Electrical Stimulation Therapy to Accelerate Nerve Regeneration Remains Effective Following Postoperative Application of Lidocaine

Abstract

Background:

Brief electrical stimulation (ES) of injured peripheral nerves for 1 hour has been shown to accelerate nerve regeneration with proximal action potential conduction to the neuron cell body, a requirement to elicit therapeutic benefit. Local anesthetic is often used to manage pain in patients. However, using lidocaine after ES therapy has been controversial. We assessed the effects of extraneural usage of lidocaine after ES therapy on nerve regeneration in a rodent nerve injury model.

Methods:

Lewis rats underwent tibial nerve transection and immediate repair and randomized to 4 groups: control (REP), extraneural lidocaine alone (REP + LIDO), 60-minute ES (60 ES), and 60-minute ES with extraneural lidocaine (60 ES + LIDO). The tibial nerve was retrograde labeled distally from the neurorrhaphy 28 days post repair. Spinal cords and dorsal root ganglia were harvested to assess motor and sensory neuron counts. Data were analyzed using 1-way analysis of variance (ANOVA) with a post-hoc Tukey correction.

Results:

Using lidocaine after nerve repair did not affect nerve regeneration in the control group (REP vs REP + LIDO) or ES group (60 ES vs 60 ES + LIDO), with motor and sensory neuron counts not statistically different between groups. Electrical stimulation therapy showed at least a 60% increase in motor and sensory neuron counts than controls, a statistically significant effect (P < .001).

Conclusions:

Extraneural usage of lidocaine after ES does not abolish the improved effect of ES on nerve regeneration. Future clinical studies should evaluate the usage of subcutaneous injection of lidocaine post ES for analgesia control.

Introduction

Peripheral nerve injuries are common, occurring in approximately 3.8% of extremity trauma patients. ¹ These injuries range from compressions to complete transections, all of which result in functional deficits, with traumatic injuries causing the most severe impairments. ² Severe injuries, such as nerve transections, have a limited potential for full functional recovery.^2,3 While peripheral nerves have intrinsic regenerative capacity, partial restoration of function leaves permanent deficits in most cases.^2,3 Currently, the standard of care for lacerated nerves is direct tension-free microsurgical repair. ⁴ Various strategies have been explored to enhance peripheral nerve regeneration, with brief 1-hour electrical stimulation (ES) emerging as the most clinically translatable approach.^5,6 This therapeutic modality has shown benefit both preclinically and clinically in animals and humans, respectively. It has been postulated that with the activation of injured axons via ES, the action potentials traveling to the neuronal cell bodies in the spinal cord or dorsal root ganglia result in an upregulation of regeneration-associated genes that ultimately lead to enhanced axon elongation.^7-13

Local anesthetics are commonly used in surgical procedures and can block action potential conduction, particularly when ES is applied near the site of administration. ⁹ To mitigate this issue, ES can be applied under general anesthesia before administering local anesthetics for postoperative pain control. In cases where a surgery is performed entirely under local block, a patient may wait until the block wears off before the implementation of the therapy, as demonstrated by Coroneos et al. ¹⁴ Recently, Keane et al ¹⁵ showed that administering local lidocaine after the application of very short duration (10 minutes) of therapeutic ES, abolished the effect of the therapy as measured through histological analysis. Although Keane et al hypothesized that lidocaine would have no impact, their findings suggest that postoperative lidocaine should be avoided following ES, potentially limiting the clinical feasibility of this therapy. However, a critical examination of the dosing of lidocaine used in the Keane study is important and the concentration used may have had an impact.

This study examined the use of postoperative local anesthesia on the regenerative effects of ES as measured by retrograde labeling of regenerated neurons. We hypothesized that subcutaneous postoperative local anesthesia administered after ES therapy would not impair its regenerative effects, if ES-induced retrograde depolarization of the neuronal soma occurs prior to lidocaine-induced conduction block.

