Neurol India Home 

Year : 2019  |  Volume : 67  |  Issue : 7  |  Page : 100--105

Use of animal models in peripheral nerve surgery and research

Chandan B Mohanty1, Dhananjaya I Bhat2, B Indira Devi2,  
1 Department of Neurosurgery, Bombay Hospital Institute of Medical Sciences and BJ Wadia Children's Hospital, Mumbai, Maharashtra, India
2 Department of Neurosurgery, National Institute of Mental Health and Neurosciences, Bengaluru, Karnataka, India

Correspondence Address:
Dr. Dhananjaya I Bhat
Department of Neurosurgery, National Institute of Mental Health and Neurosciences, Hosur Road, Bengaluru - 560 029, Karnataka


Animal models are widely used in research of peripheral nerve injury and regeneration, since there are only minor differences in the anatomy of peripheral nerves and the physiology of nerve regeneration between the humans and animals. Animal models, especially rodents, are widely used for this purpose. This narrative review provides a brief overview of the role of animal models in peripheral nerve surgery and research.

How to cite this article:
Mohanty CB, Bhat DI, Devi B I. Use of animal models in peripheral nerve surgery and research.Neurol India 2019;67:100-105

How to cite this URL:
Mohanty CB, Bhat DI, Devi B I. Use of animal models in peripheral nerve surgery and research. Neurol India [serial online] 2019 [cited 2022 Jan 21 ];67:100-105
Available from:

Full Text

Animal models are popular in nerve surgery since the anatomy of peripheral nerves and physiology of nerve regeneration is similar in animals and humans. Hence, nerve experiments are easily replicable in an animal model. The smaller sized animals are generally preferred since the length-dependent nerve regeneration and end-organ reinnervation is faster in them compared to humans.[1],[2] Ensuring the safety and efficacy of newer treatment modalities in the pre-clinical animal studies is a necessary pre-requisite before they can be used in the clinical trials involving human beings. However, the direct extrapolation of the data of nerve experiments in animal models to humans has certain caveats and limitations. Nerve suturing exercises in animal models are also extremely useful for acquisition of microsurgical skills.

In this narrative review, we provide a broad overview of the animal models commonly used in nerve surgery and the current status of the role of animal models in the study of nerve injury and nerve regeneration.

 Types and Selection of Appropriate Animal Models

The animal models in nerve surgery can be divided into rodents and non-rodents. Rodents are characterized by the presence of growing incisors in the upper and lower jaw, which includes animals like rat, mice, guinea pig, hamster, etc. Non-rodent models in nerve surgery includes rabbit, monkey, horse, cat, dog, sheep, pig, etc., The ease of availability, the lower degree of care and maintenance cost, the societal acceptance, the tolerance to captivity and relative resistance to post-surgical infections have made rat, the most widely used animal model in nerve surgery.[3],[4],[5] Larger animals like monkey, horse, and dog are difficult to procure and expensive to transport and maintain, and hence infrequently used. The commonly used rat species are Wistar, Sprague-Dawley, Lewis, etc.

Sciatic nerve is the most commonly used nerve to perform nerve experiments,[3],[6],[7] being the largest nerve, which makes its handling and repair easier.[8] The other commonly used nerves in the rat are peroneal, tibial, radial, median and ulnar nerves. Nerves of the forelimb are uncommonly used due to their smaller size, especially in smaller animals like the mice, since advanced microsurgical skills are required to handle and repair these nerves.[7] Selection of the correct 'lesion' is based on the study goals. Generally, the 'axonotomesis model' is selected to study the biology of the nerve regeneration and to test new pharmacological and therapeutic agents especially in neuropathic pain.[7] The 'neurotmesis model' is generally used in pre-clinical studies of therapeutic interventions since differences in various outcome assessment parameters will help in proving the efficacy of the intervention.[7] The superiority of either the forelimb nerve to the hindlimb nerve, or vice-versa, has not been unequivocally established.[7] Most of the experiments are performed on mixed nerves; however depending on the clinical target, a pure motor, pure sensory or an autonomic nerve can also be selected. Autonomic nerves are generally selected for urological studies.[7] A sensory nerve has less efficient nerve regeneration compared to a motor nerve.[9]

