Bioelectronic Medicine

, Volume 2, Issue 1, pp 43–48 | Cite as

Chronic Electrical Nerve Stimulation as a Therapeutic Intervention for Peripheral Nerve Repair

  • Miyuki Sakuma
  • Ivan R. Minev
  • Sandra Gribi
  • Bhagat Singh
  • Clifford J. WoolfEmail author
  • Stéphanie P. LacourEmail author
Open Access
Invited Review Article


When a peripheral nerve is injured after either trauma or a neurodegenerative disease, motor function and sensory perception are impaired. Repair strategies aim both at reconstructing the damaged nerve and in promoting regeneration to enhance target reinnervation and functional recovery. Advanced surgical procedures can enable efficient repair, but restoration of function remains challenging. Among various factors influencing nerve regeneration, electrical stimulation is often cited as a potential therapeutic approach to nerve repair, engaging regenerative transcriptional programs. In this report, we review both reported effects on axonal growth and functional outcomes of electrical stimulation on peripheral nerve repair and the techniques for chronic nerve stimulation, highlighting the challenges and opportunities of such repair strategies.


Peripheral nerve damage is an important clinical problem, with an occurrence of up to 3% of all trauma patients (1, 2, 3). Peripheral nerve injuries often result in loss in motor function, sensory function or both and are associated with a high socioeconomic impact (4, 5, 6). After nerve injury, the damaged axons of sensory and motor neurons can regenerate and, under optimal circumstances, can reinnervate peripheral target organs such as the skin and muscles (7). However, functional outcome after repair (despite modern microsurgery techniques) is often poor and incomplete. Extensive soft tissue injury, a large gap between nerves stumps, long distances to reach receptive targets and misdirected axons contribute to the challenging repair environment (8, 9, 10, 11). There is an urgent need to find new treatments for promoting regeneration after peripheral nerve injury, especially since chronic denervation results in muscle atrophy and progressive decline in Schwann cell support, making the distal nerve progressively less growth-permissible (12).

After trauma or neurodegenerative diseases, electrical stimulation of neural tissue can be used to either alleviate symptoms or restore function (13). For example, deep brain stimulation relieves the effects of Parkinson’s disease; cochlear implants convert sound waves into stream of electrical impulses that are transmitted to the auditory nerve to partially restore hearing; direct electrical stimulation enhances wound healing; and transcutaneous electrical nerve stimulation is a nonpharmacological therapy for pain relief.

Electrical stimulation has been also envisioned as a potential complementary therapy to microsurgical repair after peripheral nerve injury to improve functional outcomes by promoting a greater and faster axonal regeneration. A number of preclinical studies across a number of nerve injury models have shown that direct stimulation of injured peripheral nerves can enhance peripheral sensory (14,15) and motor (16, 17, 18) axon regeneration, increase target reinnervation (17,19) and improve electrophysiological outcomes (20,21) and functional recovery (14,15). To date, however, electrical stimulation is rarely used in a clinical context of nerve repair. One study has reported accelerated reinnervation of the thenar muscle following nerve electrical stimulation after carpal tunnel release surgery (22).

Reviewing the effect of electrical stimulation on nerve repair is challenging because results from preclinical studies are often inconsistent, and their impacts on peripheral nerve repair vary substantially. This can be attributed to the vast number of experimental protocols, for which parameters include: type of injury (crush versus transection), nature of microsurgical repair, type of stimulation used (direct current or pulsed stimulation), location of the stimulation (transcutaneous, proximal or distal nerve stump), onset of stimulation (immediately after injury or delayed), its duration (acute versus chronic), stimulation waveforms (frequency, amplitude, pulse duration, biphasic) and ways to assert physiological and functional outcomes. Given this large methodological variance and consequent inconclusive data, the justification for such interventions at the clinic remains uncertain. Furthermore, it is still not clear whether any failure to detect an improvement is due to a lack of biological impact of electrical stimulation or a technical failure, or both.

Here we review the effects of electrical stimulation on peripheral nerve repair, distinguishing regenerative effects and functional outcomes. On the basis of preliminary work that we have performed to enable chronic stimulation of the sciatic nerve after a crush injury, we discuss the challenges and opportunities for developing tools and methods for chronic stimulation of injured peripheral nerves.

