|Year : 2020 | Volume
| Issue : 4 | Page : 335-341
Low-intensity pulsed ultrasound for regenerating peripheral nerves: potential for penile nerve
Dong-Yi Peng1,2, Amanda B Reed-Maldonado1, Gui-Ting Lin1, Shu-Jie Xia3, Tom F Lue1
1 Knuppe Molecular Urology Laboratory, Department of Urology, School of Medicine, University of California, San Francisco, CA 94143, USA
2 Department of Urology, The Third Xiangya Hospital of Central South University, Changsha 410013, China
3 Department of Urology, Shanghai General Hospital, Shanghai Jiao Tong University, Shanghai 200240, China
|Date of Submission||22-Mar-2019|
|Date of Acceptance||11-Jul-2019|
|Date of Web Publication||13-Sep-2019|
Tom F Lue
Knuppe Molecular Urology Laboratory, Department of Urology, School of Medicine, University of California, San Francisco, CA 94143
Source of Support: None, Conflict of Interest: None
Peripheral nerve damage, such as that found after surgery or trauma, is a substantial clinical challenge. Much research continues in attempts to improve outcomes after peripheral nerve damage and to promote nerve repair after injury. In recent years, low-intensity pulsed ultrasound (LIPUS) has been studied as a potential method of stimulating peripheral nerve regeneration. In this review, the physiology of peripheral nerve regeneration is reviewed, and the experiments employing LIPUS to improve peripheral nerve regeneration are discussed. Application of LIPUS following nerve surgery may promote nerve regeneration and improve functional outcomes through a variety of proposed mechanisms. These include an increase of neurotrophic factors, Schwann cell (SC) activation, cellular signaling activations, and induction of mitosis. We searched PubMed for articles related to these topics in both in vitro and in vivo animal research models. We found numerous studies, suggesting that LIPUS following nerve surgery promotes nerve regeneration and improves functional outcomes. Based on these findings, LIPUS could be a novel and valuable treatment for nerve injury-induced erectile dysfunction.
Keywords: activation; cellular signaling; low-intensity pulsed ultrasound; neurotrophic factors; peripheral nerve regeneration; Schwann cells
|How to cite this article:|
Peng DY, Reed-Maldonado AB, Lin GT, Xia SJ, Lue TF. Low-intensity pulsed ultrasound for regenerating peripheral nerves: potential for penile nerve. Asian J Androl 2020;22:335-41
|How to cite this URL:|
Peng DY, Reed-Maldonado AB, Lin GT, Xia SJ, Lue TF. Low-intensity pulsed ultrasound for regenerating peripheral nerves: potential for penile nerve. Asian J Androl [serial online] 2020 [cited 2020 Jul 11];22:335-41. Available from: http://www.ajandrology.com/text.asp?2020/22/4/335/266830 - DOI: 10.4103/aja.aja_95_19
| Introduction|| |
Peripheral nerves are often damaged by compression, stretch, avulsion, or division. For example, radical prostatectomy, the gold standard for surgical treatment of prostate cancer, may damage the cavernous nerves, causing neurogenic erectile dysfunction. The prevalence of erectile dysfunction secondary to nerve damage during radical prostatectomy is estimated to be 14%–90%. In a recent study, Jo et al. showed that early penile rehabilitation with sildenafil after robot-assisted laparoscopic prostatectomy significantly improved erectile dysfunction compared with delayed treatment. However, the treatment options for nerve injury after radical prostatectomy remain limited, and the prognosis remains poor if treatment is delayed. Fortunately, previous studies have shown that low-intensity pulsed ultrasound (LIPUS) has the potential to induce nerve regeneration by stimulating neurotrophic factors and reducing neuroinflammation.,
Peripheral nerve damage is a significant clinical challenge, which leads to long-lasting morbidity, disability, and economic costs.,, When identified, peripheral nerve injuries are typically reconstructed by primary repair (direct reconnection between damaged nerve stumps), by interposition of an artificial conduit, or by autologous nerve graft if tension-free coaptation is not possible. In cases of severe nerve injury, the long distance between the lesion and the end organ may represent a limiting factor for reinnervation.,,, One approach to accelerate peripheral nerve regeneration is to stimulate the physiological processes that occur following nerve injury thereby promoting nerve regeneration.
There are many proposed methods of speeding nerve regeneration, including physical methods (such as electric stimulation, magnetic field stimulation, and laser stimulation ) and biological methods (such as administration of neurotrophic factors, vitamins, and medications ). However, there are some disadvantages to the clinical application of many of these therapies, and their clinical efficacy is unproven in many cases. Therefore, a novel and effective therapeutic approach to stimulate the physiological processes involved in nerve regeneration is needed.
Very recently, LIPUS has been successfully employed to promote tissue healing;, to inhibit inflammation and reduce pain; to provoke differentiation of stem cells; and to stimulate tissue regeneration of muscle, nerve, bone, ligament, and articular cartilage in intervertebral discs.,,,, It is believed that ultrasound waves stimulate tissue regeneration by transmitting mechanical energy which induces mechanical motion of molecules in periodically alternating phases of compression and rarefaction. Although no clinical studies examining the effects of LIPUS on nerve regeneration exist, several experimental studies have investigated the application of LIPUS treatment following peripheral motor nerve injury and report positive outcomes. This article presents a systematic review of the available preclinical literature reporting on the effects of LIPUS in peripheral nerve regeneration and discusses the potential clinical applications of LIPUS.
| Pathogenesis of Nerve Injury and Regeneration|| |
Peripheral nerves are particularly vulnerable to injury, but the nerves of the peripheral nervous system have the ability to regenerate. This is in contrast to the nerves of the central nervous system, which cannot regenerate. Currently, the pathophysiology of peripheral nerve injuries and the mechanisms involved in spontaneous regeneration are relatively well understood. There is some evidence that a conditioning lesion primes the peripheral nerve for regeneration. Despite this structural recovery, functional recovery is often incomplete.
