SCIENTIFIC ARTICLE
Short-Term Electrical Stimulation to Promote Nerve Repair and Functional Recovery in a Rat Model Colleen Calvey, MD, Wenda Zhou, MS, Kimberly Sloan Stakleff, PhD, Patricia Sendelbach-Sloan, BS, Amy B. Harkins, PhD, William Lanzinger, MD, Rebecca Kuntz Willits, PhD
Purpose To evaluate the effect of duration of electrical stimulation on peripheral nerve regeneration and functional recovery. Based on previous work, we hypothesized that applying 10 minutes of electrical stimulation to a 10-mm rat sciatic nerve defect would significantly improve nerve regeneration and functional recovery compared with the non-electrical stimulation group. Methods A silicone tube filled with a collagen gel was used to bridge a 10-mm nerve defect in rats, and either 10 minutes or 60 minutes of electrical stimulation was applied to the nerve during surgery. Controls consisted of a silicone tube with collagen gel and no electrical stimulation or an isograft. We analyzed recovery over a 12-week period, measuring sciatic functional index and extensor postural thrust scores and concluding with histological examination of the nerve. Results Functional assessment scores at week 12 increased 24% in the 10-minute group as compared to the no stimulation control group. Electrical stimulation of either 10 or 60 minutes improved the number of nerve fibers over no stimulation. Additionally, the electrical stimulation group’s histomorphometric analysis was not different from the isograft group. Conclusions Several previous studies have demonstrated the effectiveness of 60-minute stimulations on peripheral nerve regeneration. This study demonstrated that an electrical stimulation of 10 minutes enhanced several functional and histomorphometric outcomes of nerve regeneration and was overall similar to a 60-minute stimulation over 12 weeks. Clinical relevance Decreasing the electrical stimulation time from 60 minutes to 10 minutes provided a potential clinically feasible and safe method to enhance nerve regeneration and functional recovery. (J Hand Surg Am. 2015;40(2):314e322. Copyright Ó 2015 by the American Society for Surgery of the Hand. All rights reserved.) Key words Peripheral nerve regeneration, electrical stimulation, rat sciatic nerve, direct current.
Received for publication July 18, 2014; accepted in revised form October 2, 2014.
Akron, Margaret F. Donovan Endowed Chair for Women in Engineering. The authors acknowledge Amanda Pinheiro for her assistance extracting the walking track video and Kelly Bonder, BS, Bethany Noble, BS, and Marcus Dempster for their technical support during surgeries. Histology was provided by Jason Papke, BS, and Dr. Jan Ryerse at Saint Louis University.
The present address of K.S.S. is Department of Urology, Akron General Medical Center, Akron, OH.
No benefits in any form have been received or will be received related directly or indirectly to the subject of this article.
The present address of P.S.-S. is Kent State University, Research Compliance, Kent, OH.
Corresponding author: Rebecca Kuntz Willits, PhD, Department of Biomedical Engineering, The University of Akron, 260 S. Forge St., OLRC 301, Akron, OH 44325-0302; e-mail: [email protected].
From the Department of Orthopaedic Surgery and the Kenneth Calhoun Research Laboratory, Akron General Medical Center, Akron; the Department of Biomedical Engineering, The University of Akron, Akron, OH; and the Department of Pharmacological and Physiological Science, Saint Louis University, St. Louis, MO.
The present address of C.C. is University of Louisville, Christine M. Kleinert Institute for Hand & Microsurgery, Louisville, KY. Funding for this study was provided by Akron General Medical Center, Akron General Development Foundation, and the Department of Orthopedics, as well as The University of
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0363-5023/15/4002-0018$36.00/0 http://dx.doi.org/10.1016/j.jhsa.2014.10.002
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frequent and costly occurrence, resulting from traumatic, compressive, and demyelinating disease mechanisms. Peripheral nerve injuries of the upper and lower extremities occur in about 3% of trauma patients,1 with an estimated incidence of digital nerve injuries in 6.2/100,000 patients.2 Spontaneous nerve regeneration occurs at an estimated rate of 1 mm/d, and it is often complicated by inhibitory growth factors, Wallerian degeneration, neuroma formation, and disorganized growth.3 Currently, treatments such as autografts, allografts, or nerve guidance conduits provide a scaffold to help guide regeneration, but do not affect the rate of recovery, and outcomes can be unpredictable. Safe and efficacious tools that can be coupled with currently available treatments to improve outcomes can have an important impact on nerve repair clinically. The nerve is analogous to an electrical cable, with an electrically active extracellular environment generated and maintained by ionic gradients and pumps. Externally applied electrical stimulation (ES), therefore, has been examined as a method to improve nerve regeneration. ES upregulates nerve growtheassociated genes, which have been found to correlate with acceleration of axonal regeneration.4,5 Additionally, a number of in vitro and in vivo studies have examined the ability of ES to enhance nerve repair and regeneration, maintain nerve plasticity, and improve functional recovery.6e9 The ES parameters that are required to provide improved outcomes are unknown, however. Applying ES of approximately 1 hour at the time of surgical repair can enhance nerve recovery.9e13 Given the high cost of operating-room time,14 however, an ES period of 60 minutes would be impractical and costprohibitive in a wide clinical setting. In previous in vitro studies, researchers found that environmental changes such as media composition, calcium ion levels, temperature, and stimulation duration (10 vs 100 min), had minimal impact on neurite outgrowth results. Any ES resulted in a 40% increase in neurite length compared with nonstimulated controls after 10 minutes of stimulation.15,16 Therefore, in this study, we examined whether 10-minute ES would also have a significant impact on nerve regeneration in vivo. ERIPHERAL NERVE INJURIES ARE A
METHODS Study design and ES The Akron General Medical Center institutional review board and institutional animal care and use committee approved all animal surgical and experimental procedures. Rats were divided into 4 groups: 10-minute ES (N ¼ 10), 60-minute ES (N ¼ 11), no ES J Hand Surg Am.
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control (N ¼ 10), and isograft control (N ¼ 10). In the first 3 groups, a 10-mm sciatic nerve gap was created and a silicone tube filled with 1.0 mg/mL collagen was used to bridge the defect. After entubulation, 0 (no stimulation group), 10 (10-min group), or 60 (60-min group) minutes of ES were applied as appropriate for the group. In the isograft group, the nerve defect was bridged by sciatic nerve retrieved from donor rats. In the stimulation groups, looped platinum wire electrodes were placed at the proximal and distal nerve segments after repair. A direct current of 24 V/m (24 mV, w1.5mA) was applied across the electrodes and run for 10 or 60 minutes. On the contralateral, non-experimental side, sham electrodes without electrical current were applied across a 10-mm segment of sciatic nerve. The exposed tissues were kept moist with sterile saline soaked gauze during the ES. The rats were then given food and water ad libitum and housed in pairs with a 12-hour lightedark cycle. Footprint recoding and sciatic functional index calculation Rats were made to traverse a walking track every 2 weeks where a digital camera captured their footprints.17 The rat feet were marked with black ink using a marker. Video was captured for at least 3 walking cycles, and when the rat took 5 continuous steps across the middle of the track and paws were not contracted, the image would be used for sciatic functional index (SFI) analysis. SFI was used to evaluate the sciatic nerve functional recovery every 2 weeks after surgery until the animals were killed. SFI was determined using the following equation as introduced by de Medinaceli and adapted by Bain,18 where: EPL NPL ETS NTS SFI ¼ 38:3 þ 109:5 NPL NTS EIT NIT þ 13:3 8:8 NIT
In this equation, EPL is distance from the heel to the top of the third toe in the injured foot, NPL is the same distance in the non-injured foot; ETS is distance between the first and the fifth toe in the injured foot, NTS is the same distance in the non-injured foot; EIT is distance from the second to the fourth toe in the injured foot, and NIT is the same distance in the noninjured foot. In SFI analysis, 4 extracted images of clear footprints were randomly chosen at each time point per rat, and ImageJ (NIH, Bethesda, MD)19 was used to measure the appropriate distances. An average of 4 measurements was then used to calculate the SFI value. These measurements were blinded by Vol. 40, February 2015
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FIGURE 1: Biweekly SFI for the different groups was collected as a measure of post injury functional recovery (N ¼ 7). * P < .05 for the measurement at week 12 relative to week 2. # P < .05 for the 10-minute electrical stimulation group versus the no stimulation control at week 12.