Peripheral Nerve Electrical Stimulation


Electrical stimulation may be applied with surface skin electrodes or by means of invasive electrodes placed in the direct vicinity of an injured nerve. Few studies report the use of transcutaneous electrical stimulation for nerve repair, with inconsistent outcomes for both regeneration and function (23,24). Surface electrodes are often >1 cm2 cloth-like pads coated with a coupling gel above the site of nerve injury and have relatively low impedance, in the kΩ range.

Invasive nerve electrodes are either designed as wires/needles (mainly used for acute stimulation) or cuff electrodes. A metallic wire (cathode) is typically wrapped around the proximal nerve, and a second wire electrode (anode) is inserted within a distal muscle or wrapped along the nerve distally. Such an electrode configuration is commonly implemented for acute stimulation following a nerve crush injury, nerve transection with end-to-end suturing of the proximal and distal nerve stumps (16–19,25–27) or nerve repair facilitated by a biocompatible conduit sutured between both nerve stumps (21,28). Their impedance is typically in the 10-s MΩ range because of their small effective surface area.

Cuff electrodes in contrast are nerve diameter-sized tubes hosting metallic contacts with a monopolar, bipolar or tripolar configuration. This design provides selective activation of the nerve fibers. The impedance of the contacts is in the kΩ range. Chronic (long-term, >2 wks) stimulation of the injured nerve is possible, provided the cuff is secured to the tissue and induces negligible constriction of the nerve during movement. Subcutaneous wiring to a back- or head-plug enables minimally disruptive experiments. Ultimately long-term electrodes could interface a wireless, implanted microstimulator.

Parameters of Electrical Stimulation

In most cases, a pulsed stimulation is implemented. Four criteria defining electrical stimulation protocols need to be preselected: (a) pulse waveform (frequency, width, amplitude and shape), (b) duration of stimulation, (c) onset time of nerve stimulation after injury and (d) overall duration of the stimulation treatment. The most frequently used parameters are listed below:
  • Threshold stimulation current: 0.5–5 mA range, but depends on electrode type, configuration and distance from the injured nerve;

  • Cathodic pulse width: 50–400 µs;

  • Pulse frequency: 1–20 Hz;

  • Session duration: 15 min to 1 h;

  • Acute stimulation: immediately after nerve injury or after surgical repair of the injured nerve;

  • Long-term stimulation: 5 d/wk or every other day for up to 8 wks.

Different types of stimulation (direct current [DC] and alterative current [AC]) and various stimulation parameters used and their impact on peripheral nerve regeneration are summarized in Table 1.
Table 1

Reported effects on nerve injury recovery after electrical stimulation


Journal (year)

Type of current used, site of stimulation and parameters

Type of injury


Hoffman H

Aust. J. Exp. Biol. Med. Sci. (1952)

AC, 6.3 V, 50–100 cycles, 1.5 m A, 0–5,000 Ω variable resistance

Spinal cord injury

Enhanced reinnervation and sprouting

Maehlen J, NjåA

J. Physiol. (1982)

Preganglionic stimulation for 1 h immediately after the partial denervation with 100 pulses at 20 Hz every 25 s

Thoraco-cervical sympathetic trunk transection

Increased rate of sprouting

Nix WA, Hopf HC

Brain Res. (1983)

AC, 0.2 ms duration, frequency of 4 pulses per second (pps), applied (24 h daily) for 4 wks, and stimulation started 1 d postoperatively

Sciatic nerve transection

Improved electrophysiological recovery

Pockett S, Gavin R

Neurosci. Lett. (1985)

AC, proximal stump, 0.1-ms pulses, supramaximal voltage

Sciatic nerve crush

Improvement in toe spread function

McDevitt, et al.

Brain Res. (1987)

DC, 10 µA/cm2, Distal cathode, hind paw, field strength ∼100 mV/cm, 100 kΩ resistance, daily for 20 d

Sciatic nerve transection and repair

Enhanced motor axon regeneration

Román GC, et al.