 Use of Animals in Peripheral Nerve Research

Promotion of nerve regeneration

The best possible results following surgical repair of the nerve have reached a plateau in spite of improving surgical techniques. Hence, the aim of majority of the nerve studies in animals is to improve peripheral nerve regeneration following nerve repair. The study of nerve regeneration in humans and larger animals, especially primates, is marred by the slow rate of growth of the regenerating nerve and the cost and ethical considerations. Rodent models are preferred for this reason. Sciatic nerve transection in the rat, followed by an end-to-end repair using a nerve graft or nerve conduit, is the most commonly used model to study nerve regeneration.[10] Improvement of nerve regeneration can be achieved by employing one step or a combination of the following steps.[11]

a. Promotion of axonal growth

Local use of growth factors like nerve growth factor (NGF), glial growth factor (GGF), fibroblast growth factor (FGF), glial cell derived neurotrophic factor (GDNF), neurotrophin 3 (NT-3), ciliary neurotrophic factor, leupeptin and brain derived nerve growth factor (BDNF) at the proximal stump have been described to promote axonal regeneration.[12],[13],[14],[15]

Brief electrical stimulation of the proximal stump is also known to increase the expression of neurotrophic factors like BDNF and NGF in the Schwann cells and also upregulates the neuronal regeneration associated genes like BDNF and tyrosine receptor kinase B (trkB).[16],[17]

Application of gene therapy in improving peripheral nerve regeneration is an area of intense research. Gene therapy involves successful delivery of genes to the site of nerve injury to promote axonal regeneration using viral vectors. Adeno-associated viral vectors are the most commonly used viral vectors due to their low mutagenicity and immunogenicity.[10],[18] The key role played by Schwann cells in the region of the regenerating axon is well recognized and most of the gene therapy is targeted towards maintenance and sustenance of these Schwann cells using lentivirus.[10],[15],[19] Improved gait recovery, sensory recovery, and electrophysiological recovery have been reported following regulatable gene therapy in sciatic nerve models in rats.[15],[20] Regulatable gene therapy vector is essential to prevent excessive production of neurotrophic factors, to avoid local trapping of the regenerating axons, and also to prevent side effects like hypersensitivity to the growth factors. Regulatable gene therapy has an “on-off” mechanism for local GDNF production using extrinsic agents like doxycycline.[15],[20]

Schwann cells are known to be the most important cells for promotion and maintenance of regenerating axons. However, a direct Schwann cell implantation at the repair site is impractical since Schwann cells are difficult to harvest, slow to grow, difficult to isolate, and have a high immunogenicity. Hence, stem cells with the ability to transform into Schwann cells at the site of regeneration are being intensively studied. Some of the sources of these stem cells are neural stem cells, mesenchymal stem cells, fetal derived stem cells (umbilical cord, Wharton's jelly, amniotic tissue), skin derived precursor stem cells, hair follicle stem cells, dental pulp stem cells and induced pluripotent stem cells.[21],[22],[23],[24],[25] Stem cell transplantation is usually achieved by microinjection at the site of regeneration or into the conduit lumen.[21] The three-dimensional printing technique has also been used to prepare a scaffold with multiple varieties of cell types to mimic the native micro-environment.[26] The mechanism of action of stem cells is influenced by their differentiation into Schwann-type cell, by an increase in the local production of neurotrophic factors, and by an enhanced myelination of the regenerating axon.[21] Use of stem cells in the rat sciatic nerve model, or the monkey median nerve model have shown functional, histological, histomorphometric, and electrophysiological recovery better than that obtained using the current microsurgical repair techniques.[20],[22],[27]

b. Local milieu conducive to regenerating axons

Limited availability of autografts, especially in polytrauma patients, has made the use of artificial nerve conduits essential. However, in addition to its role as a graft, the use of artificial nerve conduits should allow diffusion of oxygen as well as nutrients inside the graft and should also prevent fibroblast infiltration into the graft.[28] Use of artificial conduits has been extensively studied in animals and the safety and efficacy of the process has been well established. This has led to the use of artificial nerve conduits in humans.