FIGURE 2: Biweekly EPT for different groups was collected as a measure of post injury functional recovery (N ¼ 10). * P < .05 between week 2 and week 12. # P < .05 between week 2 and week 10.
analyzing each video by date and then combined to the appropriate sample group. Extensor postural thrust measurement Extensor postural thrust (EPT) score was used as a separate evaluation of motor performance and measured on the same days as the SFI.17 EPT is classified as a postural reflex reaction and previous J Hand Surg Am.
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research has shown high correlation between SFI and EPT.20 The body of the rat was wrapped in a surgical towel with the hind limbs extending out. The animal was then placed over the platform of a digital balance. Once the hind limb made contact of the digital metatarsus, the hind limb was kept in contact with the platform for 30 seconds and the maximum force of the hind limb pushing against the platform was Vol. 40, February 2015
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FIGURE 3: The effect on the gastrocnemius muscle was examined by A box plot of weight for each sample, and histology images for B contralateral side, C no stimulation, D 10-minute electrical stimulation, E 60-minute electrical stimulation, and F isograft. The distance between the nuclei indicated the size of skeletal muscle cell. Cells in no stimulation control, 10-minute, and 60-minute groups were smaller than the isograft group, as the nucleus density was higher in those groups. Scale bar ¼ 50 mm.
recorded. The procedure was repeated 5 times and the 3 largest forces were used for analysis. Histomorphometric evaluation At the end of 12 weeks, the sciatic nerves were excised and segments were collected at the middle of the tube and 2 mm distal to the end of the silicone tube. The nerve segments were fixed in paraformaldehyde, epoxy-embedded, sectioned at 1 mm, and stained with toluidine blue. For each section, a 20x light microscope was used to capture the fascicular area. An entire nerve image was created and the perineurium edge was outlined using the software to measure total fascicular area. The number of nerve fibers and fiber area were then measured as described previously.21 Briefly, images were taken from 3 random regions per section. A specified area was processed in ImageJ to count the number of nerve fibers and measure the percentage of the area that was nerve fibers. The number of nerve fibers and percent fiber areas were J Hand Surg Am.
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calculated from a total of at least 15 images per group. Nerve fiber density was calculated by scaling the percentage of nerve fibers to the total intrafascicular area. The mean nerve fiber area was calculated by dividing nerve fiber number by total nerve fiber area. The mean fiber width was determined by relating the mean nerve fiber area to the area of a circle. In addition to the nerve, the gastrocnemius muscle was dissected from the injured side. The muscle samples were weighed and then dehydrated, embedded in paraffin, and stained with hematoxylin and eosin. A 40x light microscope was used to capture the muscle histology image. Statistics All data are expressed as mean standard deviation. Prior to institutional animal care and use committee approval, a power analysis was performed using histomorphometric data from the literature, with power ¼ 0.9 and significance level of .05, to set the Vol. 40, February 2015
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FIGURE 4: Sections of regenerated sciatic nerve at the middle of the conduit 12 weeks after surgery for the animals receiving A isograft, B no stimulation control, C 10-minute electrical stimulation, and D 60-minute electrical stimulation groups. Blood vessels (BV) were found on all groups that had a conduit. Scale bar ¼ 20 mm.
number per group at N ¼ 10.21 Comparisons within and between no stimulation, 10-minute, and 60minute groups were analyzed with one-way ANOVA with the Tukey post hoc test. Each group was then compared to the isograft with a t test. Values of P < .05 were considered statistically significant. RESULTS Functional evaluation SFI: No significant difference in SFI was found between any time points in the no stimulation group. After 12 weeks, SFI for 10-minute, 60-minute, and isograft groups were significantly higher than their respective SFI at the 2-week time point (P ¼ .002, P ¼ .02, P ¼ .01, respectively). When comparing within each group in the same time point, no significant differences were found in the early weeks (Fig. 1). SFI in the 10-minute group was significantly improved by 24% compared with the no stimulation group at week 12 (P ¼ .03). At the 12-week time point, there was no significant difference between the 10-minute and 60-minute groups (P ¼0.2) or between the ES groups and the isograft group. EPT: Similar to SFI analysis, no significant difference was found in EPT between time points for the no stimulation control (Fig. 2). EPT in the 10-minute group gradually increased with increasing recovery time. At 12 weeks, rats from the 10-minute group had J Hand Surg Am.
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a significantly higher EPT force than at the earlier time points of 2, 4, 6, and 8 weeks (P < .04). No significant difference was found between the 10- and 60-minute groups at each time point. Histomorphometric evaluation Muscle weight and histology: The extracted gastrocnemius muscle weights (in g) on the experimental side were not significantly different between the no ES control, 10-minute, and 60-minute groups. The 10-minute and 60-minute groups were significantly lower than the isograft group (P ¼ .035, P ¼ .001 respectively, Fig. 3). From the histology of gastrocnemius muscles, cell atrophy was visible in the 10-minute and 60minute groups compared with the isograft group and normal sciatic nerve. Number of nerve fibers: The overall histology of the nerves is shown in Figure 4. There was no significant difference in nerve samples from the midline of injury between the no stimulation control, 10-minute, and 60-minute groups, although they were all significantly lower than the isograft group (P < .003; Fig. 5). In the distal nerve sample, the average number of nerve fibers from the 10-minute (P ¼ .03) and 60-minute (P ¼ .026) groups was significantly higher than the no stimulation control (Fig. 6). No significant difference was found between either ES groups or the isograft group in number of distal Vol. 40, February 2015
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FIGURE 5: Histomorphometric data collected at the midline of the conduit (N 6). The data were quantified from the images for A number of nerves, B percent nerve area, C mean fiber width, and D fiber density. * P < .05 versus no stimulation control group.
nerves, but the no stimulation control group was significantly lower than the isograft group (P ¼ .013). Area of nerve fiber: The quality of nerve regeneration is represented by the percentage area of nerve fiber and was significantly higher in the 60-minute group at both the midline (Fig. 5) and distal (Fig. 6) nerve samples than the no stimulation control (P ¼ .04 and P ¼ .038, respectively). The 10-minute group did not show a significant difference compared with the no stimulation control. No significant differences existed between the 10- or 60-minute groups and the isograft group at either the midline or the distal nerve sample, but the no stimulation controls were significantly lower than isograft groups at both the midline (P ¼ .005) and the distal (P ¼ .003) nerve samples. Mean fiber width: Mean fiber width (Figs. 5, 6), an indirect measure of myelination and potentially mature nerve fibers, was not statistically different between the ES groups, although they were significantly higher than the no stimulation control at the midline nerve samples (P ¼ .006 and P ¼ .002, respectively). When compared with the isograft group, the no stimulation control was significant lower than the isograft group (P ¼ .016) at the midline, and 10minute and 60-minute groups were not statistically different from isograft group. Density: Although there was no significant difference between the 10-minute and 60-minute groups and no J Hand Surg Am.