Exp. Neurol. (1987)

DC, distal cathode implantation with a 10-µA for 3 wks

Sciatic nerve transection and repair

Increased number of myelinated axons

Zanakis MF, et al.

Acupunct. Electrother. Res. (1990)

DC, 1.4 µA (about 8 mV/cm field strength) to a nerve cuff

Sciatic nerve crush

Enhanced number of regenerating axons in the distal stump

Kerns JM, et al.

Exp. Neurol. (1992)

DC, 10 µA/cm2, Distal cathode

Sciatic nerve crush

No change in sciatic function index (SFI)

Pomeranz B, et al.

Brain Res. (1993)

DC of 10 µA, 200–270 kΩ resistor, stimulation for a month

Sciatic nerve crush

Improved electrophysiological outcomes

Al-Majed AA, et al.

J. Neurosci. (2000)

AC, proximal stump, 0.1 ms,3V, 20 Hz, 1 h immediately after repair and up to 2 wks

Femoral nerve transection and repair

Accelerated motor axons regeneration across repair site after 1 h of stimulation but no further benefits with chronic stimulation

Mendonça CA, et al.

J. Neurosci. Methods (2003)

DC, proximal stump, low-intensity continuous current circuit (1 µA), 1.5 V battery and a 1.3-MΩ resistor for 3 wks

Sciatic nerve crush

Improved SFI

Ahlborn P, et al.

Exp. Neurol. (2007)

AC, proximal stump, square 0.1-ms pulses, 20 Hz, 3–4 V, acute

Femoral nerve transection

Improved motor axon regeneration and behavioral recovery

Geremia NM, et al.

Exp. Neurol. (2007)

AC, proximal stump, 0.1 ms, 3V, 20 Hz, 1 h immediately after repair and up to 3 wks

Femoral nerve transection and repair

Improved sensory axon regeneration with acute stimulation but reduced benefits with chronic stimulation

Singh B, et al.

J. Neurosurg. (2012)

AC, proximal stump, 0.1 ms,3V, 20 Hz, 1 h immediately after repair

Sciatic nerve transection and repair

Improved axon regeneration and target reinnervation

Huang J, et al.

Eur. J. Neurosci. (2013)

AC (3 V, 20 Hz, 20 min) applied proximally to the transected nerve while repairing (2, 4, 12 and 24 wks)

Sciatic nerve transection and delayed repair

Increased motoneurons and sensory neurons regeneration, improvement in CMAP and NCV up to 24 wks of delay

Zhang X, et al.

Mol. Med. Rep. (2013)

AC (3 V, 20 Hz, 1 h) applied proximally to the nerve

Sciatic nerve crush

Improved remyelination, axon diameter and electrophysiological measures

Calvey C, et al.

J. Hand Surg. Am. (2014)

A direct current of 24 V/m (24 mV, DC 1.5 mA), applied across the electrodes for 10 min and 60 min

Sciatic nerve transection and repair

Enhanced behavioral and histological recovery

Xu C, et al.

PLoS One (2014)

AC, proximal stump, 0.1 ms, 3 V, 20 Hz, delayed nerve repair after 1 d, 1 wk, 1 month and 2 months

Sciatic nerve transection and repair at different time points

Improved electrophysiology parameters but the impact reduced with the delay

Thompson NJ, et al.

Dev. Neurobiol. (2014)

AC, short (0.1 ms) pulses at 20 Hz, 1 h immediately before nerve transection

Sciatic nerve transection and repair

Enhanced axon regeneration

An important issue lies in the identification of those fibers that need to be activated by electrical stimulation. The nerve contains motor and sensory fibers, which have different stimulation thresholds: motor axons are activated with lower stimulation amplitude than unmyelinated sensory axons. Recruitment of small myelinated or unmyelinated fibers, which signal pain, itch and thermal sensations, will produce nocifensive responses and will generally not be tolerated. As motor axons begin to reform functional neuromuscular junctions, movement will be elicited, and this may not be tolerated in a freely moving animal. The duration of the stimulation session may be important for regeneration. For example, both 1-h-long and 2-wk-long electrical stimulation have been reported to lead to a similar increase in the number of regenerated motor neurons (17), whereas a 3-wk stimulation was found to reduce the number of backlabeled sensory neurons compared with a 1-h, acute electrical stimulation session (26).