The process of spontaneous nerve regeneration starts with the initial response to an injury such as a complete nerve transection. After nerve transection, the distal nerve ending undergoes Wallerian degeneration, which is a unique and structured form of axon degeneration. At first, axonal and myelin debris are produced, and resident macrophages in the nerve tissue then differentiate into activated macrophages to phagocytose the cellular debris. In the proximal stumps of axons, activation of mRNA translation is observed. This stimulates the formation of the protein complex, importin-phosphorylated extracellular regulated protein kinase 1/2 vimentin. This complex is transported by the motor protein dynein in a retrograde direction to the cell body, and this signal informs the neuron of the axonal damage. The neuron of soma then reacts by increasing its volume and breaking up Nissl bodies to promote protein synthesis., Within a few hours of the nerve injury, the growing axonal extremity extends filopodia which are randomly oriented at first but thereafter gain unidirectionality. Next, the proximal stump sprouts processes that sample the environment for neurotrophic factors to guide them to their target.,,
Successful peripheral nerve regeneration after injury relies on both injured axons and nonneuronal cells, including Schwann cells (SCs), endoneurial fibroblasts, and macrophages, which produce a supportive microenvironment to promote successful regrowth of the proximal nerve endings. SCs play an important role in the axonal regeneration. They secret chemokines, such as monocyte chemoattractant protein-1, which recruit circulating macrophages to remove myelin and axonal debris., SCs also produce neurite-promoting proteins, such as fibronectin, laminin, tenascin, heparin sulfate, and collagen, which are incorporated to replace the extracellular matrix (ECM) lost secondary to injury. In addition, proliferating SCs align into columns and form “bands of Büngner,” which provides a physical guide for new axonal regrowth., To further support neuronal regeneration, SCs express cell adhesion molecules that interact with matrix proteins to modulate axonal outgrowth and pathfinding.,, SCs also express neurotrophic factors, such as ciliary neurotrophic factor (CNTF), brain-derived neurotrophic factor (BDNF), glial cell line-derived neurotrophic factor (GDNF), and nerve growth factor (NGF), which increase cell survival and promote nerve regeneration. Furthermore, it was very recently reported that SCs regulate peripheral nerve regeneration by secreting exosomes.
| The Physical Characteristics of Ultrasound|| |
Ultrasound is defined as sound waves with frequencies above the human hearing threshold. Ultrasound is clinically divided into two main categories: imaging ultrasound and therapeutic ultrasound. Both preclinical and clinical studies have shown that LIPUS stimulates tissue regeneration by transmitting mechanical energy. The beneficial biological effects of LIPUS likely result from the biomechanical conduction of the ultrasound vibration, which produces microturbulence within the intercellular and intracellular fluids in the vicinity of the wave. Recent developments in the science of ultrasound have improved and refined the technology, making ultrasound a promising therapy for various diseases.,,,
An ultrasound wave is a high-frequency wave that is generally 1–12 MHz. Depending on the level of ultrasonic energy, therapeutic ultrasound can be classified into two categories: high-intensity ultrasound with peak intensities of 5000–15 000 W cm −2 and low-intensity ultrasound with peak intensities of 0.5–3000 mW cm −2. LIPUS is delivered at low intensities (<0.1 W cm −2) and at a constant frequency (1–1.5 MHz)., LIPUS is both nonthermogenic and nondestructive to tissues. This is in direct contrast to high-intensity continuous ultrasound.
LIPUS has been found to have a wide range of biological effects on tissues, including promoting bone fracture healing, accelerating soft-tissue regeneration,, and inhibiting inflammatory responses. However, the potential mechanisms producing the above biological effects are still unclear and are under continuing investigation. Low-intensity extracorporeal shock wave therapy (Li-ESWT) is similar to LIPUS, but features a single, mainly positive pressure wave with high amplitude, short duration, and fast rise time.
| Effect of Lipus on Peripheral Nerve Regeneration|| |
LIPUS promotes peripheral nerve regeneration
It has been reported that peripheral nerves are very sensitive to ultrasound stimulation and that ultrasound can reversibly regulate nerve conduction. It has also been found that LIPUS can promote functional recovery of oppressive neuropathy, which suggests that LIPUS simulates damaged nerves to regenerate., Further, a study found that LIPUS stimulates the growth of SCs, thereby accelerating the recovery of damaged nerves.
Autologous nerve grafts, widely used to bridge peripheral nerve defects, serve as a standard repair technique when primary suture anastomosis is impossible. However, the limited availability of donor nerves and donor-site morbidity are major limitations of this technique. In addition, the outcomes of autologous nerve transplantation are far from ideal. With this in mind, researchers have investigated whether LIPUS can improve the outcomes of autologous nerve transplantation. In addition, significant effort has been made to generate synthetic nerve conduits,, which may promote axonal proliferation by developing a scaffold, recruiting support cells (i.e., SCs and macrophages), and producing induction factors and extracellular matrices. Multiple groups have investigated the effect of LIPUS on sciatic nerve regeneration after interposition of autologous nerve or synthetic nerve conduits. In 2016, Jiang et al. used a rat sciatic nerve defect model with a right-sided 10 mm sciatic nerve reversed autologous nerve transplantation and treated with LIPUS (1 MHz, 0.25 W cm −2 for 5 min). Functional results showed that sciatic functional index (SFI) and electrophysiological evaluation were significantly increased with LIPUS. Histologic results showed that LIPUS increased the rate of axonal regeneration significantly. These results suggested that autograft nerve regeneration was improved. The authors hypothesized that LIPUS provides appropriate mechanical stimulus to promote local neovascularization, to stimulate nerve sprouting, and to provoke the release of more neurotrophic factors. In 2004, Chang and Hsu  found that LIPUS can improve peripheral nerve regeneration on poly(DL-lactic acid-co-glycolic acid) (PLGA) nerve guidance conduits seeded with SCs. These authors interposed the seeded conduit into rats' sciatic nerve gaps, then they treated the site with LIPUS (1 MHz, 0.2 W cm −2 for 5 min). The results showed that LIPUS stimulated nerve regrowth, and the LIPUS-treated rats exhibited considerably more myelinated axons with a larger mean area at the mid-conduit than the control group. These results suggest that LIPUS may stimulate the SCs within the PLGA conduits to regenerate nerves. In 2010, Park et al. used a rat sciatic nerve defect model to explore the effect of LIPUS as a simple, noninvasive stimulus at the poly(lactic-co-glycolic acid) and Pluronic F127 (PLGA/F127) nerve guide site. The results showed that animals treated with LIPUS (1 MHz, 0.4 W cm −2 for 2 min) displayed more rapid nerve regeneration (0.71 mm per day) than the group without LIPUS treatment (0.48 mm per day). The LIPUS group also showed greater neural tissue area as well as larger axon diameter and thicker myelin sheaths than the group without LIPUS treatment, indicating improved nerve regeneration. These effects of LIPUS may be due to both the physical stimulation of SCs and the activation of the neurotrophic factors.
