduits applications, particularly by incorporating advanced biomaterials and bioengineering strategies, which may eventually overcome current limitations. As shown in studies involving biodegradable poly(glycerol sebacate) (PGS) and poly(lactide-co-glycolide) conduits, the integration of these materials with support cells or neurotrophic factors offers promising avenues for enhancing nerve repair and functional recovery [131,137–139].

Table 1.

US FDA approved nerve guide conduits and absorbable nerve cuff/protectant wrap devices.

Product	Manufacturer	Composition	Gap (cm)	Degradation	Length (cm)	Diameter (mm)	510(k) ID	FDA clearance date
Nerve Guide Conduit Devices
NeuroTube™	Synovis Micro Companies Alliance, Inc.	Polyglycolic acid	2.0–4.0	3 months	2–4	2.3–8	K983007	22 March 1999/1995
NeuraGen™	Integra LifeSciences	Type I collagen	0.5–1.7	36–48 months	2–3	1.5–7	K011168	22 June 2001
Neuroflex™	Collagen Matrix, Inc.	Type I collagen	2.5–3.0	4–8 months	2.5	2–6	K012814	21 Sept 2001
NeuroMatrix™	Collagen Matrix, Inc.	Type I collagen	2.5–3.0	4–8 months	2.5	2–6	K012814	21 Sept 2001
Neurolac™	Polyganics, Inc.	Poly(dl-lactide-ϵ-caprolactone)	≤2.0	16 months	3	1.5–10	K032115; K050573	10 October, 2003 /4th May 2005
SaluTunnel™	Salumedica, LLC	Polyvinyl Alcohol	4.0–6.35	Non absorbable	6.35	2–10	K100382	5 August 2010
Reaxon™ Direct	Medovent GmbH	Chitosan	≤1.0	2.5 years	14	2.1–6.0	K143711	2015
NeuroCap™	Polyganics, Inc.	Poly(DL-lactide-caprolactone)	≤2.0	16 months	20	N/A	K171379	2017
Reaxon® Plus	Medovent GmbH	Chitosan	≤2.0	2.5 years	20	1.5–8.0	K181409	2018

Nerve Wrap Devices
SaluBridge™	Salumedica, LLC	Polyvinyl Alcohol	N/A	Non absorbable	6.35	2–10	K002098	24 November 2000/2001
AxoGuard™ Nerve Protector	Cook Biotech Products	Porcine small intestinal submucosa	N/A	3 months	2–4	2–10	K031069	15 May 2003
NeuraWrap™	Integra LifeSciences	Type I collagen	N/A	36–48 months	2–4	3–10	K041620	16 July 2004
NeuroMend™	Collagen Matrix, Inc.	Type I collagen	N/A	4–8 months	2.5–5	4–12	K060952	14 July 2006
Axoguard™ Nerve Cap	AxoGen, Inc.	Porcine small intestinal submucosa	N/A	4–8 months	1.5–7	5–10	K183366	2018

4.4. Combining different techniques

Combining different repair techniques can synergistically enhance nerve regeneration. For example, pairing nerve grafts with conduits can offer both structural integrity and biological cues critical for axonal growth. Utilizing fibrin glue in direct nerve repairs, when integrated with nerve conduits, can reduce tension at the repair site while ensuring proper alignment and stabilization of regenerating axons. Additionally, combining bioengineered conduits with growth factor-releasing scaffolds or Schwann cells can create an enriched regenerative environment, potentially overcoming the limitations of single techniques. This multimodal approach aims to optimize the regenerative process and improve the functional outcomes of nerve repair.

Table 2 summarizes the major findings on repair strategies, including direct repair, nerve grafts and conduits.

5. Conclusion

The field of peripheral nerve regeneration has seen significant advancements, especially through the use of rodent models. These models have provided crucial insights into nerve repair mechanisms and potential therapeutic strategies. However, there are still challenges in translating these findings into effective clinical treatments. Factors such as age, sex, species, time course, injury models and repair techniques all play essential roles in determining the outcomes of nerve regeneration studies. Future research can further refine strategies for nerve repair and improve functional recovery by addressing these variables and leveraging new technologies such as biomaterials, gene editing and circadian rhythm modulation. Standardizing experimental models and clinical trials will be essential in bridging the gap between preclinical findings and clinical applications, ultimately enhancing treatment efficacy for patients with peripheral nerve injuries.

6. Future perspective

The field of peripheral nerve regeneration has made significant strides, particularly with rodent models that have provided critical insights into the mechanisms and potential therapeutic strategies for nerve repair. Despite these advancements, several areas warrant further exploration to enhance our understanding and improve clinical outcomes.