Electrical Stimulation Modulates the Molecular and Cellular Activity Involved in Nerve Regeneration

Peripheral nerve injury results in plastic changes in injured neurons. The sudden exposure to an extracellular medium triggers an immediate influx of sodium and calcium ions into the injured axons, causing a high-frequency burst of action potentials in the severed axons. A phenotypic switch of the injured neurons from a maintenance to a regenerative state then ensues, characterized by up- and downregulation of hundreds of genes. These include increased expression of brain-derived neurotrophic factor (BDNF) and its receptor tropomyosin kinase receptor B (trkB), growth associated protein 43 (GAP-43) and the cytoskeletal proteins Tα-1 tubulin and actin, and a downregulation of enzymes such as choline acetyltransferase (ChAT) and acetylcholinesterase (AchE). To facilitate retrograde protein transport along the axons, neurofilament genes (NF-L/M/H mRNA) are concomitantly downregulated (7).

Acute electrical stimulation (1 h, 20 Hz) applied immediately after injury proximal to the injured nerve appears to lead to a more robust and earlier phenotypic switch (16,17) with, for example, enhanced mRNA and protein expression of BDNF and GAP-43 in adult sensory neurons (21,26,29,30). In a recent study, androgen receptor signaling was elucidated as a possible cascade activated by electrical stimulation (31). Electrical stimulation also was found to induce an upregulation of NT-3/4 in sensory neurons, which may promote axonal regeneration (32).

The specific mechanisms by which electrical stimulation may induce improved nerve regeneration are not yet clear. However, the available data suggest that the enhanced production of growth factors, especially BDNF, may have a significant role. Blockade of the electrostimulation-induced regenerative response in both motor and sensory neurons by tetrodotoxin (TTX) application at the site of injury implies involvement of neuronal cell body mechanisms triggered by electrical activity (16,26). However, a contribution of non-neuronal cell types (Schwann cells at the site of injury and in the distal stump and periganglionic satellite cells in the dorsal root ganglia) cannot be excluded. Enhanced glial fibrillary acidic protein (GFAP) expression in Schwann cells was, for example, observed after local electrical stimulation of injured nerves (33). Furthermore, Schwann cells respond to electrical stimulation by producing increased nerve growth factor (NGF) due to rises in calcium levels (34), and this could increase growth of TrkA-expressing nociceptors. Overall, although there are several phenomenological studies indicating some benefit of acute electrical stimulation on peripheral nerve regeneration, the specific mechanisms responsible still remain elusive.

Benefits of Prolonged Electrical Stimulation

Few studies have examined the effects of chronic electrical stimulation of injured nerves over time (35, 36, 37). Changes in histological metrics (myelin thickness, axon density, blood vessel area and density, axon and fiber density, g-ratio, reinnervation specificity, mRNA and proteins expression), electrophysiological responses (compound muscle action potential, nerve conduction velocity, M/H wave ratio, latency and amplitude of electrical response) and functional outcomes (functional stance recovery, sciatic function index, thermal and mechanical algesimetry, toe-pinch test) have been reported for stimulation up to 24 wks after injury. However, results are inconsistent. Single time-point analyses have revealed either significant improvements at 2, 3 and 6 wks after repair (19,38,39) or no improvement at 4 wks (40). Over longer periods, between 6 and 12 wks after injury, no significant differences were observed between stimulated versus nonstimulated nerves. This result may reflect an action of electrical stimulation only in an earlier activation of intrinsic growth programs, which is not sustained so that the normal pattern of growth eventually “catches up” (17,39,41). Chronic stimulation delivered for 8 h a day was actually found to be counterproductive over longer periods, exacerbating nerve atrophy in a rabbit model (42). Daily stimulation of 1 h, 5 d a week, failed to promote motor reinnervation in comparison to single acute stimulation but reduced the H/M wave ratio, indicating decreased hyperreflexia 60 d after injury (27,43). Hyperreflexia is a reflection of neuronal reorganization and plasticity in the spinal cord after peripheral nerve injury (7). Whereas 1 h of stimulation applied every other day on eight occasions was found to produce significant recovery of the sciatic function index at 12 wks as well as a higher reinnervation of both sensory and motoneurons, reducing the electrical stimulation to 15 min every other day starting at 1 wk, post-injury repair did not produce any improvement of motor responses at 6 wks. However, a significant increase in blood vessel density in the nerve was observed (39).