A recent meta-analysis  reviewed ten preclinical in vivo LIPUS studies which included a total of 445 animals. The authors included four studies with sciatic nerve crush injury, one study with reverse sciatic autograft, and five studies with a conduit. The results showed that repetitive LIPUS with intensities between 200 mW cm −2 and 500 mW cm −2 significantly promoted axonal regeneration and muscle reinnervation, increased the number and myelination of axons distal to the lesion site, and improved nerve conduction velocity after nerve injury. In addition, there were no negative side effects noted. Overall, there is significant experimental evidence that LIPUS promotes both functional and structural peripheral nerve regeneration after nerve injury.
Dosage of LIPUS for peripheral nerve regeneration
In 1988, Lowdon et al. investigated the effects of therapeutic ultrasound for regeneration of the tibial nerve following a compressive lesion in a rat model. These authors demonstrated that the nerve conduction velocity recovered significantly earlier with the lower intensity of 0.5 W cm −2 and significantly later with the higher intensity of 1 W cm −2, as compared to the control group. They concluded that low-intensity therapeutic ultrasound promoted nerve regeneration, but high-intensity ultrasound delayed nerve regeneration. A similar study in 2001 used a rat sciatic nerve crush injury model followed with therapeutic ultrasound of different intensities and frequencies. These authors applied LIPUS three times a week for 1 month, and they found that nerve regeneration was enhanced with an intensity of 0.25 W cm −2 and a frequency of 2.25 MHz. Over the following decade, more and more researchers utilized the sciatic nerve injury rat model to explore the effects of LIPUS on peripheral nerve regeneration. In 2002, Crisci and Ferreira  found that LIPUS (16 mW cm −2, 1.5 MHz) stimulated faster regeneration of peripheral nerves following neurotomy. These authors suggested that the numerous thick fibers in the nerves of LIPUS-treated animals were a result of amplified SC activity, which led to earlier recovery of their myelin sheaths. In 2005, Raso et al. found that LIPUS (1 MHz, 0.4 W cm −2, 2 min duration) increased SFIs and prompted nerve regeneration after sciatic nerve crush injury. Three weeks after nerve crush injury, the SFI improved more significantly for the LIPUS-treated nerves (73%) than the control (55%). The small-diameter, thin myelin sheath fibers typical of nerve regeneration were predominant in the LIPUS-treated group, as opposed to large-diameter, thin myelin sheath fibers in the control group. This suggested that LIPUS enhanced nerve regeneration.
In 2010, Chen et al. utilized LIPUS (0.25 W cm −2, 1.0 MHz for 1 min) to treat a sciatic nerve crush rat model. Their results showed that the density of nerve fibers with myelin sheaths and the SFI of the treatment group were significantly higher than those of the control group. This suggested that LIPUS accelerated the regeneration and functional recovery of injured sciatic nerves. In 2017, another group used similar LIPUS (0.2 W cm −2, 1.0 MHz for 1 min) to treat the sciatic nerve crush injury rat model. These authors found that the LIPUS-treated rats had higher SFIs, compound muscle action potentials, wet weight ratios of the target muscle, and mRNA expression of BDNF in the crushed nerve and ipsilateral dorsal root ganglia as compared with the control group. This suggested that LIPUS might promote injured nerve regeneration by stimulating BDNF release. Overall, in these studies, the effective dosage of LIPUS for nerve regeneration ranged from 0.016 W cm −2 to 1 W cm −2.
LIPUS promotes recovery of erectile function
In 2015, Lei et al. used an erectile dysfunction (ED) rat model with streptozotocin-induced diabetes mellitus (DM) to explore the effect of LIPUS on erectile function. After 2 weeks of LIPUS treatment with different low energy levels (100, 200, and 300 mW cm −2; 3 times per week), intracavernous pressure (ICP) was measured, and neuronal nitric oxide synthase (nNOS) expression in penile tissue was examined by histology and Western blot. The results showed that LIPUS enhanced the ICP levels and increased the nNOS expression in both dorsal and cavernous nerves. These results indicated that LIPUS can significantly improve erectile function in diabetic rats. In 2019, Chiang and Yang  hypothesized that LIPUS could have a therapeutic effect on erectile function deriving from cavernous nerve injury based on its neuroregenerative and protective effects. Thus far, little research has been done to further investigate this hypothesis or the potential efficacy of LIPUS for the treatment of ED.
| Lipus Activates Schwann Cells|| |
The effects of LIPUS on peripheral nerve regeneration are positive. However, the potential mechanisms producing the effects remain unclear and are under further investigation. As SCs play a predominant role in the processes of peripheral nerve regeneration,,,, many researchers have focused on the effects of LIPUS on SCs.
In in vivo studies, many researchers have demonstrated that LIPUS can activate SCs at the site of nerve injury. In 2002, Crisci and Ferreira  found increased numbers of SCs exhibiting morphological characteristics consistent with increased metabolic activity in LIPUS-treated animals after sciatic nerve neurotomy as compared with the control group. This indicated that LIPUS stimulated SCs during the regeneration of the sciatic nerve and that the increased SC activity accelerated the recovery of myelin sheaths. This study was the first to describe that LIPUS activated SCs in vivo. A similar result was subsequently found by Raso et al. in 2005. These authors found that LIPUS stimulated increased SC activity and an increase in SC nuclei with the characteristic reactional appearance of synthesis activity as compared to the control group. Moreover, in 2010, Chen et al. found that LIPUS improved SC proliferation at an earlier stage (first 4 weeks) after nerve injury.