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stimulation control at the midline, the density of the 60-minute group was significantly higher than the no stimulation control at the distal end (P ¼ .015) (Fig. 6). The 10-minute and 60-minute groups were not statistically different from the isograft group (Figs. 5, 6), whereas the no stimulation control was significantly lower than the isograft (P ¼ .005) at the distal end. DISCUSSION The application of ES to nerves provides an additional tool to promote nerve regeneration and functional recovery after nerve injury. Researchers have examined several ES parameters for one hour’s duration since 2000, when Gordon and colleagues demonstrated that application of ES for 1 hour was as effective as weeks of ES to promote nerve regeneration.8 For example, 1 hour of ES using pulsed direct current (square 0.1-ms pulses [4V] at 20 Hz) accelerated motor functional recovery in a larger rat nerve defect (15 mm).22 Additionally, changing the frequency of an alternating current stimulation (1 mA) impacted nerve regeneration.23 We extend these previous reports by demonstrating that application of direct current, rather than alternating current or pulsed direct current, similarly improved sciatic outcome parameters in rats after ES of 60 or 10 minutes. The low current applied here, which is 100 to 1,000 times lower than applied for TENS24,25 and Vol. 40, February 2015
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FIGURE 6: Histomorphometric data collected at 2 mm distal to the repair conduit (N 5). The data were quantified from the images for A number of nerves, B percent nerve area, C mean fiber width, and D fiber density. * P < .05 versus the no stimulation control.
lower than previous studies,26 reduces the time and improves the safety of application while not reducing the efficacy. Although low degrees of late-onset injury or axonal degeneration have been associated with ES (up to 8 hours, 50 Hz, and 2100e4500 mA),27e29 longer durations of ES would increase surgical risks due to increased time under anesthesia and risk of infection. In addition, increased time in the operating room increases the costs associated with implementing a new procedure. Improving safety and reducing cost while maintaining efficacy are important benefits of decreasing the duration of ES. In an effort to mitigate safety concerns with the application of 60 minutes of ES, researchers have investigated the effect of reducing the duration of ES. A 30-minute stimulation (20-Hz pulse rate, 2mA amplitude) promoted nerve regeneration after a nerve crush injury in a rat model.30 After 3 weeks, SFI increased by 28% in 30-minute stimulation groups compared with controls.30 Although this result was promising, the outcomes of interventions after crush injury do not always match the outcome after transection injury. In a 5-mm transection, a 20-minute stimulation (3V, 20 Hz) accelerated nerve regeneration in delayed nerve injury repair rat model.31 Using only 10 minutes of stimulation, we saw improved SFI and EPT functional recovery measures, demonstrating efficacy. With the equation used to calculate J Hand Surg Am.
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SFI, a value close to zero indicates that the rat’s toe spread is equivalent on both injured and non-injured side, and a value close to e100 indicates total impairment. After 1 year, functional recovery after sciatic nerve repair with optimal transection conditions is limited to 41% of function, or a score of e41.32 After 12 weeks, our SFI scores approached 50%. Compared with the no stimulation control, the 10-minute stimulation increased the SFI by 24% after 12 weeks, and SFI in the 60-minute group was increased by 13%. This improvement in functional recovery over shorter recovery time frames was comparable to the crush injury results,30 as well as results of studies after 60 minutes of stimulation.33 Although the effectiveness of short-duration stimulation is controversial,3 both our functional recovery and histomorphometric results showed that 10 minutes of stimulation was similar to 60 minutes of stimulation. The histological sections provided several evaluations of nerve recovery. The blood vessels found in the midline of conduits would provide nutrition to regrowing nerves (Fig. 4). The mean fiber width, which represents the maturity of regenerating nerve fibers, indicated that the 10-minute stimulation was as good as a stimulation of 60 minutes. The density of nerve fibers in the 60-minute group was also greater than no stimulation control, which is consistent with previous literature.34 No Vol. 40, February 2015
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significant differences were found between the stimulation groups and the isograft group in number of distal nerves, area of nerve percent, and fiber density at distal end, whereas the no stimulation group was significantly lower than isograft group on those parameters. This result suggests that the distal nerves are better sustained with ES than without, making the group similar to an isograft. Although the isograft group had an increased number of nerves at the midline, this result would be expected as the isograft itself contains nerve fibers. At the midline, the area of nerve fibers for the 60-minute group and the mean fiber width of both stimulation groups were significantly higher than the no stimulation control, suggesting that ES could enhance the quality and maturity of nerve regeneration. Overall, we did not find any significant differences between the 10- and 60-minute groups in the histomorphometric parameters studied, suggesting that a 10-minute stimulation may suitably replace a 60-minute stimulation. There were several limitations to the current study. A collagen gel was placed into the conduit prior to implantation, and collagen offers a suitable scaffold for the regeneration of axons.35 Therefore, synergies between the mechanism of ES and growth within a collagen tube should be further explored. The weight of the gastrocnemius muscle indirectly evaluates nerve regeneration.36 In this study, we found the muscle weight in the isograft group was higher than the other groups. Although this result was similar to other studies,37,38 it would be important to determine the cause of these weight differences. Previous studies have shown that ES supports motor neuron recovery and survival.8,11,39 The sciatic nerve is a mixed sensory and motor nerve, yet functional recovery, as measured here, was dependent on motor neuron survival and regeneration. Future experiments should be directed at examining sensory nerve recovery. Toe contractures were found in all groups, reducing the number of animals analyzed from 10 to 7 in SFI. Although SFI reliability may be compromised by removal of animals, it was not the case in this study as shown by significance found at week 12 with a 10minute stimulation. Because contractures indicate a lack of physical therapy rather than a lack of function,40,41 SFI is still a useful noninvasive tool to measure functional recovery.42 Daily exercise training should be used to increase the functional recovery and SFI numbers.43 In this study, histomorphometric samples were lost due to damage of some embedding process, thereby reducing the numbers as noted in the results. Post hoc power analysis was used to confirm any significant differences. J Hand Surg Am.