Table 1 summarizes different experimental protocols used to test if electrical stimulation alters regeneration and lists the major findings for each study.

Challenges of Chronic Peripheral Nerve Stimulation

We performed a pilot study to evaluate the technical challenges of chronic nerve stimulation in the rat. A cuff electrode was applied proximal to a sciatic nerve crush injury site and the nerve was stimulated for 2–3 wks after the injury.

We tested several types of silicone cuff electrode, which had subtle variations in their geometry. All implants were prototyped by using silicone rubber, multistrand stainless steel wires (300-µm outer diameter [o.d.]) and an Omnetic connector mounted on the head of the animals. After braiding three wires equally spaced around a polystyrene cylindrical mandrel (tripolar configuration, 1.5 mm diameter, matching the sciatic nerve diameter), the structure was flooded with quick-cure silicone (KWIK-SIL, World Precision Instruments, Sarasota, FL, USA). The distal ends of the wires were then cut and the plastic mandrel dissolved in acetone. Small additional amounts of silicone were added to cover any exposed portion of the steel wires. The cuff was then cut to the desired length. The cuff endings were terminated with either flat or tapered edges. To form the soft edge, the cuff was briefly dipped in freshly mixed PDMS (polydimethylsiloxane) before removal of the mandrel. This ensured formation of a gradually tapered edge. Next, a longitudinal slit was made in the cuff with fine surgical scissors, allowing for easy opening with a pair of forceps and gentle insertion of the nerve.

Implantation of the soft cuff electrodes designed to wrap around the rat sciatic nerve proved challenging. Anchoring the cuff implant to the soft tissue surrounding the nerve did not provide a stable way to secure the cuff in position. The stretching or twisting of the nerve within the electrode cuff during leg movement often triggered mechanical damage to the nerve and inflammation. “Hard,” straight, edge cuffs induced much more damage than soft. However, extremely soft cuffs then proved difficult to anchor to the surrounding tissues with nylon sutures, without compressing the nerve.

Another challenge associated with long-term peripheral nerve stimulation was management of the wiring cable connecting the cuff electrodes to the head-plug connector. As the rat grew over the course of the experiment, tension in the cable in some cases pulled the connector wires out of the cuff. Adding length to the cables to provide slack did not solve this issue because the cables became surrounded with scar tissue and fixed in position.

Our experience indicates that the technical challenges associated with long-term peripheral nerve interfaces in the rat are considerable and should first be solved before physiological, anatomical and functional assessment of nerve regeneration and its modulation by electrical stimulation can be performed. Nevertheless, we feel that this should be possible and needs to be done.


Although considerable effort has been devoted by many investigators to the pre-clinical analysis of the effects of peripheral nerve stimulation on axonal regeneration in rodents, these effects have not yet translated into therapy. To better understand the extent to which electrical stimulation may enhance sensory and motor axon regeneration and if there is a correlation between such regeneration and improved functional outcome, further neurotechnology developmental efforts are needed to enable the necessary long-term chronic stimulation studies to be conducted. Minimizing nerve damage by using soft, nerve-like materials and ultra-compliant wiring, together with the development of new surgical techniques for inserting and stabilizing the electrodes, as well as development of sensitive behavioral outcome measures accurately reflecting successful sensory and motor recovery in freely moving animals, would enable the long-term preclinical studies needed to assess if chronic stimulation is a beneficial strategy for treating axon damage in peripheral nerves and what stimulation protocol is most effective.


The authors declare that they have no competing interests as defined by Bioelectronic Medicine, or other interests that might be perceived to influence the results and discussion reported in this paper.



Financial support was provided by the Foundation Bertarelli (SP Lacour and CJ Woolf), the Funds National Suisse (FNS) and a Yonemitsu Foundation Fellowship in Kumamoto Kino Hospital to M Sakuma. B Singh holds a Canadian Institute of Health Research (CIHR) postdoctoral fellowship. We thank Fengfeng Bei for suggestions in improving head-plug fixation and Lee Barrett for assisting in animal dissection and microscopy.