As activation of SCs by LIPUS in vivo is proposed to be one of the primary mechanisms by which LIPUS promotes nerve regeneration, many groups have investigated the effect of LIPUS on SCs in vitro. In 2005, Chang et al. cultured SCs in serum deprivation culture medium to simulate an environment of mechanical trauma on a nerve injury site. They then treated the cells with LIPUS, and the results showed that LIPUS reduced the level of lactate dehydrogenase (LDH) in comparison with the sham group. This indicated a protective effect of SCs. At the same time, the 3-(4,5-dimethylthiazol-2-yl)2,5-diphenyltetrazolium bromide (MTT) assay showed increased numbers of living cells after LIPUS treatment, suggesting that LIPUS enhanced the activity of SCs. In 2009, Zhang et al. cultured rat SCs to explore how the SCs respond to in vitro LIPUS (1 MHz, 100 mW cm −2 for 5 min). The results revealed that LIPUS increased SC proliferation, indicating that SCs may become mitogenic in response to LIPUS in vitro. A similar result was found by Tsuang et al. who reported lower levels of LDH and increased values of MTT. These authors concluded that LIPUS promoted SC proliferation and prevented cell death, which is consistent with results of previous studies. In 2018, Ren et al. once again confirmed that LIPUS promoted SC viability and proliferation and explored the mechanisms of LIPUS. These authors asserted that the effects of LIPUS were a result of activation of the glycogen synthase kinase 3 beta (GSK-3β)/β-catenin signaling pathway.
Extensive studies provide evidence that LIPUS activates and promotes SC proliferation both in vivo and in vitro and that LIPUS promotes SC survival in serum deprivation culture medium, which simulates the environment of mechanical trauma on sites of injury nerve. This is of great significance for peripheral nerve regeneration and may be one of the mechanisms by which LIPUS promotes peripheral nerve regeneration after injury. Very recently, it was reported that LIPUS enhanced the secretion of exosomes from bone marrow dendritic cells (BMDCs). This may be another mechanism by which LIPUS promotes peripheral nerve regeneration.
| Lipus Induces Neurotrophic Factors|| |
A class of secreted proteins called neurotrophic factors (NFs) are essential during the development and differentiation of the central nervous system (CNS) and the peripheral nervous system (PNS). NFs consist of NGF, BDNF, and neurotrophin-3 (NT-3), among others. Since the discovery of NFs in the 1950s by Levi-Montalcini, in vitro and in vivo animal experiments have elucidated their ability to elicit positive survival and functional responses in neurons of the CNS and PNS. After nerve injury, NFs are essential in controlling the survival, proliferation, and differentiation of neural and nonneural cells involved in nerve regeneration., NGF was the first identified NF and is the dominant NF in the PNS. During peripheral nerve regeneration, NGF promotes the proliferation and differentiation of neurons and the repair of injured nerves., Upregulation of NGF leads to SC differentiation and proliferation to form regenerating neurites. In a sciatic nerve injury rat model, Chen et al. found that in vivo LIPUS increased NGF expression compared to the control group throughout the entire postinjury period (2–8 weeks). Ren et al. found that in vitro LIPUS promoted SCs secretion of NGF at both the mRNA and protein levels. In 2017, Xia et al. found that LIPUS upregulated the expression of NGFR in cultured induced pluripotent stem cell-derived neural crest stem cells. NGFR can bind to NGF, BDNF, NT-3, and NT-4 and mediate both the survival and the death of neural cells. In summary, LIPUS can promote NGF secretion both in vivo and in vitro and enhance the effects of NGF, which may be one of the mechanisms through which LIPUS enhances nerve regeneration.
BDNF plays an important role in the survival of existing neurons and in the differentiation of new neurons. BDNF is associated with axonal regeneration, myelinogenesis of medullated nerve fibers, and SC regeneration during the repair of nerve injury. In a rat sciatic nerve injury model, Ni et al. found that in vivo LIPUS increased mRNA expression of BDNF in the crushed nerve and the ipsilateral dorsal root ganglia. In 2017, in a mouse model of traumatic brain injury, Su et al. found that LIPUS increased BDNF protein levels and inhibited the progression of apoptosis. Thus, investigators have found that LIPUS can promote the expression of BDNF in both the PNS and CNS. Ren et al. found that in vitro LIPUS can promote SCs to secrete BDNF at both the mRNA and protein levels. Contrary to these findings, a study in 2009 showed that the mRNA expression of BDNF at the mRNA level was very slightly decreased after LIPUS treatment; these results seem implausible when compared to all other findings. In summary, we suggest that LIPUS can promote BDNF secretion both in vivo and in vitro and that the increase in BDNF may be beneficial for nerve regeneration after injury.
NT-3, a key NF in the PNS, is an important regulator of neural survival, development, function, and differentiation  and an important autocrine factor supporting SC survival and differentiation in the absence of axons. As recent studies have shown, NT-3 has a strong effect on neurite outgrowth,, and SCs which overexpress NT-3 induce a significantly increased number of axons at the site of injury. In 2009, Zhang et al. treated cultured SCs with LIPUS and found that the mRNA expression of NT-3 was significantly upregulated compared with the control 14 days after LIPUS stimulation. These authors postulated that the increased expression of NT-3 induced by LIPUS might establish an environment that promotes axonal sprouting and SC migration after peripheral nerve injury.
| Effect of Lipus on Cellular Signaling for Cell Activation and Mitosis|| |
It is well established that ultrasound therapy can promote cultured SC survival and proliferation. To define the mechanisms by which LIPUS activates SCs and promotes nerve regeneration after injury, cellular signaling was the focus of recent investigation [Figure 1].