One promising area for future research is the development of more sophisticated rodent models that better mimic human nerve injuries. Current models, while valuable, often do not fully replicate the complexity of human peripheral nerve damage. Advanced genetic engineering techniques, such as CRISPR-Cas9, could be utilized to create more accurate models that reflect the genetic and molecular intricacies of human neuropathies [140,141]. For example, incorporating circadian rhythm genes like Bmal1 to study their role in nerve regeneration can offer deeper insights, highlighting the potential for genetic tools, like CRISPR-Cas9, to reveal and modulate specific genetic pathways to study their involvement in nerve regeneration [58].

Additionally, incorporating comorbid conditions, such as diabetes or autoimmune disorders, into these models could provide investigators a more clinically representative model for investigating the roles and influences various conditions have on nerve regeneration [142–144]. For instance, diabetic neuropathy presents unique challenges in nerve repair due to prolonged inflammation and impaired blood flow, which are not typically present in standard rodent models [142].

The anatomical and functional variability within the peripheral nervous system also requires attention. Different nerves may exhibit distinct regenerative capacities due to variations in their anatomical structures, functions and local microenvironments. For example, sensory nerves often regenerate differently than motor nerves due to differences in their cellular compositions and regenerative demands [115,116]. Furthermore, nerve injuries in proximal regions (e.g., brachial plexus) often result in more complex repair challenges and different outcomes compared with distal injuries (e.g., digital nerves) due to the larger nerve gap and the need for more extensive tissue reconstruction [91–93].

The role of age and sex in nerve regeneration also presents a rich avenue for future investigation. While current studies have highlighted the impact of these factors, a deeper understanding of the underlying molecular and hormonal mechanisms is needed. Longitudinal studies that track regenerative processes across different life stages and between sexes could elucidate critical periods and conditions that optimize nerve repair [9,45,145,146]. Moreover, examining how age-related changes in the immune system influence regeneration could uncover novel therapeutic targets [8,147,148].

Another critical area for future research is the development and optimization of biomaterials and scaffolds for nerve repair. While autologous nerve grafts remain the gold standard, synthetic and bioengineered conduits hold great promise for overcoming the limitations of donor nerve availability and size mismatch [130,149]. Future studies should focus on enhancing the biocompatibility, mechanical properties and functional integration of these materials [150,151]. Incorporating growth factors, extracellular matrix components and cells into these scaffolds could significantly enhance their regenerative potential [133,152–154]. For example, the Brooke Army Medical Center conducted a clinical trial (NCT ID: NCT03964129), Phase I human safety study, to evaluate the sequential treatments of the decellularized peripheral nerve graft, Avance Nerve Graft with autologous bone marrow concentrate. However, achieving successful regeneration across gaps exceeding 3 cm continues to be a significant challenge, especially for patients with proximal nerve injuries. In such cases, biomaterial systems must ensure a precisely timed and spatial release of growth factors aligned with the rate of regeneration to prevent complications such as axonal trapping [155].

Another clinical trial is occurring at the University of Miami. Their active clinical trial (NCT ID: NCT05541250) aims to assess the safety and efficacy of autologous human Schwann cell augmentation in severe peripheral nerve injuries.

Ultimately, the translation of findings from rodent models to human clinical applications remains a significant challenge. Bridging this translational gap requires more sophisticated animal models and well-designed clinical trials that consider the complexities of human nerve injuries [156]. Collaboration between basic scientists, clinicians and engineers will be essential to developing and refining therapies that can effectively translate to patient care [157]. Additionally, developing clinically translatable drug delivery systems that provide controlled release of therapeutic agents during the entire nerve regeneration process is crucial. Biomaterials that emulate the three-dimensional structure of tissues incorporating surface topography, biochemical signals, and electrical properties, have shown significant potential in enhancing tissue regeneration [158,159]. However, these engineered drug delivery systems must ensure sustained and controlled drug release until functional reinnervation occurs to achieve clinical translation [155,160].

Author contributions

T Olsen: Investigation, writing – original draft, writing – review & editing. J LaGuardia: Conceptualization, investigation, writing – original draft, writing – review & editing, supervision, project administration. D Chen: Investigation, writing – original draft. R Lebens: Writing – original draft. K Huang: Investigation. D Milek: Conceptualization, writing – review & editing. M Noble: Conceptualization, writing – review & editing. J Leckenby: Conceptualization, resources, writing – review & editing, visualization, supervision, project administration.

Financial disclosure

This paper was not funded.

Competing interests disclosure

The authors have no relevant affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript. This includes employment, consultancies, honoraria, stock ownership or options, expert testimony, grants or patents received or pending, or royalties.

Writing disclosure

No writing assistance was utilized in the production of this manuscript.