In addition to nerve conduits, use of amniotic membrane, fibrin glue and Nd-YAG laser to seal the site of coaptation have shown that these measures separate the regenerating axons from the surrounding tissue leading to an increase in the local neurotrophic factors. It also prevents the entry of fibroblasts and inflammatory cells into the coaptation site, thus preventing perineural scarring and fibrosis and leading to an improved nerve regeneration. A biomechanical study on sciatic nerve coaptation of rabbits had shown that the use of fibrin glue alone had a poorer tensile strength compared to the repair with sutures.[29] However, a systematic review by Sameem et al., (included human and animal studies) revealed that fibrin glue has at least equal efficacy as suture repair, if not being superior to it.[30]

c. Delaying or preventing the occurrence of Wallerian degeneration

Prevention of Wallerian degeneration would mean that the regenerating axons need to travel for a shorter distance to reach the distal stump, leading to improved outcomes. Invertebrates like earthworm and crayfish are able to delay or completely prevent Wallerian degeneration, unlike the situation in mammals.[31] Advances in the field of microsurgery, cell biology, biochemical and biomolecular engineering have enabled axonal membrane fusion using polyethylene glycol (PEG) of the severed proximal and distal ends of the axonal membrane. This technique has been extensively researched starting in earthworms, followed by in vitro and subsequently in vivo experimentation in rats, rabbits and guinea pigs.[32] Extensive research on the rat sciatic nerve model has shown improved behavioral recovery, electrophysiological recovery, and survival of the distal segment using PEG assisted axonal fusion. The exact mechanism of action of technique is still unknown; however, the reports do indicate recovery of function within days to weeks, which is much faster compared to any other technique.[32]

d. Reduction of duration of muscle denervation

Chronic denervation leads to fibrosis, muscle atrophy and endplate changes, which leads to a poor neurological recovery. Williams, using continuous electrical stimulation of the denervated muscle by an implantable electrical stimulator in rabbits and dogs, showed that following microsurgical nerve repair, these animals showed a good functional recovery in the denervated muscles.[33] The author also noted a good tolerability of the implanted device in the animals.

Study of neural plasticity

Neural plasticity plays an important role in improving the overall outcome after nerve repair.[34],[35],[36] However, there still exist numerous knowledge gaps in the understanding of pathophysiology of neural plasticity, where the use of animal models is critical. Interestingly, in the study of neural plasticity, the upper extremity nerves in monkeys are commonly used as experimental models, probably due to their closer resemblance to humans and their ability to carry out fine distal motor movements.[36],[37] Functional magnetic resonance imaging (fMRI) and diffusion tensor imaging (DTI) are usually used to record the changes in brain plasticity following nerve injury, where the intra- and interhemispheric cortical map expansion can be studied and are correlated with the degree of axonal regeneration in the injured nerve. Maladaptive neural plasticity following nerve injury has also been studied extensively in rat models. Long-term potentiation in the dorsal horn of the spinal cord by activation of N-methyl D-aspartate receptors leading to hyperalgesia and allodynia even after complete nerve recovery has been shown in numerous rat models.[38]

Study of neuropathic pain

Neuropathic pain has been described as the 'most terrible of all tortures.'[39] Replication of neuropathic pain in humans may produce irreversible nerve damage and hence the process is not performed in humans.[39] Rat is the most commonly used animal for experiments. Neuropathic pain can be due to numerous causes, hence numerous models have been described to replicate the clinical scenario in animals. These include the complete nerve transection model, chronic constriction injury model using ligatures around the nerve, the partial sciatic nerve ligation model, spared nerve injury model (tibial, peroneal nerve is completely transected while the sural nerve is left intact), sciatic nerve cryoneurolysis (the sciatic nerve is frozen to -60 C with a cryoprobe), sciatic inflammatory neuritis (injection of tumour necrosis factor alpha or zymosan or seaweed protein to induce neuritis), laser induced sciatic nerve injury, photochemical induced nerve injury, various drug induced neuropathy models, diabetic neuropathy model (injection of pancreatic beta cell toxins like streptozocin), and post-herpetic neuralgia model (injection of varicella zoster virus in rat paw.[39] These models have been studied to test the pharmacological efficacy of new drugs to treat neuropathic pain.