|Figure 1: Cellular signaling pathways regulated by low-intensity pulsed ultrasound for peripheral nerve regeneration. LIPUS: low-intensity pulsed ultrasound; Pax: Paxillin; FAK: focal adhesion kinase; GSK3β: glycogen synthase kinase 3 beta; BDNF: brain-derived neurotrophic factor; NT-3: neurotrophin-3; PI3K: phosphatidylinositol 3-OH kinase; NGF: nerve growth factor; Trk: tyrosine kinase; MEK: mitogen-activated protein kinase/ERK kinase; ERK: extracellular signal-regulated protein kinase; CREB: cAMP-regulated enhancer B.|
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Phosphatidylinositol 3-OH kinase (PI3K)/Akt pathway
In 2008, Takeuchi et al. found that LIPUS activates PI3K/Akt pathway via an integrin in cultured chondrocytes. These authors hypothesized that LIPUS transmits signals into the cell via an integrin acting as a mechanoreceptor on the cell membrane and promotes the attachment of various focal adhesion adaptor proteins. Focal adhesion kinase (FAK) and Paxillin are then phosphorylated to initiate signal transduction to PI3K/Akt, which is known to be involved in various cellular functions including cell survival, proliferation, motility, control of cell size, and metabolism.,
Additional studies revealed that the activation of FAK stimulates the phosphorylation of GSK3β and stabilizes the Wnt/β-catenin protein to promote its nuclear translocation and to activate target-gene expression., Ren et al. also investigated the GSK-3β/β-catenin/cyclin D1 signaling pathway to investigate the mechanisms of improved SC proliferation after LIPUS. Their results indicated that LIPUS promotes phosphorylated GSK3β at serine-9, which regulates the nuclear accumulation of β-catenin to control cell biofunctions, such as gene expression, protein synthesis, and cell viability. The subsequent increased expression of cyclin D1 stimulated SC proliferation.
Extracellular signal-regulated protein kinase (ERK1/2)–cAMP-regulated enhancer B (CREB)–Trx-1 pathway
In 2016, Zhao et al. found that LIPUS activates the ERK1/2–CREB–Trx-1 pathway to promote neurite outgrowth in cultured PC12 cells. LIPUS significantly increased the levels of both phosphorylated ERK1/2 through stretch-activated ion channels and phosphorylated AKT through activation of tyrosine kinase A (TrkA) by increasing NGF. The activation of AKT and ERK1/2 phosphorylated CREB and increased expression of Trx-1. Trx-1 has several biological functions, including antioxidant, neurotrophic cofactor, cell growth promoter, and cellular apoptosis suppressor.
In 2017, Su et al. found that LIPUS treatment increased BDNF levels in a mouse model of traumatic brain injury and that BDNF mediates its effect through its high affinity for the TrkB receptor. The activation of TrkB triggers the downstream PI3K/Akt signaling pathway and increases the phosphorylation of CREB, a key transcription factor for neuroprotection and BDNF production.
| Conclusions|| |
There is significant evidence supporting the application of LIPUS to promote nerve regeneration and improve functional outcomes after surgery or trauma. The benefits of LIPUS in peripheral nerve regeneration are likely secondary to increased production of neurotrophic factors, activation of SCs, and stimulation of cellular signaling pathways for cell activation and mitosis. Given the preclinical benefits of LIPUS in the absence of any negative side effects, LIPUS shows promise as a potential clinical therapy following nerve surgery or trauma.
| Author Contributions|| |
TFL and SJX generated the concept. DYP and GTL collected the information and DYP drafted the manuscript. GTL, ABRM, SJX, and TFL reviewed and edited the manuscript. All authors read and approved the final manuscript.
| Competing Interests|| |
All authors declared no competing interests.
| Acknowledgments|| |
This article was supported by Army, Navy, NIH, Air Force, VA and Health Affairs to support the AFIRM II effort, under Award number W81XWH-13-2-0052, and NIDDK of the National Institutes of Health under award number 1R01DK105097-01A1. The U. S. Army Medical Research Acquisition Activity, 820 Chandler Street, Fort Detrick MD 21702-5014, is the awarding and administering acquisition office. Opinions, interpretations, conclusions, and recommendations are those of the author and are not necessarily endorsed by the Department of Defense and do not necessarily represent the official views of the National Institutes of Health.
| References|| |
Salonia A, Castagna G, Capogrosso P, Castiglione F, Briganti A, et al.
Prevention and management of post prostatectomy erectile dysfunction. Transl Androl Urol
2015; 4: 421–37.
Jo JK, Jeong SJ, Oh JJ, Lee SW, Lee S, et al.
Effect of starting penile rehabilitation with sildenafil immediately after robot-assisted laparoscopic radical prostatectomy on erectile function recovery: a prospective randomized trial. J Urol
2018; 199: 1600–6.
Glina S. Erectile dysfunction after radical prostatectomy: treatment options. Drugs Aging
2011; 28: 257–66.
Lei H, Xin H, Guan R, Xu Y, Li H, et al.
Low-intensity pulsed ultrasound improves erectile function in streptozotocin-induced type I diabetic rats. Urology
2015; 86: 1241, e11–8.
Huang SL, Chang CW, Lee YH, Yang FY. Protective effect of low-intensity pulsed ultrasound on memory impairment and brain damage in a rat model of vascular dementia. Radiology
2017; 282: 113–22.
Ciaramitaro P, Mondelli M, Logullo F, Grimaldi S, Battiston B, et al.
Traumatic peripheral nerve injuries: epidemiological findings, neuropathic pain and quality of life in 158 patients. J Peripher Nerv Syst
2010; 15: 120–7.
Kingham PJ, Terenghi G. Bioengineered nerve regeneration and muscle reinnervation. J Anat
2006; 209: 511–26.
Torres RY, Miranda GE. Epidemiology of traumatic peripheral nerve injuries evaluated by electrodiagnostic studies in a tertiary care hospital clinic. Bol Asoc Med P R
2015; 107: 79–84.
Fu SY, Gordon T. Contributing factors to poor functional recovery after delayed nerve repair: prolonged denervation. J Neurosci
1995; 15: 3886–95.
Gordon T, Tyreman N, Raji MA. The basis for diminished functional recovery after delayed peripheral nerve repair. J Neurosci
2011; 31: 5325–34.
Kobayashi J, Mackinnon SE, Watanabe O, Ball DJ, Gu XM, et al.
The effect of duration of muscle denervation on functional recovery in the rat model. Muscle Nerve
1997; 20: 858–66.
Aydin MA, Mackinnon SE, Gu XM, Kobayashi J, Kuzon WM Jr. Force deficits in skeletal muscle after delayed reinnervation. Plast Reconstr Surg
2004; 113: 1712–8.
Elzinga K, Tyreman N, Ladak A, Savaryn B, Olson J, et al.
Brief electrical stimulation improves nerve regeneration after delayed repair in Sprague Dawley rats. Exp Neurol
2015; 269: 142–53.