  1. 1.
    Taylor CA, Braza D, Rice JB, Dillingham T. (2008) The incidence of peripheral nerve injury in extremity trauma. Am. J. Phys. Med. Rehabil. 87:381–5.CrossRefGoogle Scholar
  2. 2.
    Asplund M, Nilsson M, Jacobsson A, von Holst H. (2009) Incidence of traumatic peripheral nerve injuries and amputations in Sweden between 1998 and 2006. Neuroepidemiology. 32:217–28.CrossRefGoogle Scholar
  3. 3.
    Pfister BJ, et al. (2011) Biomedical engineering strategies for peripheral nerve repair: surgical applications, state of the art, and future challenges. Crit Rev Biomed Eng 39:81–124.CrossRefGoogle Scholar
  4. 4.
    Missios S, Bekelis K, & Spinner RJ (2014) Traumatic peripheral nerve injuries in children: epidemiology and socioeconomics. J. Neurosurg. Pediatr 14:688–94.CrossRefGoogle Scholar
  5. 5.
    Lad SP, Nathan JK, Schubert RD, & Boakye M (2010) Trends in median, ulnar, radial, and brachioplexus nerve injuries in the United States. Neurosurgery 66:953–60.CrossRefGoogle Scholar
  6. 6.
    Ciaramitaro P, et al. (2010) Traumatic peripheral nerve injuries: epidemiological findings, neuropathic pain and quality of life in 158 patients. J Peripher Nerv Syst 15:120–7.CrossRefGoogle Scholar
  7. 7.
    Navarro X, Vivo M, & Valero-Cabre A (2007) Neural plasticity after peripheral nerve injury and regeneration. Prog Neurobiol 82:163–201.CrossRefGoogle Scholar
  8. 8.
    Brown PW (1972) Factors influencing the success of the surgical repair of peripheral nerves. Surg Clin North Am 52:1137–55.CrossRefGoogle Scholar
  9. 9.
    Ruijs AC, Jaquet JB, Kalmijn S, Giele H, Hovius SE (2005) Median and ulnar nerve injuries: a meta-analysis of predictors of motor and sensory recovery after modern microsurgical nerve repair. Plast Reconstr Surg 116:484–94; discussion 95–6.CrossRefGoogle Scholar
  10. 10.
    Murovic JA (2009) Upper-extremity peripheral nerve injuries: a Louisiana State University Health Sciences Center literature review with comparison of the operative outcomes of 1837 Louisiana State University Health Sciences Center median, radial, and ulnar nerve lesions. Neurosurgery 65:A11–7.CrossRefGoogle Scholar
  11. 11.
    He B, et al. (2014) Factors predicting sensory and motor recovery after the repair of upper limb peripheral nerve injuries. Neural Regen Res 9:661–72.CrossRefPubMedPubMedCentralGoogle Scholar
  12. 12.
    Gordon T, Tyreman N, & Raji MA (2011) The basis for diminished functional recovery after delayed peripheral nerve repair. J. Neurosci. 31:5325–34.CrossRefGoogle Scholar
  13. 13.
    Borton D, Micera S, Millan Jdel R, & Courtine G (2013) Personalized neuroprosthetics. Sci Transl Med 5:210rv2.CrossRefGoogle Scholar
  14. 14.
    Ahlborn P, Schachner M, & Irintchev A (2007) One hour electrical stimulation accelerates functional recovery after femoral nerve repair. Exp Neurol 208:137–44.CrossRefGoogle Scholar
  15. 15.
    Singh B, et al. (2012) Accelerated axon outgrowth, guidance, and target reinnervation across nerve transection gaps following a brief electrical stimulation paradigm. J. Neurosurg. 116:498–512.CrossRefGoogle Scholar
  16. 16.
    Al-Majed AA, Brushart TM, Gordon T. (2000) Electrical stimulation accelerates and increases expression of BDNF and trkB mRNA in regenerating rat femoral motoneurons. Eur. J. Neurosci. 12:4381–90.PubMedGoogle Scholar
  17. 17.