Skills training

Animal models are an imperative part of skills training laboratory. Sciatic nerve end-to-end repair in rats is one of the most common exercises to gain skills in microneurosurgery. The sciatic nerve is usually exposed in the standard fashion in the rat hind limb by splitting the hamstring muscles. A clean nerve transection is performed and the nerve coaptation is performed using 9-0 or 10-0 polypropylene epineurial sutures in a tensionless manner. An end-to-side coaptation can also be performed by dissecting the sciatic nerve more distally till its trifurcation to the peroneal, tibial and sural branches is seen. One of these branches can then be transected, which can be looped and anastomosed to one of the other branches in an end-to-side fashion.

The candidate learns to handle the micro-instruments, gains familiarity with the use of microscope and the effective use of zoom-in and zoom-out function, tie knots under high magnification, and learns the nuances of securing the knot with the necessary tension without damaging the neural tissue. Thus, the use of rat model provides one of the best and time-tested models for advanced microneurosurgical skill acquisition.[40]

 Ethics in Animal Experiments

The International Committee for Laboratory Animal Science (ICLAS) has laid down clear guidelines for the experimentation and the necessary training of the researchers involved in animal handling.[41]

The three 'R's of use of animals in experimental studies is based on the principles of: “Replacement” (avoid or replace the use of animals in experiments by suitable in vitro models or simulation models or other animals like invertebrates or nematodes); “Reduction” (Reduce the number of animals to be used in the experiments by modifying the methodology, as well as sharing of data and results); and “Refinement” (refine the procedure to minimize the pain and suffering of the animals). Committee for the Purpose of Control and Supervision of Experiments on Animals (CPCSEA), the nodal regulatory body in India for the use of experimental animals added the 4th 'R' namely “Rehabilitation” (rehabilitation and after-care of animals after the experiment, with the researcher having a moral responsibility towards the animals) emphasizing the concept of 'Ahimsa' (philosophy of non-violence) reflecting the religious, cultural and social traditions of India.[42] Rehabilitation of experimental animals is a legal requirement in India.

 Outcome Assessment

Outcome assessment is important to correctly document the degree of nerve regeneration making the experimental groups comparable to one another.

Functional assessment

Motor functional assessment is one of the most common methods to assess outcome. Gait can be analyzed by numerous parameters like sciatic functional index, step length ratio, walking track analysis, ladder rung analysis, Basso-Beattie-Bresnahan score, and computerized gait analysis (Cat Walk system).[43] Forelimb motor function can be assessed by grasping test, Irvine-Beatties-Bresnahan (IBB) forelimb scale, and staircase test etc.[7],[44],[45]

Sensory function assessment can be performed by using thermal sensitivity testing and Von-Frey test.[20],[46]


Electrophysiological assessment using electromyography (EMG) and compound motor action potential (CMAP) helps in assessment of the nerve function.[20],[47],[48] CMAPs that are recorded proximal and distal to the site of nerve injury, help to assess the functional nerve recovery.

Histology and histomorphometry

Histological and histomorphometric analysis of the nerve segment complement the functional and electrophysiological assessment. The histological assessment can identify foreign body reactions and granulomas, if foreign substances like artificial grafts are used. It is also extremely useful in assessing the number of regenerating axons, the presence and thickness of the myelin sheath, perineural adhesions and fibrosis.[43]


In vivo imaging techniques like ultrasonography and MRI are being recently investigated as potential investigations to assess nerve regeneration. Ultrasonography allows the assessment of the regenerating axon within an artificial nerve conduit.[43],[49] However, the cost incurred and the technical difficulties encountered in obtaining useful images in small animals remains a challenge.