Methods

Animals

A total of 40 adult male Lewis rats (weight 250-350 g) were used and were equally divided between the groups (n = 10 per group). The number of rats selected was based on previous work by the authors and other studies using retrograde labeling techniques. ⁹ This strain of rats were chosen for experiments because they showed the lowest level of auto-mutilation following nerve transection injuries. ¹⁶ Rats were housed in temperature-controlled rooms, under standard 12-h light cycle and given standard rat chow and water ad libitum. Experimental procedures were approved by the Animal Care Committee at McMaster University and were performed according to guidelines set out by the Canadian Council on Animal Care.

Experimental Groups

All rats underwent tibial nerve transection and immediate repair. Rats were randomly assigned into 4 groups: (1) direct repair only (REP); (2) direct repair followed by lidocaine application (REP + LIDO); (3) 60 minutes of ES (60 ES); and (4) 60 minutes of ES followed by lidocaine application (60 ES + LIDO).

Surgical Procedures

All surgical procedures were performed aseptically under 2% isoflurane anesthetic. All rats undergoing nerve injury and repair underwent the same surgical procedures, with the right hindlimb serving as the experimental limb. A lateral thigh incision was made to expose the tibial nerve. The biceps femoris muscle was then dissected from the femur, and the tibial nerve was isolated from surrounding connective tissue. The nerve was transected 1 mm distal to the trifurcation of the nerve using microscissors, and the 2 tibial nerve stumps were immediately reconnected using 2 Nylon 9-0 epineurial sutures under 10× surgical microscope magnification.

ES Paradigm

In groups receiving ES, stimulation was delivered using the PeriPulse nerve stimulation system (Epineuron Technologies Inc., Ontario, Canada) for 60 minutes. An introducer tool was used to create a subcutaneous tunnel to feed the electrode lead through. The electrode lead was then placed next to the repaired tibial nerve. This method of placement combined with the material properties of the electrode lead prevents it from inadvertently migrating from the site of application. The electrode lead was then connected to the stimulator, and the appropriate ES therapy was applied. Stimulus parameters included charge balanced constant current stimulation at 20 Hz with 100 µs rectangular pulse widths and stimulus amplitudes (1.5-2.5 mA) titrated to maximize dorsiflexion via volume conduction and activation of the uninjured common peroneal nerve.

Lidocaine Application

Immediately following ES therapy, 0.15 to 0.20 mL of 1% lidocaine hydrochloride, depending on rat weight, was injected directly into the submuscular space as a tibial nerve block after closure of the biceps femoris using 3-0 Vicryl sutures. The dose was calculated in accordance with McMaster University’s Standard Operating Procedures for rodent dosing guidelines, ensuring it did not exceed 7 mg/kg of body weight. Our institution’s framework is based on the University of British Columbia’s Standard Operating Procedure on local analgesia/anesthesia in adult rodents. ¹⁷ This approach was selected to ensure effective nerve blockade and consistency across specimens, while maintaining proper retention within the surgical site. For rats weighing 250 to 350 g (0.25-0.35 kg), the maximum allowable dose of lidocaine is 1.75 mg (for a 250 g rat) to 2.45 mg (for a 350 g rat). For 1% lidocaine hydrochloride (10 mg/mL), this equates to an allowable volume of 0.175 mL (for a 250 g rat) to 0.245 mL (for a 350 g rat). Hence, our administered volumes of 1% lidocaine hydrochloride (0.15-0.20 mL) for 250 to 350 g rats are below the maximum allowable dosage. Following lidocaine application, the skin incision was closed using 3-0 Vicryl sutures.