Rusovan A, Kanje M. Magnetic fields stimulate peripheral nerve regeneration in hypophysectiomized rats. Neuroreport
1992; 3: 1039–41.
Sene GA, Sousa FF, Fazan VS, Barbieri CH. Effects of laser therapy in peripheral nerve regeneration. Acta Ortop Bras
2013; 21: 266–70.
Ma F, Xiao Z, Chen B, Hou X, Dai J, et al.
Linear ordered collagen scaffolds loaded with collagen-binding basic fibroblast growth factor facilitate recovery of sciatic nerve injury in rats. Tissue Eng Part A
2014; 20: 1253–62.
Montava M, Garcia S, Mancini J, Jammes Y, Courageot J, et al.
Vitamin D3 potentiates myelination and recovery after facial nerve injury. Eur Arch Otorhinolaryngol
2015; 272: 2815–23.
Wang Y, Shen W, Yang L, Zhao H, Gu W, et al.
The protective effects of achyranthes bidentata polypeptides on rat sciatic nerve crush injury causes modulation of neurotrophic factors. Neurochem Res
2013; 38: 538–46.
Ying ZM, Lin T, Yan SG. Low-intensity pulsed ultrasound therapy: a potential strategy to stimulate tendon-bone junction healing. J Zhejiang Univ Sci B
2012; 13: 955–63.
Khanna A, Nelmes RT, Gougoulias N, Maffulli N, Gray J. The effects of LIPUS on soft-tissue healing: a review of literature. Br Med Bull
2009; 89: 169–82.
Lv Y, Zhao P, Chen G, Sha Y, Yang L. Effects of low-intensity pulsed ultrasound on cell viability, proliferation and neural differentiation of induced pluripotent stem cells-derived neural crest stem cells. Biotechnol Lett
2013; 35: 2201–12.
Lv Y, Nan P, Chen G, Sha Y, Xia B, et al
. In vivo
repair of rat transected sciatic nerve by low-intensity pulsed ultrasound and induced pluripotent stem cells-derived neural crest stem cells. Biotechnol Lett
2015; 37: 2497–506.
Hannemann PF, Mommers EH, Schots JP, Brink PR, Poeze M. The effects of low-intensity pulsed ultrasound and pulsed electromagnetic fields bone growth stimulation in acute fractures: a systematic review and meta-analysis of randomized controlled trials. Arch Orthop Trauma Surg
2014; 134: 1093–106.
Jia XL, Chen WZ, Zhou K, Wang ZB. Effects of low-intensity pulsed ultrasound in repairing injured articular cartilage. Chin J Traumatol
2005; 8: 175–8.
Takakura Y, Matsui N, Yoshiya S, Fujioka H, Muratsu H, et al.
Low-intensity pulsed ultrasound enhances early healing of medial collateral ligament injuries in rats. J Ultrasound Med
2002; 21: 283–8.
Kristiansen TK, Ryaby JP, McCabe J, Frey JJ, Roe LR. Accelerated healing of distal radial fractures with the use of specific, low-intensity ultrasound. A multicenter, prospective, randomized, double-blind, placebo-controlled study. J Bone Joint Surg Am
1997; 79: 961–73.
Hoffman PN. A conditioning lesion induces changes in gene expression and axonal transport that enhance regeneration by increasing the intrinsic growth state of axons. Exp Neurol
2010; 223: 11–8.
Hanz S, Fainzilber M. Retrograde signaling in injured nerve--the axon reaction revisited. J Neurochem
2006; 99: 13–9.
Stoll G, Griffin JW, Li CY, Trapp BD. Wallerian degeneration in the peripheral nervous system: participation of both Schwann cells and macrophages in myelin degradation. J Neurocytol
1989; 18: 671–83.
Yudin D, Hanz S, Yoo S, Iavnilovitch E, Willis D, et al.
Localized regulation of axonal RanGTPase controls retrograde injury signaling in peripheral nerve. Neuron
2008; 59: 241–52.
Jones S, Eisenberg HM, Jia X. Advances and future applications of augmented peripheral nerve regeneration. Int J Mol Sci
2016; 17: 1494.
Marx J. Helping neurons find their way. Science
1995; 268: 971–3.
Geuna S, Raimondo S, Ronchi G, Di Scipio F, Tos P, et al.
Chapter 3: histology of the peripheral nerve and changes occurring during nerve regeneration. Int Rev Neurobiol
2009; 87: 27–46.
Lee SK, Wolfe SW. Peripheral nerve injury and repair. J Am Acad Orthop Surg
2000; 8: 243–52.
Caillaud M, Richard L, Vallat JM, Desmouliere A, Billet F. Peripheral nerve regeneration and intraneural revascularization. Neural Regen Res
2019; 14: 24–33.
Toews AD, Barrett C, Morell P. Monocyte chemoattractant protein 1 is responsible for macrophage recruitment following injury to sciatic nerve. J Neurosci Res
1998; 53: 260–7.
Klein D, Martini R. Myelin and macrophages in the PNS: an intimate relationship in trauma and disease. Brain Res
2016; 1641: 130–8.
Fu SY, Gordon T. The cellular and molecular basis of peripheral nerve regeneration. Mol Neurobiol
1997; 14: 67–116.
Burnett MG, Zager EL. Pathophysiology of peripheral nerve injury: a brief review. Neurosurg Focus
2004; 16: E1.
Gaudet AD, Popovich PG, Ramer MS. Wallerian degeneration: gaining perspective on inflammatory events after peripheral nerve injury. J Neuroinflammation
2011; 8: 110.
Madl CM, Heilshorn SC. Matrix interactions modulate neurotrophin-mediated neurite outgrowth and pathfinding. Neural Regen Res
2015; 10: 514–7.
Hopker VH, Shewan D, Tessier-Lavigne M, Poo M, Holt C. Growth-cone attraction to netrin-1 is converted to repulsion by laminin-1. Nature
1999; 401: 69–73.
Qing L, Chen H, Tang J, Jia X. Exosomes and their MicroRNA cargo: new players in peripheral nerve regeneration. Neurorehabil Neural Repair
2018; 32: 765–76.