Retrograde labelling

Retrograde labelling of the neurons in the spinal cord of the rat or mouse is another method to assess nerve regeneration. This involves the use of a tracer like fast blue or fluro gold or fluoro ruby (which is introduced by an intramuscular injection, or by application of the crystal, or by conduit reservoir where the proximal end of the transected nerve is placed in a conduit which acts as a reservoir of the dye). After this procedure, the stained neurons in the spinal cord are counted. Thus, retrograde labelling provides a quantitative proof of reconnection between the nerve distal to the injury site and the spinal cord.[50] Retrograde labelling is also useful to determine the quantity of the misdirected regenerating axons which may prevent a functional recovery.[51] The femoral nerve model in rats is commonly used to study retrograde labelling.[51]

 Limitations and Future Directions

The success gained in the animal models of nerve regeneration have not progressed to the clinical settings. The main limitations that hamper the use of animal models in peripheral nerve research is the inability to directly extrapolate data from the animal studies to humans due to the subtle physiological differences in peripheral nerve regeneration in animals and humans. The primate physiology more closely resembles that of humans. Due to the cost of procurement, transportation, maintenance and ethical considerations involved in the use of primates, any experimentation on them is usually avoided. Another limitation of the animal studies is the relatively short duration of the experimental study, due to which the long-term animal data is lacking. Majority of the experiments have a follow-up data of less than 6 months. Stem cells in animals have been used extensively with proven efficacy in improving peripheral nerve regeneration; however, the long-term effects of using pluripotent stem cells or the transduced viruses are still unknown. There is also a lack of consensus on a standardized outcome assessment to measure nerve recovery due to the varied parameters employed by the researchers. Animal experiments are generally performed on very young animals who are free of other diseases, which does not match with the clinical scenario. Hence, some researchers have performed experiments with older animals and animals with diseases like diabetes, infections etc.[52],[53] Most of the animal studies in nerve regeneration use a nerve transection and coaptation model with a nerve gap of less than 1 cm in an acute setting.[3] Delayed nerve repair models in animals have been infrequently described.[22] However, most of nerve injuries in the clinical setting have a delayed presentation, longer nerve gaps and a clean transection rarely occurs in nature. Stretch, laceration, neuroma formation or a combination of all these modes of injury are seen in the clinical settings; however, they are infrequently described in the animal models.[5],[54] Hence, the remarkable results of nerve regeneration in the animal models must be interpreted with these limitations in mind.

The successful use of artificial nerve conduits in animals have resulted in the wide-spread use of these products in human nerve injuries.[3] The safety and efficacy of fibrin glue was proven in animal models and was subsequently adopted in humans. Similarly, following the successful implementation of electrical stimulation in the rodent model, this technique has also been introduced in humans. In this method, a brief electrical stimulation of the median nerve following carpal tunnel release surgery showed a significant improvement in the axonal regeneration and target reinnervation.[55] Use of transgenic animals (mouse, rat) is another exciting development in the nerve research. The utilization of transgenic animals involves the insertion of foreign deoxyribose nucleic acid (DNA) into the host chromosome and thus the animal is able to phenotypically mimic the disease or the condition to be studied.[56] Transgenic animals have been instrumental in improving our understanding of the biology of nerve regeneration by overexpression or deletion of certain proteins or genes at the target site. Transgenic animals have also been genetically altered to express fluorescent proteins at the site of regenerating axons to allow the direct visualization and tracking of nerve regeneration in live animals.[57],[58],[59]

Standardization of the animals, animal strains, nerve injury models and the outcome assessment criteria of nerve regeneration and recovery is critical to make the animal experimental data comparable to each other. This will result in an enhanced translation of various therapies into the clinical settings.

Financial support and sponsorship


Conflicts of interest

There are no conflicts of interest.