    Al-Majed AA, Neumann CM, Brushart TM, Gordon T. (2000) Brief electrical stimulation promotes the speed and accuracy of motor axonal regeneration. J. Neurosci. 20:2602–8.CrossRefGoogle Scholar
  18. 18.
    Al-Majed AA, Tam SL, Gordon T. (2004) Electrical stimulation accelerates and enhances expression of regeneration-associated genes in regenerating rat femoral motoneurons. Cell. Mol. Neurobiol. 24:379–402.CrossRefGoogle Scholar
  19. 19.
    Brushart TM, Jari R, Verge V, Rohde C, Gordon T. (2005) Electrical stimulation restores the specificity of sensory axon regeneration. Exp. Neurol. 194:221–9.CrossRefGoogle Scholar
  20. 20.
    Nix WA, Hopf HC. (1983) Electrical stimulation of regenerating nerve and its effect on motor recovery. Brain Res. 272:21–5.CrossRefGoogle Scholar
  21. 21.
    Xu C, et al. (2014) Electrical stimulation promotes regeneration of defective peripheral nerves after delayed repair intervals lasting under one month. PLoS One. 9:e105045.CrossRefPubMedPubMedCentralGoogle Scholar
  22. 22.
    Gordon T, Brushart TM, Chan KM. (2008) Augmenting nerve regeneration with electrical stimulation. Neurol. Res. 30:1012–22.CrossRefGoogle Scholar
  23. 23.
    Cavalcante Miranda de Assis D, et al. (2014) The parameters of transcutaneous electrical nerve stimulation are critical to its regenerative effects when applied just after a sciatic crush lesion in mice. Biomed. Res. Int. 2014:572949.CrossRefPubMedPubMedCentralGoogle Scholar
  24. 24.
    del Valle J, Navarro X. (2013) Interfaces with the peripheral nerve for the control of neuroprostheses. Int. Rev. Neurobiol. 109:63–83.CrossRefGoogle Scholar
  25. 25.
    Brushart TM, et al. (2002) Electrical stimulation promotes motoneuron regeneration without increasing its speed or conditioning the neuron. J. Neurosci. 22:6631–8.CrossRefGoogle Scholar
  26. 26.
    Geremia NM, Gordon T, Brushart TM, Al-Majed AA, Verge VM. (2007) Electrical stimulation promotes sensory neuron regeneration and growth-associated gene expression. Exp. Neurol. 205:347–59.CrossRefGoogle Scholar
  27. 27.
    Asensio-Pinilla E, Udina E, Jaramillo J, Navarro X. (2009) Electrical stimulation combined with exercise increase axonal regeneration after peripheral nerve injury. Exp. Neurol. 219:258–65.CrossRefGoogle Scholar
  28. 28.
    Huang J, et al. (2013) Dog tibial nerve regeneration across a 30-mm defect bridged by a PRGD/PDLLA/beta-TCP/NGF sustained-release conduit. J. Reconstr. Microsurg. 29:77–87.CrossRefGoogle Scholar
  29. 29.
    Wang JH, et al. (2009) Effect of electroacupuncture of different acupoints on the excitability of detrusor muscle and the expression of BDNF and TrkB in the spinal cord of rats with urinary retention due to spinal cord injury [in Chinese]. Zhen Ci Yan Jiu. 34:387–92.PubMedGoogle Scholar
  30. 30.
    Huang J, et al. (2012) Electrical stimulation to conductive scaffold promotes axonal regeneration and remyelination in a rat model of large nerve defect. PLoS One. 7:e39526.CrossRefPubMedPubMedCentralGoogle Scholar
  31. 31.
    Thompson NJ, Sengelaub DR, English AW. (2014) Enhancement of peripheral nerve regeneration due to treadmill training and electrical stimulation is dependent on androgen receptor signaling. Dev. Neurobiol. 74:531–40.CrossRefGoogle Scholar
  32. 32.
    English AW, Schwartz G, Meador W, Sabatier MJ, Mulligan A. (2007) Electrical stimulation promotes peripheral axon regeneration by enhanced neuronal neurotrophin signaling. Dev. Neurobiol. 67:158–72.CrossRefPubMedPubMedCentralGoogle Scholar