Fry WJ. Intense ultrasound; a new tool for neurological research. J Ment Sci
1954; 100: 85–96.
Newell JA. Ultrasonics in Medicine. Phys Med Biol
1963; 8: 241–64.
Wells PN. Ultrasonics in medicine and biology. Phys Med Biol
1977; 22: 629–69.
Kremkau FW. Cancer therapy with ultrasound: a historical review. J Clin Ultrasound
1979; 7: 287–300.
ter Haar G. Therapeutic applications of ultrasound. Prog Biophys Mol Biol
2007; 93: 111–29.
Lin G, Reed-Maldonado AB, Lin M, Xin Z, Lue TF. Effects and mechanisms of low-intensity pulsed ultrasound for chronic prostatitis and chronic pelvic pain syndrome. Int J Mol Sci
2016; 17: 1057.
Ikeda K, Takayama T, Suzuki N, Shimada K, Otsuka K, et al.
Effects of low-intensity pulsed ultrasound on the differentiation of C2C12 cells. Life Sci
2006; 79: 1936–43.
Urita A, Iwasaki N, Kondo M, Nishio Y, Kamishima T, et al.
Effect of low-intensity pulsed ultrasound on bone healing at osteotomy sites after forearm bone shortening. J Hand Surg Am
2013; 38: 498–503.
Zhou S, Schmelz A, Seufferlein T, Li Y, Zhao J, et al.
Molecular mechanisms of low intensity pulsed ultrasound in human skin fibroblasts. J Biol Chem
2004; 279: 54463–9.
Ikai H, Tamura T, Watanabe T, Itou M, Sugaya A, et al.
Low-intensity pulsed ultrasound accelerates periodontal wound healing after flap surgery. J Periodontal Res
2008; 43: 212–6.
Nakao J, Fujii Y, Kusuyama J, Bandow K, Kakimoto K, et al.
Low-intensity pulsed ultrasound (LIPUS) inhibits LPS-induced inflammatory responses of osteoblasts through TLR4-MyD88 dissociation. Bone
2014; 58: 17–25.
Daeschler SC, Harhaus L, Schoenle P, Boecker A, Kneser U, et al.
Ultrasound and shock-wave stimulation to promote axonal regeneration following nerve surgery: a systematic review and meta-analysis of preclinical studies. Sci Rep
2018; 8: 3168.
Young RR, Henneman E. Functional effects of focused ultrasound on mammalian nerves. Science
1961; 134: 1521–2.
Mourad PD, Lazar DA, Curra FP, Mohr BC, Andrus KC, et al.
Ultrasound accelerates functional recovery after peripheral nerve damage. Neurosurgery
2001; 48: 1136–40.
Crisci AR, Ferreira AL. Low-intensity pulsed ultrasound accelerates the regeneration of the sciatic nerve after neurotomy in rats. Ultrasound Med Biol
2002; 28: 1335–41.
Chang CJ, Hsu SH, Lin FT, Chang H, Chang CS. Low-intensity-ultrasound-accelerated nerve regeneration using cell-seeded poly(D, L-lactic acid-co
-glycolic acid) conduits: an in vivo
and in vitro
study. J Biomed Mater Res B Appl Biomater
2005; 75: 99–107.
Terzis JK, Sun DD, Thanos PK. Historical and basic science review: past, present, and future of nerve repair. J Reconstr Microsurg
1997; 13: 215–25.
Tsuang YH, Liao LW, Chao YH, Sun JS, Cheng CK, et al.
Effects of low intensity pulsed ultrasound on rat Schwann cells metabolism. Artif Organs
2011; 35: 373–83.
Tountas CP, Bergman RA, Lewis TW, Stone HE, Pyrek JD, et al.
A comparison of peripheral nerve repair using an absorbable tubulization device and conventional suture in primates. J Appl Biomater
1993; 4: 261–8.
Aldini NN, Perego G, Cella GD, Maltarello MC, Fini M, et al.
Effectiveness of a bioabsorbable conduit in the repair of peripheral nerves. Biomaterials
1996; 17: 959–62.
Evans GR, Brandt K, Katz S, Chauvin P, Otto L, et al.
Bioactive poly(L-lactic acid) conduits seeded with Schwann cells for peripheral nerve regeneration. Biomaterials
2002; 23: 841–8.
Jiang W, Wang Y, Tang J, Peng J, Wang Y, et al.
Low-intensity pulsed ultrasound treatment improved the rate of autograft peripheral nerve regeneration in rat. Sci Rep
2016; 6: 22773.
Chang CJ, Hsu SH. The effects of low-intensity ultrasound on peripheral nerve regeneration in poly(DL-lactic acid-co-glycolic acid) conduits seeded with Schwann cells. Ultrasound Med Biol
2004; 30: 1079–84.
Park SC, Oh SH, Seo TB, Namgung U, Kim JM, et al.
Ultrasound-stimulated peripheral nerve regeneration within asymmetrically porous PLGA/Pluronic F127 nerve guide conduit. J Biomed Mater Res B Appl Biomater
2010; 94: 359–66.
Lowdon IM, Seaber AV, Urbaniak JR. An improved method of recording rat tracks for measurement of the sciatic functional index of de Medinaceli. J Neurosci Methods
1988; 24: 279–81.
Raso VV, Barbieri CH, Mazzer N, Fasan VS. Can therapeutic ultrasound influence the regeneration of peripheral nerves? J Neurosci Methods
2005; 142: 185–92.
Chen WZ, Qiao H, Zhou W, Wu J, Wang ZB. Upgraded nerve growth factor expression induced by low-intensity continuous-wave ultrasound accelerates regeneration of neurotometicly injured sciatic nerve in rats. Ultrasound Med Biol
2010; 36: 1109–17.
Ni XJ, Wang XD, Zhao YH, Sun HL, Hu YM, et al.
The effect of low-intensity ultrasound on brain-derived neurotropic factor expression in a rat sciatic nerve crushed injury model. Ultrasound Med Biol
2017; 43: 461–8.
Chiang PK, Yang FY. A potential treatment of low intensity pulsed ultrasound on cavernous nerve injury for erectile dysfunction. Med Hypotheses
2019; 122: 19–21.