Retrograde Labeling

All animals underwent a retrograde labeling surgical procedure to label regenerated motor neurons 28 days following initial nerve injury. This time duration was selected to allow a sufficient number of regenerated axons to cross the labeling site as the regeneration process is staggered and typically more time is needed for all regenerated axons to cross the surgical repair site as discussed by Al-Majed et al. ⁹ Surgical procedures were performed aseptically under anesthetic (2% isoflurane). The tibial nerve was exposed, and a ruler was used to measure 10 mm from the site of nerve repair. The tibial nerve was transected at this location and the proximal portion of the nerve labeled with Fluoro-Ruby using a standard well technique. ¹⁸ Briefly, the transected proximal tibial nerve stump was placed in a 5 × 5 mm petroleum jelly well that was constructed on top of a piece of parafilm. The well was filled with 10 µL of 8% solution of Fluoro-Ruby dextran (ThermoFisher Catalog number: D1817 or equivalent) dissolved in sterile saline solution (Figure 1). The nerve was left within the well for 1 hour and then the proximal stump and wound rinsed 3 times with sterile saline to remove any residual dye and prevent staining of the adjacent muscle or nerve. The wound was then closed with 3-0 Vicryl suture for muscle and skin, respectively, as before.

Figure 1.

Retrograde labeling of motor and sensory neurons by placing the distal tibial nerve stump, cut 10 mm from the nerve repair site, in a well filled with Fluoro-Ruby for 1 hour.

Tissue Harvest

Seven days following the retrograde labeling procedure, the animals were deeply anesthetized using sodium pentobarbital and transcardially perfused with saline followed by 4% paraformaldehyde. The lumbar spinal cords were then harvested and placed in 4% paraformaldehyde solution for 24 hours. Cryoprotection was performed by transferring the tissue to a 20% sucrose solution in 4% paraformaldehyde, and spinal cords were placed in cryomolds and the molds filled with optimal cutting temperature (OCT) compound. The tissue was then frozen over a liquid nitrogen bath. Serial sections were cut at 40 µm thickness for both spinal cord and dorsal root ganglia tissue using a cryostat (Leica CM 3050 S) and placed on glass slides.

Analysis

Labeled motor and sensory neurons were counted on each slide by a blinded observer using a LEICA CTR6000 fluorescent microscope. A correction factor of 0.45 was applied to all counts according to Abercrombie. ¹⁹ Representative slides were then chosen from each group and imaged using a ZEISS LSM980 confocal microscope at 10× magnification. Postprocessing was performed in ImageJ (version 1.54f) and involved adjustment for contrast and brightness to reduce background. A 5 pixel mean filter was applied to reduce noise.

Statistical Methods

All statistical analysis was performed in GraphPad Prism Version 10. Values are represented as mean ± standard deviation. Motor neuron counts were analyzed using a 1-way analysis of variance (ANOVA) with a post-hoc Tukey analysis to control for multiple comparisons. Statistical significance was set to P < .05.

Results

One rat in the 60ES + LIDO group did not survive the retrograde labeling surgery. Some samples were lost during dissection or tissue sectioning and sample sizes are reflected in the figures.

Lidocaine Does Not Impair Nerve Regeneration

A clinically relevant dose of lidocaine hydrochloride applied directly on the nerve did not impair motor nerve regeneration in rats that had their tibial nerve immediately cut and repaired (Figure 2). Motor neuron counts in rats that had their tibial nerves immediately repaired (REP) were 331 ± 26 compared to 295 ± 44 for rats that had lidocaine applied immediately after the nerve repair was performed (REP + LIDO), a difference that was not statistically significant (P = .63).

Figure 2.

Labeled motor neuron counts 28 days after nerve injury for rats that underwent immediate nerve repair (REP, n = 8), immediate nerve repair followed by lidocaine application (REP + LIDO. n = 10), electrical stimulation for 60 minutes (60 ES, n = 8), and electrical stimulation for 60 minutes followed by lidocaine application (60 ES + LIDO, n = 9).

Note. REP = immediate nerve repair; REP + LIDO = immediate nerve repair followed by lidocaine application; 60 ES = electrical stimulation for 60 minutes; 60 ES + LIDO = electrical stimulation for 60 minutes followed by lidocaine application.

Levels of statistical significance are represented by asterisks with *P < .05. **P < .01. ***P < .001. ****P < .0001.