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Despite these limitations, our findings suggest that 10 minutes of ES enhanced nerve regeneration and functional recovery compared with no ES. Decreasing the duration of stimulation has made this method a safe, efficient, and feasible tool for future clinical use. REFERENCES 1. Kang JR, Zamorano DP, Gupta R. Limb salvage with major nerve injury: current management and future directions. J Am Acad Orthop Surg. 2011;19(Suppl 1):S28eS34. 2. Thorsen F, Rosberg HE, Steen Carlsson K, Dahlin LB. Digital nerve injuries: epidemiology, results, costs, and impact on daily life. J Plast Surg Hand Surg. Sep 2012;46(3e4):184e190. 3. Gordon T, Sulaiman O, Boyd JG. Experimental strategies to promote functional recovery after peripheral nerve injuries. J Peripher Nerv Syst. 2003;8(4):236e250. 4. Al-Majed AA, Brushart TM, Gordon T. Electrical stimulation accelerates and increases expression of BDNF and trkB mRNA in regenerating rat femoral motoneurons. EurJ Neurosci. 2000;12(12):4381e4390. 5. Geremia NM, Gordon T, Brushart TM, Al-Majed AA, Verge VM. Electrical stimulation promotes sensory neuron regeneration and growth-associated gene expression. Exp Neurol. 2007;205(2): 347e359. 6. English AW, Schwartz G, Meador W, Sabatier MJ, Mulligan A. Electrical stimulation promotes peripheral axon regeneration by enhanced neuronal neurotrophin signaling. Dev Neurobiol. 2007;67(2):158e172. 7. Brushart TM, Jari R, Verge V, Rohde C, Gordon T. Electrical stimulation restores the specificity of sensory axon regeneration. Exp Neurol. 2005;194(1):221e229. 8. Al-Majed AA, Neumann CM, Brushart TM, Gordon T. Brief electrical stimulation promotes the speed and accuracy of motor axonal regeneration. J Neurosci. 2000;20(7):2602e2608. 9. Gordon 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(1):192e202. 10. Al-Majed AA, Tam SL, Gordon T. Electrical stimulation accelerates and enhances expression of regeneration-associated genes in regenerating rat femoral motoneurons. Cell Mol Neurobiol. 2004;24(3): 379e402. 11. Brushart TM, Hoffman PN, Royall RM, Murinson BB, Witzel C, Gordon T. Electrical stimulation promotes motoneuron regeneration without increasing its speed or conditioning the neuron. J Neurosci. 2002;22(15):6631e6638. 12. Gordon T, Brushart TM, Amirjani N, Chan KM. The potential of electrical stimulation to promote functional recovery after peripheral nerve injury—comparisons between rats and humans. Acta Neurochir Suppl. 2007;100:3e11. 13. Udina E, Furey M, Busch S, Silver J, Gordon T, Fouad K. Electrical stimulation of intact peripheral sensory axons in rats promotes outgrowth of their central projections. Exp Neurol. 2008;210(1): 238e247. 14. Macario A. What does one minute of operating room time cost? J Clin Anesth. 2010;22(4):233e236. 15. Wood M, Willits RK. Short-duration, DC electrical stimulation increases chick embryo DRG neurite outgrowth. Bioelectromagnetics. 2006;27(4):328e331. 16. Adams RD, Rendell SR, Counts LR, Papke JB, Willits RK, Harkins AB. Electrical and neurotrophin enhancement of neurite outgrowth within a 3d collagen scaffold. Ann Biomed Eng. 2014;42(6):1282e1291. 17. Varejao ASP, Melo-Pinto P, Meek MF, Filipe VA, Bulas-Cruz J. Methods for the experimental functional assessment of rat sciatic nerve regeneration. Neurol Res. 2004;26(2):186e194.
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18. Hare GMT, Evans PJ, Mackinnon SE, et al. Walking track analysis— a long-term assessment of peripheral-nerve recovery. Plast Reconstr Surg. 1992;89(2):251e258. 19. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis. Nature Methods. 2012;9(7):671e675. 20. Koka R, Hadlock TA. Quantification of functional recovery following rat sciatic nerve transection. Exp Neurol. 2001;168(1):192e195. 21. Lee AC, Yu VM, Lowe JB 3rd, et al. Controlled release of nerve growth factor enhances sciatic nerve regeneration. Exp Neurol. 2003;184(1):295e303. 22. Huang J, Lu L, Hu X, et al. Electrical stimulation accelerates motor functional recovery in the rat model of 15-mm sciatic nerve gap bridged by scaffolds with longitudinally oriented microchannels. Neurorehabil Neural Repair. 2010;24(8):736e745. 23. Lu M-C, Ho C-Y, Hsu S-F, et al. Effects of electrical stimulation at different frequencies on regeneration of transected peripheral nerve. Neurorehab Neural Re. 2008;22(4):367e373. 24. Cramp AFL, Gilsenan C, Lowe AS, Walsh DM. The effect of highand low-frequency transcutaneous electrical nerve stimulation upon cutaneous blood flow and skin temperature in healthy subjects. Clin Physiol. 2000;20(2):150e157. 25. Sluka KA, Walsh D. Transcutaneous electrical nerve stimulation: basic science mechanisms and clinical effectiveness. J Pain. 2003;4(3):109e121. 26. Lu MC, Tsai CC, Chen SC, Tsai FJ, Yao CH, Chen YS. Use of electrical stimulation at different current levels to promote recovery after peripheral nerve injury in rats. J Trauma. 2009;67(5):1066e1072. 27. Agnew WF, McCreery DB, Yuen TG, Bullara LA. Evolution and resolution of stimulation-induced axonal injury in peripheral nerve. Muscle Nerve. 1999;22(10):1393e1402. 28. McCreery DB, Agnew WF, Yuen TG, Bullara LA. Damage in peripheral nerve from continuous electrical stimulation: comparison of two stimulus waveforms. Med Biol Eng Comput. 1992;30(1): 109e114. 29. McCreery DB, Agnew WF, Yuen TG, Bullara LA. Relationship between stimulus amplitude, stimulus frequency and neural damage during electrical stimulation of sciatic nerve of cat. Med Biol Eng Comput. 1995;33(3 Spec No):426e429. 30. Alrashdan MS, Park JC, Sung MA, et al. Thirty minutes of low intensity electrical stimulation promotes nerve regeneration after sciatic nerve crush injury in a rat model. Acta Neurol Belg. 2010;110(2):168e179. 31. Huang JH, Zhang YG, Lu L, Hu XY, Luo ZJ. Electrical stimulation accelerates nerve regeneration and functional recovery in delayed
J Hand Surg Am.
r
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
peripheral nerve injury in rats. Eur J Neurosci. 2013;38(12): 3691e3701. Hare GM, Evans PJ, Mackinnon SE, et al. Walking track analysis: a long-term assessment of peripheral nerve recovery. Plast Reconstr Surg. 1992;89(2):251e258. Ahlborn P, Schachner M, Irintchev A. One hour electrical stimulation accelerates functional recovery after femoral nerve repair. Exp. Neurol. 2007;208(1):137e144. Yao CH, Chang RL, Chang SL, Tsai CC, Tsai FJ, Chen YS. Electrical stimulation improves peripheral nerve regeneration in streptozotocin-induced diabetic rats. J Trauma Acute Care Surg. 2012;72(1):199e205. Chen YS, Hsieh CL, Tsai CC, et al. Peripheral nerve regeneration using silicone rubber chambers filled with collagen, laminin and fibronectin. Biomaterials. 2000;21(15):1541e1547. Evans GRD, Brandt K, Katz S, et al. Bioactive poly(L-lactic acid) conduits seeded with Schwann cells for peripheral nerve regeneration. Biomaterials. 2002;23(3):841e848. Evans GR, Brandt K, Widmer MS, et al. In vivo evaluation of poly(L-lactic acid) porous conduits for peripheral nerve regeneration. Biomaterials. 1999;20(12):1109e1115. Evans GR, Brandt K, Niederbichler AD, et al. Clinical long-term in vivo evaluation of poly(L-lactic acid) porous conduits for peripheral nerve regeneration. J Biomater Sci Polym Ed. 2000;11(8): 869e878. Singh B, Xu QG, Franz CK, et al. Accelerated axon outgrowth, guidance, and target reinnervation across nerve transection gaps following a brief electrical stimulation paradigm. J. Neurosurg. 2012;116(3):498e512. Chamberlain LJ, Yannas IV, Hsu HP, Strichartz GR, Spector M. Near-terminus axonal structure and function following rat sciatic nerve regeneration through a collagen-GAG matrix in a tenmillimeter gap. J Neurosci Res. 2000;60(5):666e677. Kobayashi J, Mackinnon SE, Watanabe O, et al. The effect of duration of muscle denervation on functional recovery in the rat model. Muscle Nerve. 1997;20(7):858e866. Varejao AS, Meek MF, Ferreira AJ, Patricio JA, Cabrita AM. Functional evaluation of peripheral nerve regeneration in the rat: walking track analysis. J Neurosci Methods. 15 2001;108(1):1e9. van Meeteren NL, Brakkee JH, Hamers FP, Helders PJ, Gispen WH. Exercise training improves functional recovery and motor nerve conduction velocity after sciatic nerve crush lesion in the rat. Arch Phys Med Rehabil. 1997;78(1):70e77.