1Dolenc V, Janko M. Nerve regeneration following primary repair. Acta Neurochir (Wien) 1976;34:223-4.
2Buchthal F, Kuhl V. Nerve conduction, tactile sensibility, and the electromyogram after suture or compression of peripheral nerve: A longitudinal study in man. J Neurol Neurosurg Psychiatry 1979;42:436-51.
3Angius D, Wang H, Spinner RJ, Gutierrez-Cotto Y, Yaszemski MJ, Windebank AJ. A systematic review of animal models used to study nerve regeneration in tissue-engineered scaffolds. Biomaterials. 2012;33:8034-9.
4Schimandle JH, Boden SD. The use of animal models to study spinal fusion. Spine (Phila Pa 1976) 1994;19:1998-2006.
5Toia F, Giesen T, Giovanoli P, Calcagni M. A systematic review of animal models for experimental neuroma. J Plast Reconstr Aesthet Surg 2015;68:1447-63.
6Griffin JW, Pan B, Polley MA, Hoffman PN, Farah MH. Measuring nerve regeneration in the mouse. Exp Neurol 2010;223:60-71.
7Tos P, Ronchi G, Papalia I, Sallen V, Legagneux J, Geuna S, et al. Chapter 4: Methods and protocols in peripheral nerve regeneration experimental research: Part I-experimental models. Int Rev Neurobiol 2009;87:47-79.
8Varejão AS, Melo-Pinto P, Meek MF, Filipe VM, Bulas-Cruz J. Methods for the experimental functional assessment of rat sciatic nerve regeneration. Neurol Res 2004;26:186-94.
9Moradzadeh A, Borschel GH, Luciano JP, Whitlock EL, Hayashi A, Hunter DA, et al. The impact of motor and sensory nerve architecture on nerve regeneration. Exp Neurol 2008;212:370-6.
10Hoyng SA, de Winter F, Tannemaat MR, Blits B, Malessy MJ, Verhaagen J. Gene therapy and peripheral nerve repair: A perspective. Frontiers in Molecular Neuroscience 2015;8:32.
11Grinsell D, Keating CP. Peripheral nerve reconstruction after injury: A review of clinical and experimental therapies. BioMed Research International 2014;2014:698256.
12Konofaos P, ver Halen JP. Nerve repair by means of tubulization: Past, present, future. J Reconstructive Microsurg 2013;29:149-64.
13Lee SK, Wolfe SW. Peripheral nerve injury and repair. J Am Acad Orthop Surg 2000;8:243-52.
14Kemp SW, Walsh SK, Midha R. Growth factor and stem cell enhanced conduits in peripheral nerve regeneration and repair. Neurol Res 2008;30:1030-8.
15Shakhbazau A, Mohanty C, Shcharbin D, Bryszewska M, Caminade AM, Majoral JP, et al. Doxycycline-regulated GDNF expression promotes axonal regeneration and functional recovery in transected peripheral nerve. J Control Release 2013;172:841-51.
16Khuong HT, Midha R. Advances in nerve repair. Curr Neurol Neurosci Rep 2013;13:322.
17Gordon T, English AW. Strategies to promote peripheral nerve regeneration: Electrical stimulation and/or exercise. Eur J Neurosci 2016;43:336-50.
18Hastie E, Samulski RJ. Adeno-associated virus at 50: A golden anniversary of discovery, research and gene therapy success-a personal perspective. Hum Gene Ther 2015;26:257-65.
19Shakhbazau A, Kawasoe J, Hoyng SA, Kumar R, Van MJ, Verhaagen J, et al. Early regenerative effects of NGF-transduced Schwann cells in peripheral nerve repair. Mol Cell Neurosci 2012;50:103-12.
20Shakhbazau A, Mohanty C, Kumar R, Midha R. Sensory recovery after cell therapy in peripheral nerve repair: effects of naïve and skin precursor-derived Schwann cells. J Neurosurg 2014;121:423-31.
21Jiang L, Jones S, Jia X. Stem cell transplantation for peripheral nerve regeneration: Current options and opportunities. Int J Mol Sci 2017;18:94.