Direct application of lidocaine hydrochloride also did not impair nerve regeneration of sensory nerves when assessing the dorsal root ganglion neuron counts of these rodents (Figure 3). Dorsal root ganglion neuron counts in rats that had their tibial nerves immediately repaired (REP) were 467 ± 147 compared to 455 ± 51, for rats that had lidocaine applied immediately after the nerve repair was performed (REP + LIDO), a difference that was not statistically significant (P = .99).

Figure 3.

Labeled sensory neuron counts within the dorsal root ganglia 28 days after nerve injury for rats that underwent immediate nerve repair (REP, n = 8), immediate nerve repair followed by lidocaine application (REP + LIDO, n = 8), electrical stimulation for 60 minutes (60 ES, n = 8), and electrical stimulation for 60 minutes followed by lidocaine application (60 ES + LIDO, n = 7).

Note. REP = immediate nerve repair; REP + LIDO = immediate nerve repair followed by lidocaine application; 60 ES = electrical stimulation for 60 minutes; 60 ES + LIDO = electrical stimulation for 60 minutes followed by lidocaine application.

Levels of statistical significance are represented by asterisks with *P < .05. **P < .01. ***P < .001. ****P < .0001.

ES Therapy Is Not Impacted by Postoperative Lidocaine Application

Rats that underwent tibial nerve repair followed by 60 minutes of ES (60 ES) had a mean motor neuron count of 543 ± 74, compared to 539 ± 89 in those that received ES for 60 minutes followed by lidocaine application (60 ES + LIDO) (Figure 2). This difference was not statistically significant (P = .99). However, both 60 ES groups showed significantly greater motor neuron counts than the repair-only group, with increases of 188 to 273 neurons (63%-83%, P < .0001).

Similarly, when assessing the dorsal root ganglion neuron counts, rats that underwent tibial nerve repair followed by 60 minutes of ES (60 ES) had a mean sensory neuron count of 1413 ± 152 compared to 1365 ± 72 in those that received 60 ES followed by lidocaine application (Figure 3). This difference was not statistically significant (P = .43). However, both 60 ES groups showed significantly greater sensory neuron counts than the repair-only group, with increases of 854 to 1007 neurons (200%-202%, P < .0001).

Representative longitudinal spinal cord sections cut at 40-µm thickness showing labeled motor and sensory neurons from each group are shown in Figures 4 and 5, respectively.

Figure 4.

Representative longitudinal sections of rat spinal cord cut at 40 µm thickness with Fluro-Ruby labeled motor neurons.

Note. REP = immediate nerve repair; REP + LIDO = immediate nerve repair followed by lidocaine application; 60 ES = electrical stimulation for 60 minutes; 60 ES + LIDO = electrical stimulation for 60 minutes followed by lidocaine application.

Figure 5.

Representative longitudinal sections of rat dorsal root ganglia cut at 40 µm thickness with Fluro-Ruby labeled sensory neurons.

Note. REP = immediate nerve repair; REP + LIDO = immediate nerve repair followed by lidocaine application; 60 ES = electrical stimulation for 60 minutes; 60 ES + LIDO = electrical stimulation for 60 minutes followed by lidocaine application.

Discussions

The use of brief 1-hour ES in peripheral nerve injury has been shown to significantly enhance nerve regeneration and is widely regarded as a therapy with minimal barriers to clinical translation. Notably, 4 randomized clinical trials using 1-hour of ES have demonstrated substantial improvements in both sensory and motor outcomes across various injury types, including compression, traction, and transection.^10-13 In these studies, ES therapy was administered either intraoperatively or postoperatively, with most surgeries performed under general anesthesia, avoiding preoperative nerve blocks or local anesthesia that could compromise the therapeutic efficacy of the ES treatment.