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Short-Term Electrical Stimulation to Promote Nerve Repair and Functional Recovery in a Rat Model Colleen Calvey, MD, Wenda Zhou, MS, Kimberly Sloan Stakleff, PhD, Patricia Sendelbach-Sloan, BS, Amy B. Harkins, PhD, William Lanzinger, MD, Rebecca Kuntz Willits, PhD
Purpose To evaluate the effect of duration of electrical stimulation on peripheral nerve regeneration and functional recovery. Based on previous work, we hypothesized that applying 10 minutes of electrical stimulation to a 10-mm rat sciatic nerve defect would significantly improve nerve regeneration and functional recovery compared with the non-electrical stimulation group. Methods A silicone tube filled with a collagen gel was used to bridge a 10-mm nerve defect in rats, and either 10 minutes or 60 minutes of electrical stimulation was applied to the nerve during surgery. Controls consisted of a silicone tube with collagen gel and no electrical stimulation or an isograft. We analyzed recovery over a 12-week period, measuring sciatic functional index and extensor postural thrust scores and concluding with histological examination of the nerve. Results Functional assessment scores at week 12 increased 24% in the 10-minute group as compared to the no stimulation control group. Electrical stimulation of either 10 or 60 minutes improved the number of nerve fibers over no stimulation. Additionally, the electrical stimulation group’s histomorphometric analysis was not different from the isograft group. Conclusions Several previous studies have demonstrated the effectiveness of 60-minute stimulations on peripheral nerve regeneration. This study demonstrated that an electrical stimulation of 10 minutes enhanced several functional and histomorphometric outcomes of nerve regeneration and was overall similar to a 60-minute stimulation over 12 weeks. Clinical relevance Decreasing the electrical stimulation time from 60 minutes to 10 minutes provided a potential clinically feasible and safe method to enhance nerve regeneration and functional recovery. (J Hand Surg Am. 2015;40(2):314e322. Copyright Ó 2015 by the American Society for Surgery of the Hand. All rights reserved.) Key words Peripheral nerve regeneration, electrical stimulation, rat sciatic nerve, direct current.
Received for publication July 18, 2014; accepted in revised form October 2, 2014.
Akron, Margaret F. Donovan Endowed Chair for Women in Engineering. The authors acknowledge Amanda Pinheiro for her assistance extracting the walking track video and Kelly Bonder, BS, Bethany Noble, BS, and Marcus Dempster for their technical support during surgeries. Histology was provided by Jason Papke, BS, and Dr. Jan Ryerse at Saint Louis University.
The present address of K.S.S. is Department of Urology, Akron General Medical Center, Akron, OH.
No benefits in any form have been received or will be received related directly or indirectly to the subject of this article.
The present address of P.S.-S. is Kent State University, Research Compliance, Kent, OH.
Corresponding author: Rebecca Kuntz Willits, PhD, Department of Biomedical Engineering, The University of Akron, 260 S. Forge St., OLRC 301, Akron, OH 44325-0302; e-mail: [email protected].
From the Department of Orthopaedic Surgery and the Kenneth Calhoun Research Laboratory, Akron General Medical Center, Akron; the Department of Biomedical Engineering, The University of Akron, Akron, OH; and the Department of Pharmacological and Physiological Science, Saint Louis University, St. Louis, MO.
The present address of C.C. is University of Louisville, Christine M. Kleinert Institute for Hand & Microsurgery, Louisville, KY. Funding for this study was provided by Akron General Medical Center, Akron General Development Foundation, and the Department of Orthopedics, as well as The University of
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0363-5023/15/4002-0018$36.00/0 http://dx.doi.org/10.1016/j.jhsa.2014.10.002
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frequent and costly occurrence, resulting from traumatic, compressive, and demyelinating disease mechanisms. Peripheral nerve injuries of the upper and lower extremities occur in about 3% of trauma patients,1 with an estimated incidence of digital nerve injuries in 6.2/100,000 patients.2 Spontaneous nerve regeneration occurs at an estimated rate of 1 mm/d, and it is often complicated by inhibitory growth factors, Wallerian degeneration, neuroma formation, and disorganized growth.3 Currently, treatments such as autografts, allografts, or nerve guidance conduits provide a scaffold to help guide regeneration, but do not affect the rate of recovery, and outcomes can be unpredictable. Safe and efficacious tools that can be coupled with currently available treatments to improve outcomes can have an important impact on nerve repair clinically. The nerve is analogous to an electrical cable, with an electrically active extracellular environment generated and maintained by ionic gradients and pumps. Externally applied electrical stimulation (ES), therefore, has been examined as a method to improve nerve regeneration. ES upregulates nerve growtheassociated genes, which have been found to correlate with acceleration of axonal regeneration.4,5 Additionally, a number of in vitro and in vivo studies have examined the ability of ES to enhance nerve repair and regeneration, maintain nerve plasticity, and improve functional recovery.6e9 The ES parameters that are required to provide improved outcomes are unknown, however. Applying ES of approximately 1 hour at the time of surgical repair can enhance nerve recovery.9e13 Given the high cost of operating-room time,14 however, an ES period of 60 minutes would be impractical and costprohibitive in a wide clinical setting. In previous in vitro studies, researchers found that environmental changes such as media composition, calcium ion levels, temperature, and stimulation duration (10 vs 100 min), had minimal impact on neurite outgrowth results. Any ES resulted in a 40% increase in neurite length compared with nonstimulated controls after 10 minutes of stimulation.15,16 Therefore, in this study, we examined whether 10-minute ES would also have a significant impact on nerve regeneration in vivo. ERIPHERAL NERVE INJURIES ARE A
METHODS Study design and ES The Akron General Medical Center institutional review board and institutional animal care and use committee approved all animal surgical and experimental procedures. Rats were divided into 4 groups: 10-minute ES (N ¼ 10), 60-minute ES (N ¼ 11), no ES J Hand Surg Am.
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control (N ¼ 10), and isograft control (N ¼ 10). In the first 3 groups, a 10-mm sciatic nerve gap was created and a silicone tube filled with 1.0 mg/mL collagen was used to bridge the defect. After entubulation, 0 (no stimulation group), 10 (10-min group), or 60 (60-min group) minutes of ES were applied as appropriate for the group. In the isograft group, the nerve defect was bridged by sciatic nerve retrieved from donor rats. In the stimulation groups, looped platinum wire electrodes were placed at the proximal and distal nerve segments after repair. A direct current of 24 V/m (24 mV, w1.5mA) was applied across the electrodes and run for 10 or 60 minutes. On the contralateral, non-experimental side, sham electrodes without electrical current were applied across a 10-mm segment of sciatic nerve. The exposed tissues were kept moist with sterile saline soaked gauze during the ES. The rats were then given food and water ad libitum and housed in pairs with a 12-hour lightedark cycle. Footprint recoding and sciatic functional index calculation Rats were made to traverse a walking track every 2 weeks where a digital camera captured their footprints.17 The rat feet were marked with black ink using a marker. Video was captured for at least 3 walking cycles, and when the rat took 5 continuous steps across the middle of the track and paws were not contracted, the image would be used for sciatic functional index (SFI) analysis. SFI was used to evaluate the sciatic nerve functional recovery every 2 weeks after surgery until the animals were killed. SFI was determined using the following equation as introduced by de Medinaceli and adapted by Bain,18 where: EPL NPL ETS NTS SFI ¼ 38:3 þ 109:5 NPL NTS EIT NIT þ 13:3 8:8 NIT
In this equation, EPL is distance from the heel to the top of the third toe in the injured foot, NPL is the same distance in the non-injured foot; ETS is distance between the first and the fifth toe in the injured foot, NTS is the same distance in the non-injured foot; EIT is distance from the second to the fourth toe in the injured foot, and NIT is the same distance in the noninjured foot. In SFI analysis, 4 extracted images of clear footprints were randomly chosen at each time point per rat, and ImageJ (NIH, Bethesda, MD)19 was used to measure the appropriate distances. An average of 4 measurements was then used to calculate the SFI value. These measurements were blinded by Vol. 40, February 2015
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FIGURE 1: Biweekly SFI for the different groups was collected as a measure of post injury functional recovery (N ¼ 7). * P < .05 for the measurement at week 12 relative to week 2. # P < .05 for the 10-minute electrical stimulation group versus the no stimulation control at week 12.