22Khuong HT, Kumar R, Senjaya F, Grochmal J, Ivanovic A, Shakhbazau A, et al. Skin derived precursor Schwann cells improve behavioral recovery for acute and delayed nerve repair. Exp Neurol 2014;254:168-79.
23Raheja A, Suri V, Suri A, Sarkar C, Srivastava A, Mohanty S, et al. Dose-dependent facilitation of peripheral nerve regeneration by bone marrow-derived mononuclear cells: A randomized controlled study: Laboratory investigation. J. Neurosurg 2012;117:1170-81.
24Uemura T, Ikeda M, Takamatsu K, Yokoi T, Okada M, Nakamura H. Long-term efficacy and safety outcomes of transplantation of induced pluripotent stem cell-derived neurospheres with bioabsorbable nerve conduits for peripheral nerve regeneration in mice. Cells Tissues Organs 2014;200:78-91.
25Zarbakhsh S, Goudarzi N, Shirmohammadi M, Safari M. Histological study of bone marrow and umbilical cord stromal cell transplantation in regenerating rat peripheral nerve. Cell J 2016;17:668-77.
26Johnson BN, Jia X. 3D printed nerve guidance channels: Computer-aided control of geometry, physical cues, biological supplements and gradients. Neural Regen Res 2016;11:1568-9.
27Hu N, Wu H, Xue C, Gong Y, Wu J, Xiao Z, et al. Long-term outcome of the repair of 50 mm long median nerve defects in rhesus monkeys with marrow mesenchymal stem cells-containing, chitosan-based tissue engineered nerve grafts. Biomaterials 2013;34:100-11.
28Chang C, Hsu S, Yen H, Chang H. Effects of unidirectional permeability in asymmetric poly (DL-lactic acid-co-glycolic acid) conduits on peripheral nerve regeneration: An in vitro and in vivo study. Journal of Biomedical Materials Research B Applied Biomaterials 2007;83:206-15.
29Temple CL, Ross DC, Dunning CE, Johnson JA. Resistance to disruption and gapping of peripheral nerve repairs: An in vitro biomechanical assessment of techniques. Journal of Reconstructive Microsurgery 2004;20:645-50.
30Sameem M, Wood TJ, Bain JR. A systematic review on the use of fibrin glue for peripheral nerve repair. Plastic and Reconstructive Surgery 2011;127:2381-90.
31Bittner GD, Schallert T, Peduzzi JD. Degeneration, trophic interactions, and repair of severed axons: A reconsideration of some common assumptions. Neuroscientist 2000;6:88-109.
32Bittner GD, Sengelaub DR, Trevino RC, Peduzzi JD, Mikesh M, Ghergherehchi CL, et al. The curious ability of PEG-fusion technologies to restore lost behaviors after nerve severance. J Neurosci Res 2016;94:207-30.
33Williams HB. The value of continuous electrical muscle stimulation using a completely implantable system in the preservation of muscle function following motor nerve injury and repair: An experimental study. Microsurgery 1996;17:589-96.
34Mohanty CB, Midha R. Nerve section causes brain reaction. World Neurosurg 2015;84:886-8.
35Mohanty CB. Central plasticity in brachial plexus injury: A neural domino effect. World Neurosurg 2016;86:22-4.
36Mohanty CB, Bhat D, Devi BI. Role of central plasticity in the outcome of peripheral nerve regeneration. Neurosurgery 2015;77:418-23.
37Navarro X. Neural plasticity after nerve injury and regeneration. Int Rev Neurobiol 2009;87:483-505.
38Bahari Z, Sadr SS, Meftahi GH, Ghasemi M, Manaheji H, Mohammadi A, et al. Nerve injury-induced plasticity in the nociceptive pathways. Arch Neurosci 2015;2:e18214.
39Jaggi AS, Jain V, Singh N. Animal models of neuropathic pain. Fundam Clin Pharmacol 2011;25:1-28.
40Shurey S, Akelina Y, Legagneux J, Malzone G, Jiga L, Ghanem AM. The rat model in microsurgery education: Classical exercises and new horizons. Arch Plast Surg 2014;41:201-8.