The underlying mechanism of action of ES therapy is thought to be mediated through retrograde action potential conduction that targets the cell bodies of injured nerves resulting in a BDNF-mediated response. ²⁰ Local anesthetics are largely sodium channel blockers that prevent the conduction of action potentials and thus impede the therapeutic benefit of ES. Geremia et al ⁷ showed that using tetrodotoxin (TTX) to block nerve action potentials proximal to the site of stimulation, thus preventing APs from reaching the cell body, abolishes the treatment effect. Ward et al ²¹ corroborated this finding using optical stimulation instead of ES in thy-1-ChR2/YFP mice. In these rodents, nerves expressing channelrhodopsin (ChR2) are light sensitive and can become activated using blue light. This provides the advantage of investigating the effects of selective activation of nerves instead of the traditional use of electric fields which may influence neighboring structures such as glial cells. Ward et al’s conclusion was that the direct activation of the axons was key to eliciting the pro-regenerative effect of stimulation.

Postoperative pain management is required following surgical procedures, and peripheral nerve blocks have demonstrated efficacy in reducing postoperative pain and decreasing opioid use. ²² Thus, to ensure successful clinical translation of ES therapy, the effects of postoperative local anesthesia on ES should be elucidated. The dose of local anesthesia is of critical importance as amounts over the defined safe and tolerable range may lead to toxicity.^23,24 For postoperative peripheral nerve blocks and pain management purposes, the recommended maximum allowable dose range of lidocaine in humans is 4.5 to 7 mg/kg.^23,25 Generally, the maximum allowable dose of lidocaine in rats is also 7 mg/kg.^17,26 A recent study by Keane et al ¹⁵ described that the application of lidocaine after delivering ES therapy can abolish the effect of stimulation-mediated regeneration. However, the applied dose was 3 mL of 2% lidocaine for 5 minutes, which was then irrigated. For rats weighing 250 to 350 g, this dose equates to 171 to 240 mg/kg of lidocaine administered, which is approximately 24 to 34 times the maximum allowable limit of 7 mg/kg. In our study, we applied a clinically relevant dose of 0.15 to 0.20 mL of 1% lidocaine to maintain approximately 6 mg/kg of lidocaine administered per rat, which is well within the allowable limits to ensure a nontoxic dose is administered. Furthermore, the dose was administered directly into the submuscular space surrounding the nerve, rather than irrigating the nerve followed by a washout, as this better reflects what is typically performed for postoperative pain management in the clinical setting. However, some differences exist between our study and Keane et al, namely the outcome measures employed to assess nerve regeneration and the stimulation parameters. Keane et al used histological methods to assess regeneration which captures newly regenerated axons along with their collateral sprouts. In this article, we used a more direct method to label regenerated axons where collateral sprouts from a parent axon do not add to axon counts. With respect to the stimulation parameters, Keane et al used a 16-Hz constant current charge balanced waveform, whereas in the current experiment, we used a 20-Hz constant current charge balanced waveform.

Nonetheless, in line with our hypothesis, our results demonstrate that lidocaine administered postoperatively and after ES therapy does not affect enhanced nerve regeneration from ES therapy. It may be that the initiation cascade for the upregulation of key neurotrophic and regeneration-associated factors, including brain-derived neurotrophic factor (BDNF), Tα1-tubulin, and GAP-43, would have been triggered before lidocaine exerts any inhibitory effects on the regeneration process itself. Some studies have shown that BDNF mRNA is increased following mechanical damage as early as 15 minutes following the stimulus. ²⁷ Future work may explore the dynamics related to the neuron and surrounding glial cells following ES therapy. Some limitations of this study include the lack of control groups to demonstrate the efficacy of the application of lidocaine in blocking nerve conduction. These results provide preliminary data to support clinical utility of integrating ES therapy with current postoperative pain management techniques. As preoperative regional anesthetic peripheral nerve block techniques, such as wide-awake local anesthesia no tourniquet (WALANT), become more heavily adopted as standards of care, future work should include studying how ES therapy may be integrated into these surgical workflows (stimulator placement intraoperatively and ES following recovery from regional block).

Conclusions

Electrical stimulation therapy has clear evidence to support its ability to enhance peripheral nerve regeneration. In this study, we demonstrate that the beneficial effects of ES therapy are not abolished with the administration of postoperative local anesthesia allowing surgeons to benefit from implementing ES therapy within their existing postoperative pain management protocols.