FIGURE 2: Biweekly EPT for different groups was collected as a measure of post injury functional recovery (N ¼ 10). * P < .05 between week 2 and week 12. # P < .05 between week 2 and week 10.
analyzing each video by date and then combined to the appropriate sample group. Extensor postural thrust measurement Extensor postural thrust (EPT) score was used as a separate evaluation of motor performance and measured on the same days as the SFI.17 EPT is classified as a postural reflex reaction and previous J Hand Surg Am.
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research has shown high correlation between SFI and EPT.20 The body of the rat was wrapped in a surgical towel with the hind limbs extending out. The animal was then placed over the platform of a digital balance. Once the hind limb made contact of the digital metatarsus, the hind limb was kept in contact with the platform for 30 seconds and the maximum force of the hind limb pushing against the platform was Vol. 40, February 2015
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FIGURE 3: The effect on the gastrocnemius muscle was examined by A box plot of weight for each sample, and histology images for B contralateral side, C no stimulation, D 10-minute electrical stimulation, E 60-minute electrical stimulation, and F isograft. The distance between the nuclei indicated the size of skeletal muscle cell. Cells in no stimulation control, 10-minute, and 60-minute groups were smaller than the isograft group, as the nucleus density was higher in those groups. Scale bar ¼ 50 mm.
recorded. The procedure was repeated 5 times and the 3 largest forces were used for analysis. Histomorphometric evaluation At the end of 12 weeks, the sciatic nerves were excised and segments were collected at the middle of the tube and 2 mm distal to the end of the silicone tube. The nerve segments were fixed in paraformaldehyde, epoxy-embedded, sectioned at 1 mm, and stained with toluidine blue. For each section, a 20x light microscope was used to capture the fascicular area. An entire nerve image was created and the perineurium edge was outlined using the software to measure total fascicular area. The number of nerve fibers and fiber area were then measured as described previously.21 Briefly, images were taken from 3 random regions per section. A specified area was processed in ImageJ to count the number of nerve fibers and measure the percentage of the area that was nerve fibers. The number of nerve fibers and percent fiber areas were J Hand Surg Am.
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calculated from a total of at least 15 images per group. Nerve fiber density was calculated by scaling the percentage of nerve fibers to the total intrafascicular area. The mean nerve fiber area was calculated by dividing nerve fiber number by total nerve fiber area. The mean fiber width was determined by relating the mean nerve fiber area to the area of a circle. In addition to the nerve, the gastrocnemius muscle was dissected from the injured side. The muscle samples were weighed and then dehydrated, embedded in paraffin, and stained with hematoxylin and eosin. A 40x light microscope was used to capture the muscle histology image. Statistics All data are expressed as mean standard deviation. Prior to institutional animal care and use committee approval, a power analysis was performed using histomorphometric data from the literature, with power ¼ 0.9 and significance level of .05, to set the Vol. 40, February 2015
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FIGURE 4: Sections of regenerated sciatic nerve at the middle of the conduit 12 weeks after surgery for the animals receiving A isograft, B no stimulation control, C 10-minute electrical stimulation, and D 60-minute electrical stimulation groups. Blood vessels (BV) were found on all groups that had a conduit. Scale bar ¼ 20 mm.
number per group at N ¼ 10.21 Comparisons within and between no stimulation, 10-minute, and 60minute groups were analyzed with one-way ANOVA with the Tukey post hoc test. Each group was then compared to the isograft with a t test. Values of P < .05 were considered statistically significant. RESULTS Functional evaluation SFI: No significant difference in SFI was found between any time points in the no stimulation group. After 12 weeks, SFI for 10-minute, 60-minute, and isograft groups were significantly higher than their respective SFI at the 2-week time point (P ¼ .002, P ¼ .02, P ¼ .01, respectively). When comparing within each group in the same time point, no significant differences were found in the early weeks (Fig. 1). SFI in the 10-minute group was significantly improved by 24% compared with the no stimulation group at week 12 (P ¼ .03). At the 12-week time point, there was no significant difference between the 10-minute and 60-minute groups (P ¼0.2) or between the ES groups and the isograft group. EPT: Similar to SFI analysis, no significant difference was found in EPT between time points for the no stimulation control (Fig. 2). EPT in the 10-minute group gradually increased with increasing recovery time. At 12 weeks, rats from the 10-minute group had J Hand Surg Am.
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a significantly higher EPT force than at the earlier time points of 2, 4, 6, and 8 weeks (P < .04). No significant difference was found between the 10- and 60-minute groups at each time point. Histomorphometric evaluation Muscle weight and histology: The extracted gastrocnemius muscle weights (in g) on the experimental side were not significantly different between the no ES control, 10-minute, and 60-minute groups. The 10-minute and 60-minute groups were significantly lower than the isograft group (P ¼ .035, P ¼ .001 respectively, Fig. 3). From the histology of gastrocnemius muscles, cell atrophy was visible in the 10-minute and 60minute groups compared with the isograft group and normal sciatic nerve. Number of nerve fibers: The overall histology of the nerves is shown in Figure 4. There was no significant difference in nerve samples from the midline of injury between the no stimulation control, 10-minute, and 60-minute groups, although they were all significantly lower than the isograft group (P < .003; Fig. 5). In the distal nerve sample, the average number of nerve fibers from the 10-minute (P ¼ .03) and 60-minute (P ¼ .026) groups was significantly higher than the no stimulation control (Fig. 6). No significant difference was found between either ES groups or the isograft group in number of distal Vol. 40, February 2015
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FIGURE 5: Histomorphometric data collected at the midline of the conduit (N 6). The data were quantified from the images for A number of nerves, B percent nerve area, C mean fiber width, and D fiber density. * P < .05 versus no stimulation control group.
nerves, but the no stimulation control group was significantly lower than the isograft group (P ¼ .013). Area of nerve fiber: The quality of nerve regeneration is represented by the percentage area of nerve fiber and was significantly higher in the 60-minute group at both the midline (Fig. 5) and distal (Fig. 6) nerve samples than the no stimulation control (P ¼ .04 and P ¼ .038, respectively). The 10-minute group did not show a significant difference compared with the no stimulation control. No significant differences existed between the 10- or 60-minute groups and the isograft group at either the midline or the distal nerve sample, but the no stimulation controls were significantly lower than isograft groups at both the midline (P ¼ .005) and the distal (P ¼ .003) nerve samples. Mean fiber width: Mean fiber width (Figs. 5, 6), an indirect measure of myelination and potentially mature nerve fibers, was not statistically different between the ES groups, although they were significantly higher than the no stimulation control at the midline nerve samples (P ¼ .006 and P ¼ .002, respectively). When compared with the isograft group, the no stimulation control was significant lower than the isograft group (P ¼ .016) at the midline, and 10minute and 60-minute groups were not statistically different from isograft group. Density: Although there was no significant difference between the 10-minute and 60-minute groups and no J Hand Surg Am.