Reichert F, Saada A, Rotshenker S. Peripheral nerve injury induces Schwann cells to express two macrophage phenotypes: phagocytosis and the galactose-specific lectin MAC-2. J Neurosci
1994; 14: 3231–45.
Ronchi G, Nicolino S, Raimondo S, Tos P, Battiston B, et al.
Functional and morphological assessment of a standardized crush injury of the rat median nerve. J Neurosci Methods
2009; 179: 51–7.
Vrbova G, Mehra N, Shanmuganathan H, Tyreman N, Schachner M, et al.
Chemical communication between regenerating motor axons and Schwann cells in the growth pathway. Eur J Neurosci
2009; 30: 366–75.
Rath EM, Kelly D, Bouldin TW, Popko B. Impaired peripheral nerve regeneration in a mutant strain of mice (Enr) with a Schwann cell defect. J Neurosci
1995; 15: 7226–37.
Zhang H, Lin X, Wan H, Li JH, Li JM. Effect of low-intensity pulsed ultrasound on the expression of neurotrophin-3 and brain-derived neurotrophic factor in cultured Schwann cells. Microsurgery
2009; 29: 479–85.
Ren C, Chen X, Du N, Geng S, Hu Y, et al.
Low-intensity pulsed ultrasound promotes Schwann cell viability and proliferation via the GSK-3beta/beta-catenin signaling pathway. Int J Biol Sci
2018; 14: 497–507.
Weissmiller AM, Wu C. Current advances in using neurotrophic factors to treat neurodegenerative disorders. Transl Neurodegener
2012; 1: 14.
Levi-Montalcini R, Hamburger V. Selective growth stimulating effects of mouse sarcoma on the sensory and sympathetic nervous system of the chick embryo. J Exp Zool
1951; 116: 321–61.
Hausner T, Nogradi A. The use of shock waves in peripheral nerve regeneration: new perspectives? Int Rev Neurobiol
2013; 109: 85–98.
Petruska JC, Mendell LM. The many functions of nerve growth factor: multiple actions on nociceptors. Neurosci Lett
2004; 361: 168–71.
Unezaki S, Yoshii S, Mabuchi T, Saito A, Ito S. Effects of neurotrophic factors on nerve regeneration monitored by in vivo
imaging in thy1-YFP transgenic mice. J Neurosci Methods
2009; 178: 308–15.
Campbell WW. Evaluation and management of peripheral nerve injury. Clin Neurophysiol
2008; 119: 1951–65.
Xia B, Zou Y, Xu Z, Lv Y. Gene expression profiling analysis of the effects of low-intensity pulsed ultrasound on induced pluripotent stem cell-derived neural crest stem cells. Biotechnol Appl Biochem
2017; 64: 927–37.
Tomellini E, Lagadec C, Polakowska R, Le Bourhis X. Role of p75 neurotrophin receptor in stem cell biology: more than just a marker. Cell Mol Life Sci
2014; 71: 2467–81.
Acheson A, Conover JC, Fandl JP, DeChiara TM, Russell M, et al.
A BDNF autocrine loop in adult sensory neurons prevents cell death. Nature
1995; 374: 450–3.
Alrashdan MS, Sung MA, Kwon YK, Chung HJ, Kim SJ, et al.
Effects of combining electrical stimulation with BDNF gene transfer on the regeneration of crushed rat sciatic nerve. Acta Neurochir (Wien)
2011; 153: 2021–9.
Yi S, Yuan Y, Chen Q, Wang X, Gong L, et al.
Regulation of schwann cell proliferation and migration by miR-1 targeting brain-derived neurotrophic factor after peripheral nerve injury. Sci Rep
2016; 6: 29121.
Su WS, Wu CH, Chen SF, Yang FY. Transcranial ultrasound stimulation promotes brain-derived neurotrophic factor and reduces apoptosis in a mouse model of traumatic brain injury. Brain Stimul
2017; 10: 1032–41.
McAllister AK, Lo DC, Katz LC. Neurotrophins regulate dendritic growth in developing visual cortex. Neuron
1995; 15: 791–803.
McAllister AK, Katz LC, Lo DC. Neurotrophins and synaptic plasticity. Annu Rev Neurosci
1999; 22: 295–318.
Sahenk Z, Nagaraja HN, McCracken BS, King WM, Freimer ML, et al.
NT-3 promotes nerve regeneration and sensory improvement in CMT1A mouse models and in patients. Neurology
2005; 65: 681–9.
Markus A, Patel TD, Snider WD. Neurotrophic factors and axonal growth. Curr Opin Neurobiol
2002; 12: 523–31.
Takeuchi R, Ryo A, Komitsu N, Mikuni-Takagaki Y, Fukui A, et al.
Low-intensity pulsed ultrasound activates the phosphatidylinositol 3 kinase/Akt pathway and stimulates the growth of chondrocytes in three-dimensional cultures: a basic science study. Arthritis Res Ther
2008; 10: R77.
Downward J. PI 3-kinase, Akt and cell survival. Semin Cell Dev Biol
2004; 15: 177–82.
Gustin JA, Korgaonkar CK, Pincheira R, Li Q, Donner DB. Akt regulates basal and induced processing of NF-kappaB2 (p100) to p52. J Biol Chem
2006; 281: 16473–81.
Gao C, Chen G, Kuan SF, Zhang DH, Schlaepfer DD, et al.
FAK/PYK2 promotes the Wnt/beta-catenin pathway and intestinal tumorigenesis by phosphorylating GSK3beta. Elife
2015; 4: e10072.
Fonar Y, Frank D. FAK and WNT signaling: the meeting of two pathways in cancer and development. Anticancer Agents Med Chem
2011; 11: 600–6.
Zhao L, Feng Y, Hu H, Shi A, Zhang L, et al.
Low-intensity pulsed ultrasound enhances nerve growth factor-induced neurite outgrowth through mechanotransduction-mediated ERK1/2-CREB-Trx-1 signaling. Ultrasound Med Biol
2016; 42: 2914–25.
Jeon SJ, Rhee SY, Seo JE, Bak HR, Lee SH, et al.
Oroxylin a increases BDNF production by activation of MAPK-CREB pathway in rat primary cortical neuronal culture. Neurosci Res
2011; 69: 214–22.