  33. 33.
    McLean NA, Popescu BF, Gordon T, Zochodne DW, Verge VM. (2014) Delayed nerve stimulation promotes axon-protective neurofilament phos-phorylation, accelerates immune cell clearance and enhances remyelination in vivo in focally demyelinated nerves. PLoS One. 9:e110174.CrossRefPubMedPubMedCentralGoogle Scholar
  34. 34.
    Huang F, et al. (2007) GM1 and NGF modulate Ca2+ homeostasis and GAP43 mRNA expression in cultured dorsal root ganglion neurons with excitotoxicity induced by glutamate. Nutr. Neurosci. 10:105–11.CrossRefGoogle Scholar
  35. 35.
    Li YT, et al. (2013) Application of implantable wireless biomicrosystem for monitoring nerve impedance of rat after sciatic nerve injury. IEEE Trans. Neural. Syst. Rehabil. Eng. 21:121–8.CrossRefGoogle Scholar
  36. 36.
    Zorko B, Rozman J, Seliskar A. (2000) Influence of electrical stimulation on regeneration of the radial nerve in dogs. Acta. Vet. Hung. 48:99–105.CrossRefGoogle Scholar
  37. 37.
    Branner A, Stein RB, Fernandez E, Aoyagi Y, Normann RA. (2004) Long-term stimulation and recording with a penetrating microelectrode array in cat sciatic nerve. IEEE Trans. Biomed. Eng. 51:146–57.CrossRefGoogle Scholar
  38. 38.
    Haastert-Talini K, et al. (2011) Electrical stimulation accelerates axonal and functional peripheral nerve regeneration across long gaps. J. Neurotrauma. 28:661–74.CrossRefGoogle Scholar
  39. 39.
    Chen YS, et al. (2001) Effects of percutaneous electrical stimulation on peripheral nerve regeneration using silicone rubber chambers. J. Biomed. Mater. Res. 57:541–9.CrossRefGoogle Scholar
  40. 40.
    Lee TH, et al. (2010) Functional regeneration of a severed peripheral nerve with a 7-mm gap in rats through the use of an implantable electrical stimulator and a conduit electrode with collagen coating. Neuromodulation. 13:299–304.CrossRefGoogle Scholar
  41. 41.
    Huang J, et al. (2009) Electrical stimulation accelerates motor functional recovery in autograft-repaired 10 mm femoral nerve gap in rats. J. Neurotrauma. 26:1805–13.CrossRefGoogle Scholar
  42. 42.
    Gordon T, Gillespie J, Orozco R, Davis L. (1991) Axotomy-induced changes in rabbit hindlimb nerves and the effects of chronic electrical stimulation. J. Neurosci. 11:2157–69.CrossRefGoogle Scholar
  43. 43.
    Vivo M, et al. (2008) Immediate electrical stimulation enhances regeneration and reinnervation and modulates spinal plastic changes after sciatic nerve injury and repair. Exp. Neurol. 211:180–93.CrossRefGoogle Scholar

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Authors and Affiliations

  • Miyuki Sakuma
    • 1
  • Ivan R. Minev
    • 2
  • Sandra Gribi
    • 2
  • Bhagat Singh
    • 1
  • Clifford J. Woolf
    • 1
    Email author
  • Stéphanie P. Lacour
    • 3
    Email author
  1. 1.F.M. Kirby Neurobiology CenterBoston Children’s Hospital, Harvard Medical SchoolBostonUSA
  2. 2.Laboratory for Soft Bioelectronic Interfaces, Institute of Microengineering and Institute of Bioengineering, Centre for NeuroprostheticsEcole Polytechnique Fédérale de Lausanne (EPFL)LausanneSwitzerland
  3. 3.Bertarelli Foundation Chair in Neuroprosthetic Technology, Laboratory for Soft Bioelectronic Interfaces, Institute of Microengineering and Institute of Bioengineering, Centre for NeuroprostheticsEcole Polytechnique Fédérale de Lausanne (EPFL)LausanneSwitzerland

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