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stimulation control at the midline, the density of the 60-minute group was significantly higher than the no stimulation control at the distal end (P ¼ .015) (Fig. 6). The 10-minute and 60-minute groups were not statistically different from the isograft group (Figs. 5, 6), whereas the no stimulation control was significantly lower than the isograft (P ¼ .005) at the distal end. DISCUSSION The application of ES to nerves provides an additional tool to promote nerve regeneration and functional recovery after nerve injury. Researchers have examined several ES parameters for one hour’s duration since 2000, when Gordon and colleagues demonstrated that application of ES for 1 hour was as effective as weeks of ES to promote nerve regeneration.8 For example, 1 hour of ES using pulsed direct current (square 0.1-ms pulses [4V] at 20 Hz) accelerated motor functional recovery in a larger rat nerve defect (15 mm).22 Additionally, changing the frequency of an alternating current stimulation (1 mA) impacted nerve regeneration.23 We extend these previous reports by demonstrating that application of direct current, rather than alternating current or pulsed direct current, similarly improved sciatic outcome parameters in rats after ES of 60 or 10 minutes. The low current applied here, which is 100 to 1,000 times lower than applied for TENS24,25 and Vol. 40, February 2015
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FIGURE 6: Histomorphometric data collected at 2 mm distal to the repair conduit (N 5). The data were quantified from the images for A number of nerves, B percent nerve area, C mean fiber width, and D fiber density. * P < .05 versus the no stimulation control.
lower than previous studies,26 reduces the time and improves the safety of application while not reducing the efficacy. Although low degrees of late-onset injury or axonal degeneration have been associated with ES (up to 8 hours, 50 Hz, and 2100e4500 mA),27e29 longer durations of ES would increase surgical risks due to increased time under anesthesia and risk of infection. In addition, increased time in the operating room increases the costs associated with implementing a new procedure. Improving safety and reducing cost while maintaining efficacy are important benefits of decreasing the duration of ES. In an effort to mitigate safety concerns with the application of 60 minutes of ES, researchers have investigated the effect of reducing the duration of ES. A 30-minute stimulation (20-Hz pulse rate, 2mA amplitude) promoted nerve regeneration after a nerve crush injury in a rat model.30 After 3 weeks, SFI increased by 28% in 30-minute stimulation groups compared with controls.30 Although this result was promising, the outcomes of interventions after crush injury do not always match the outcome after transection injury. In a 5-mm transection, a 20-minute stimulation (3V, 20 Hz) accelerated nerve regeneration in delayed nerve injury repair rat model.31 Using only 10 minutes of stimulation, we saw improved SFI and EPT functional recovery measures, demonstrating efficacy. With the equation used to calculate J Hand Surg Am.
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SFI, a value close to zero indicates that the rat’s toe spread is equivalent on both injured and non-injured side, and a value close to e100 indicates total impairment. After 1 year, functional recovery after sciatic nerve repair with optimal transection conditions is limited to 41% of function, or a score of e41.32 After 12 weeks, our SFI scores approached 50%. Compared with the no stimulation control, the 10-minute stimulation increased the SFI by 24% after 12 weeks, and SFI in the 60-minute group was increased by 13%. This improvement in functional recovery over shorter recovery time frames was comparable to the crush injury results,30 as well as results of studies after 60 minutes of stimulation.33 Although the effectiveness of short-duration stimulation is controversial,3 both our functional recovery and histomorphometric results showed that 10 minutes of stimulation was similar to 60 minutes of stimulation. The histological sections provided several evaluations of nerve recovery. The blood vessels found in the midline of conduits would provide nutrition to regrowing nerves (Fig. 4). The mean fiber width, which represents the maturity of regenerating nerve fibers, indicated that the 10-minute stimulation was as good as a stimulation of 60 minutes. The density of nerve fibers in the 60-minute group was also greater than no stimulation control, which is consistent with previous literature.34 No Vol. 40, February 2015
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significant differences were found between the stimulation groups and the isograft group in number of distal nerves, area of nerve percent, and fiber density at distal end, whereas the no stimulation group was significantly lower than isograft group on those parameters. This result suggests that the distal nerves are better sustained with ES than without, making the group similar to an isograft. Although the isograft group had an increased number of nerves at the midline, this result would be expected as the isograft itself contains nerve fibers. At the midline, the area of nerve fibers for the 60-minute group and the mean fiber width of both stimulation groups were significantly higher than the no stimulation control, suggesting that ES could enhance the quality and maturity of nerve regeneration. Overall, we did not find any significant differences between the 10- and 60-minute groups in the histomorphometric parameters studied, suggesting that a 10-minute stimulation may suitably replace a 60-minute stimulation. There were several limitations to the current study. A collagen gel was placed into the conduit prior to implantation, and collagen offers a suitable scaffold for the regeneration of axons.35 Therefore, synergies between the mechanism of ES and growth within a collagen tube should be further explored. The weight of the gastrocnemius muscle indirectly evaluates nerve regeneration.36 In this study, we found the muscle weight in the isograft group was higher than the other groups. Although this result was similar to other studies,37,38 it would be important to determine the cause of these weight differences. Previous studies have shown that ES supports motor neuron recovery and survival.8,11,39 The sciatic nerve is a mixed sensory and motor nerve, yet functional recovery, as measured here, was dependent on motor neuron survival and regeneration. Future experiments should be directed at examining sensory nerve recovery. Toe contractures were found in all groups, reducing the number of animals analyzed from 10 to 7 in SFI. Although SFI reliability may be compromised by removal of animals, it was not the case in this study as shown by significance found at week 12 with a 10minute stimulation. Because contractures indicate a lack of physical therapy rather than a lack of function,40,41 SFI is still a useful noninvasive tool to measure functional recovery.42 Daily exercise training should be used to increase the functional recovery and SFI numbers.43 In this study, histomorphometric samples were lost due to damage of some embedding process, thereby reducing the numbers as noted in the results. Post hoc power analysis was used to confirm any significant differences. J Hand Surg Am.