41Mandal J, Parija SC. Ethics of involving animals in research. Tropical Parasitology 2013;3:4-6.
42Pereira S, Tettamanti M. Ahimsa and alternatives-the concept of the 4th R. The CPCSEA in India. ALTEX 2005;22:3-6.
43Geuna S. The sciatic nerve injury model in pre-clinical research. J Neurosci Methods 2015;243:39-46.
44Speck AE, Ilha J, do Espírito Santo CC, Aguiar AS Jr, Dos Santos AR, Swarowsky A. The IBB forelimb scale as a tool to assess functional recovery after peripheral nerve injury in mice. J Neurosci Methods 2014;226:66-72.
45Stößel M, Rehra L, Haastert-Talini K. Reflex-based grasping, skilled forelimb reaching, and electrodiagnostic evaluation for comprehensive analysis of functional recovery—The 7-mm rat median nerve gap repair model revisited. Brain Behav 2017;7:e00813.
46Cobianchi S, de Cruz J, Navarro X. Assessment of sensory thresholds and nociceptive fiber growth after sciatic nerve injury reveals the differential contribution of collateral reinnervation and nerve regeneration to neuropathic pain. Exp Neurol 2014;255:1-11.
47Navarro X, Udina E. Methods and protocols in peripheral nerve regeneration experimental research: Part III-electrophysiological evaluation. Int Rev Neurobiol 2009;87:105-26.
48Gramsbergen A, IJkema-Paassen J, Meek MF. Sciatic nerve transection in the adult rat: Abnormal EMG patterns during locomotion by aberrant innervation of hindleg muscles. Exp Neurol 2000;161:183-93.
49Chen XY, Yin YF, Zhang TT, Zhao YH, Yang YM, Yu XM, et al. Ultrasound imaging of chitosan nerve conduits that bridge sciatic nerve defects in rats. Neural Regen Res 2014;9:1386-8.
50Hayashi A, Moradzadeh A, Hunter DA, Kawamura DH, Puppala VK, Tung TH, et al. Retrograde labeling in peripheral nerve research: it is not all black and white. J Reconstr Microsurg 2007;23:381-9.
51Yu Y, Li H, Zhang P, Yin X, Han N, Kou Y, et al. Comparison of commonly used retrograde tracers in rat spinal motor neurons. Neural Regen Res 2015;10:1700-5.
52Jolivalt CG, Vu Y, Mizisin LM, Mizisin AP, Calcutt NA. Impaired prosaposin secretion during nerve regeneration in diabetic rats and protection of nerve regeneration by a prosaposin-derived peptide. J Neuropathol Exp Neurol 2008;67:702-10.
53Clavijo-Alvarez JA, Nguyen VT, Santiago LY, Doctor JS, Lee WP, Marra KG. Comparison of biodegradable conduits within aged rat sciatic nerve defects. Plast Reconstr Surg 2007;119:1839-51.
54Alant JD, Kemp SW, Khu KJ, Kumar R, Webb AA, Midha R. Traumatic neuroma in continuity injury model in rodents. J Neurotrauma 2012;29:1691-703.
55Gordon T, Amirjani N, Edwards DC, Chan KM. Brief post-surgical electrical stimulation accelerates axon regeneration and muscle reinnervation without affecting the functional measures in carpal tunnel syndrome patients. Exp Neurol 2010;223:192-202.
56Kumar TR, Larson M, Wang H, McDermott J, Bronshteyn I. Transgenic mouse technology: Principles and methods. Methods Mol Biol 2009;590:335-62.
57Bhat DI. Animal models for cerebral vasospasm: Where do we stand?. Neurol India 2017;65:1043-5.
58Myckatyn TM, Mackinnon SE, Hunter DA, Brakefield D, Parsadanian A. A novel model for the study of peripheral-nerve regeneration following common nerve injury paradigms. J Reconstr Microsurg 2004;20:533-44.
59Unezaki S, Yoshii S, Mabuchi T, Saito A, Ito S. Effects of neurotrophic factors on nerve regeneration monitored by in vivo imaging in thy1-YFP transgenic mice. J Neurosci Methods 2009;178:308-15.