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Despite these limitations, our findings suggest that 10 minutes of ES enhanced nerve regeneration and functional recovery compared with no ES. Decreasing the duration of stimulation has made this method a safe, efficient, and feasible tool for future clinical use. REFERENCES 1. Kang JR, Zamorano DP, Gupta R. Limb salvage with major nerve injury: current management and future directions. J Am Acad Orthop Surg. 2011;19(Suppl 1):S28eS34. 2. Thorsen F, Rosberg HE, Steen Carlsson K, Dahlin LB. Digital nerve injuries: epidemiology, results, costs, and impact on daily life. J Plast Surg Hand Surg. Sep 2012;46(3e4):184e190. 3. Gordon T, Sulaiman O, Boyd JG. Experimental strategies to promote functional recovery after peripheral nerve injuries. J Peripher Nerv Syst. 2003;8(4):236e250. 4. Al-Majed AA, Brushart TM, Gordon T. Electrical stimulation accelerates and increases expression of BDNF and trkB mRNA in regenerating rat femoral motoneurons. EurJ Neurosci. 2000;12(12):4381e4390. 5. Geremia NM, Gordon T, Brushart TM, Al-Majed AA, Verge VM. Electrical stimulation promotes sensory neuron regeneration and growth-associated gene expression. Exp Neurol. 2007;205(2): 347e359. 6. English AW, Schwartz G, Meador W, Sabatier MJ, Mulligan A. Electrical stimulation promotes peripheral axon regeneration by enhanced neuronal neurotrophin signaling. Dev Neurobiol. 2007;67(2):158e172. 7. Brushart TM, Jari R, Verge V, Rohde C, Gordon T. Electrical stimulation restores the specificity of sensory axon regeneration. Exp Neurol. 2005;194(1):221e229. 8. Al-Majed AA, Neumann CM, Brushart TM, Gordon T. Brief electrical stimulation promotes the speed and accuracy of motor axonal regeneration. J Neurosci. 2000;20(7):2602e2608. 9. Gordon 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(1):192e202. 10. Al-Majed AA, Tam SL, Gordon T. Electrical stimulation accelerates and enhances expression of regeneration-associated genes in regenerating rat femoral motoneurons. Cell Mol Neurobiol. 2004;24(3): 379e402. 11. Brushart TM, Hoffman PN, Royall RM, Murinson BB, Witzel C, Gordon T. Electrical stimulation promotes motoneuron regeneration without increasing its speed or conditioning the neuron. J Neurosci. 2002;22(15):6631e6638. 12. Gordon T, Brushart TM, Amirjani N, Chan KM. The potential of electrical stimulation to promote functional recovery after peripheral nerve injury—comparisons between rats and humans. Acta Neurochir Suppl. 2007;100:3e11. 13. Udina E, Furey M, Busch S, Silver J, Gordon T, Fouad K. Electrical stimulation of intact peripheral sensory axons in rats promotes outgrowth of their central projections. Exp Neurol. 2008;210(1): 238e247. 14. Macario A. What does one minute of operating room time cost? J Clin Anesth. 2010;22(4):233e236. 15. Wood M, Willits RK. Short-duration, DC electrical stimulation increases chick embryo DRG neurite outgrowth. Bioelectromagnetics. 2006;27(4):328e331. 16. Adams RD, Rendell SR, Counts LR, Papke JB, Willits RK, Harkins AB. Electrical and neurotrophin enhancement of neurite outgrowth within a 3d collagen scaffold. Ann Biomed Eng. 2014;42(6):1282e1291. 17. Varejao ASP, Melo-Pinto P, Meek MF, Filipe VA, Bulas-Cruz J. Methods for the experimental functional assessment of rat sciatic nerve regeneration. Neurol Res. 2004;26(2):186e194.
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18. Hare GMT, Evans PJ, Mackinnon SE, et al. Walking track analysis— a long-term assessment of peripheral-nerve recovery. Plast Reconstr Surg. 1992;89(2):251e258. 19. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis. Nature Methods. 2012;9(7):671e675. 20. Koka R, Hadlock TA. Quantification of functional recovery following rat sciatic nerve transection. Exp Neurol. 2001;168(1):192e195. 21. Lee AC, Yu VM, Lowe JB 3rd, et al. Controlled release of nerve growth factor enhances sciatic nerve regeneration. Exp Neurol. 2003;184(1):295e303. 22. Huang J, Lu L, Hu X, et al. Electrical stimulation accelerates motor functional recovery in the rat model of 15-mm sciatic nerve gap bridged by scaffolds with longitudinally oriented microchannels. Neurorehabil Neural Repair. 2010;24(8):736e745. 23. Lu M-C, Ho C-Y, Hsu S-F, et al. Effects of electrical stimulation at different frequencies on regeneration of transected peripheral nerve. Neurorehab Neural Re. 2008;22(4):367e373. 24. Cramp AFL, Gilsenan C, Lowe AS, Walsh DM. The effect of highand low-frequency transcutaneous electrical nerve stimulation upon cutaneous blood flow and skin temperature in healthy subjects. Clin Physiol. 2000;20(2):150e157. 25. Sluka KA, Walsh D. Transcutaneous electrical nerve stimulation: basic science mechanisms and clinical effectiveness. J Pain. 2003;4(3):109e121. 26. Lu MC, Tsai CC, Chen SC, Tsai FJ, Yao CH, Chen YS. Use of electrical stimulation at different current levels to promote recovery after peripheral nerve injury in rats. J Trauma. 2009;67(5):1066e1072. 27. Agnew WF, McCreery DB, Yuen TG, Bullara LA. Evolution and resolution of stimulation-induced axonal injury in peripheral nerve. Muscle Nerve. 1999;22(10):1393e1402. 28. McCreery DB, Agnew WF, Yuen TG, Bullara LA. Damage in peripheral nerve from continuous electrical stimulation: comparison of two stimulus waveforms. Med Biol Eng Comput. 1992;30(1): 109e114. 29. McCreery DB, Agnew WF, Yuen TG, Bullara LA. Relationship between stimulus amplitude, stimulus frequency and neural damage during electrical stimulation of sciatic nerve of cat. Med Biol Eng Comput. 1995;33(3 Spec No):426e429. 30. Alrashdan MS, Park JC, Sung MA, et al. Thirty minutes of low intensity electrical stimulation promotes nerve regeneration after sciatic nerve crush injury in a rat model. Acta Neurol Belg. 2010;110(2):168e179. 31. Huang JH, Zhang YG, Lu L, Hu XY, Luo ZJ. Electrical stimulation accelerates nerve regeneration and functional recovery in delayed
J Hand Surg Am.
r
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
peripheral nerve injury in rats. Eur J Neurosci. 2013;38(12): 3691e3701. Hare GM, Evans PJ, Mackinnon SE, et al. Walking track analysis: a long-term assessment of peripheral nerve recovery. Plast Reconstr Surg. 1992;89(2):251e258. Ahlborn P, Schachner M, Irintchev A. One hour electrical stimulation accelerates functional recovery after femoral nerve repair. Exp. Neurol. 2007;208(1):137e144. Yao CH, Chang RL, Chang SL, Tsai CC, Tsai FJ, Chen YS. Electrical stimulation improves peripheral nerve regeneration in streptozotocin-induced diabetic rats. J Trauma Acute Care Surg. 2012;72(1):199e205. Chen YS, Hsieh CL, Tsai CC, et al. Peripheral nerve regeneration using silicone rubber chambers filled with collagen, laminin and fibronectin. Biomaterials. 2000;21(15):1541e1547. Evans GRD, Brandt K, Katz S, et al. Bioactive poly(L-lactic acid) conduits seeded with Schwann cells for peripheral nerve regeneration. Biomaterials. 2002;23(3):841e848. Evans GR, Brandt K, Widmer MS, et al. In vivo evaluation of poly(L-lactic acid) porous conduits for peripheral nerve regeneration. Biomaterials. 1999;20(12):1109e1115. Evans GR, Brandt K, Niederbichler AD, et al. Clinical long-term in vivo evaluation of poly(L-lactic acid) porous conduits for peripheral nerve regeneration. J Biomater Sci Polym Ed. 2000;11(8): 869e878. Singh B, Xu QG, Franz CK, et al. Accelerated axon outgrowth, guidance, and target reinnervation across nerve transection gaps following a brief electrical stimulation paradigm. J. Neurosurg. 2012;116(3):498e512. Chamberlain LJ, Yannas IV, Hsu HP, Strichartz GR, Spector M. Near-terminus axonal structure and function following rat sciatic nerve regeneration through a collagen-GAG matrix in a tenmillimeter gap. J Neurosci Res. 2000;60(5):666e677. Kobayashi J, Mackinnon SE, Watanabe O, et al. The effect of duration of muscle denervation on functional recovery in the rat model. Muscle Nerve. 1997;20(7):858e866. Varejao AS, Meek MF, Ferreira AJ, Patricio JA, Cabrita AM. Functional evaluation of peripheral nerve regeneration in the rat: walking track analysis. J Neurosci Methods. 15 2001;108(1):1e9. van Meeteren NL, Brakkee JH, Hamers FP, Helders PJ, Gispen WH. Exercise training improves functional recovery and motor nerve conduction velocity after sciatic nerve crush lesion in the rat. Arch Phys Med Rehabil. 1997;78(1):70e77.
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