Next Article in Journal
Aptamers Targeting Cardiac Biomarkers as an Analytical Tool for the Diagnostics of Cardiovascular Diseases: A Review
Next Article in Special Issue
A Comparative Feasibility Study for Transcranial Extracorporeal Shock Wave Therapy
Previous Article in Journal
Implication of Different Tumor Biomarkers in Drug Resistance and Invasiveness in Primary and Metastatic Colorectal Cancer Cell Lines
Previous Article in Special Issue
Translational Applications of Extracorporeal Shock Waves in Dental Medicine: A Literature Review
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Biomedicines
Volume 10
Issue 5
10.3390/biomedicines10051084
biomedicines-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Systematic Review

The Effects of the Exposure of Musculoskeletal Tissue to Extracorporeal Shock Waves

by
Tobias Wuerfel
,
Christoph Schmitz
* and
Leon L. J. Jokinen
Extracorporeal Shock Wave Research Unit, Chair of Neuroanatomy, Institute of Anatomy, Faculty of Medicine, Ludwig-Maximilians-University of Munich, Munich 80336, Germany
*
Author to whom correspondence should be addressed.
Biomedicines 2022, 10(5), 1084; https://doi.org/10.3390/biomedicines10051084
Submission received: 20 April 2022 / Revised: 1 May 2022 / Accepted: 4 May 2022 / Published: 6 May 2022
(This article belongs to the Special Issue Translational Research in Shock Wave Medicine)
Browse Figure
Versions Notes

Abstract

:
Extracorporeal shock wave therapy (ESWT) is a safe and effective treatment option for various pathologies of the musculoskeletal system. Many studies address the molecular and cellular mechanisms of action of ESWT. However, to date, no uniform concept could be established on this matter. In the present study, we perform a systematic review of the effects of exposure of musculoskeletal tissue to extracorporeal shock waves (ESWs) reported in the literature. The key results are as follows: (i) compared to the effects of many other forms of therapy, the clinical benefit of ESWT does not appear to be based on a single mechanism; (ii) different tissues respond to the same mechanical stimulus in different ways; (iii) just because a mechanism of action of ESWT is described in a study does not automatically mean that this mechanism is relevant to the observed clinical effect; (iv) focused ESWs and radial ESWs seem to act in a similar way; and (v) even the most sophisticated research into the effects of exposure of musculoskeletal tissue to ESWs cannot substitute clinical research in order to determine the optimum intensity, treatment frequency and localization of ESWT.

1. Introduction

Extracorporeal shock wave therapy (ESWT) is a safe and effective treatment option for various pathologies of the musculoskeletal system. The beginning of the use of extracorporeal shock waves (ESWs) in medicine was in kidney stone fragmentation; the corresponding method is called Extracorporeal Shock Wave Lithotrypsy (ESWL). After ESWL was performed on dogs for the first time in 1976, four years later, the first human patient was successfully freed from his kidney stone disease using ESWL [1]. Expanded to other stone diseases in the gallbladder [2], pancreas [3], bile duct [4] and salivary glands [5], urologists found (more or less by chance) that the application of ESWs in the area of ureteral stones caused changes in the os ileum [6]. Specifically, when bones were exposed to ESWs, primary osteocyte damage followed by osteoblast stimulation was observed [6]. This resulted in the demonstration of the stimulation of fracture healing with ESWs in animal models [7]. Since these beginnings, the application of ESWs has been expanded to a variety of pathologies of the musculoskeletal system, with the treatment of non-unions (reviewed in [8]) and tendinopathies (reviewed in [9,10,11]) being, by far, the largest groups of indications. The treatment of pathologies of the musculoskeletal system with ESWs is commonly referred to as Extracorporeal Shock Wave Therapy (ESWT) and is thus distinguished from ESWL.
This short description of the history of ESWT demonstrates that the development of this treatment modality has not followed the classical drug discovery process, from initial target identification and validation, through assay development, high throughput screening, hit identification, lead optimization and finally the selection of a candidate molecule for clinical development [12]. Rather, progress in clinical research on ESWT was either accompanied or followed by basic and preclinical research into the potential mechanisms of action of ESWs on the target tissue. The latter was addressed in several recent reviews (e.g., [13,14,15,16,17]). Considering the fact that this study summarizes and discusses 181 studies addressing the effects of the exposure of musculoskeletal tissue on ESWs [6,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48,49,50,51,52,53,54,55,56,57,58,59,60,61,62,63,64,65,66,67,68,69,70,71,72,73,74,75,76,77,78,79,80,81,82,83,84,85,86,87,88,89,90,91,92,93,94,95,96,97,98,99,100,101,102,103,104,105,106,107,108,109,110,111,112,113,114,115,116,117,118,119,120,121,122,123,124,125,126,127,128,129,130,131,132,133,134,135,136,137,138,139,140,141,142,143,144,145,146,147,148,149,150,151,152,153,154,155,156,157,158,159,160,161,162,163,164,165,166,167,168,169,170,171,172,173,174,175,176,177,178,179,180,181,182,183,184,185,186,187,188,189,190,191,192,193,194,195,196,197], the limited number of references in the aforementioned reviews (between 38 [13] and 93 [16]) indicate that these reviews are either outdated or incomplete.
The aim of this study is to provide clinicians, basic science researchers and other stakeholders in healthcare with a comprehensive overview of what is known today regarding the effects of the exposure of musculoskeletal tissue to ESWs. This should help to further understand this fascinating, non-invasive treatment modality that is highly efficient and has a very good safety profile in the treatment of many pathologies of the musculoskeletal system. Because of the variety of different tissues that make up the musculoskeletal system, as well as of the different motivations for performing ESWT (ranging from pain relief to tissue regeneration), we divided our review into three areas focusing on bone and cartilage, connective tissue and muscle/nerve tissue.

2. Materials and Methods

PubMed and Web of Science were searched for “shock wave OR shock waves OR shockwave OR shockwaves NOT urol* NOT stone NOT review NOT clinical trial” from the days of inception of these databases until 30 September 2021, according to the PRISMA (Preferred Reporting Items for Systematic Reviews and Meta-Analyses) [198] guidelines. Duplicates were excluded.
For each identified publication, it was determined by reading the title and abstract whether the publication represented a study on the effects of exposure of musculoskeletal tissue to extracorporeal shock waves; the studies only addressing the treatment of skin with ESWT were excluded. All this was independently undertaken by T.W. and C.S. The results were compared and discussed until an agreement was achieved.
Subsequently, all the selected studies were classified with regard to the type of tissue (bone and cartilage, connective tissue or muscle/nerve tissue, respectively) that was exposed to ESWs. Furthermore, it was determined for each selected study whether (i) morphological, functional and radiological findings, (ii) findings of molecular biological investigations and/or (iii) findings of histological investigations were reported. All this was independently undertaken by T.W. and L.J., and the results were compared and discussed until an agreement was achieved.
The strategy of the literature search is summarized in Figure 1.

3. Results

The results of this systematic review are summarized in Table 1, Table 2 and Table 3, with a distinction being made between effects of the exposure of bone and cartilage tissue (Table 1), connective tissue (Table 2) and muscle and nerve tissue (Table 3) to ESWs. Within each table, the results are arranged chronologically, with the most recent findings presented first. More details of the studies listed in Table 1, Table 2 and Table 3 are provided in Tables S1–S3.

3.1. Effects of the Exposure of Bone and Cartilage Tissue to Extracorporeal Shock Waves

Our systematic review revealed 100 studies that addressed the effects of ESWs on bone and cartilage tissue [6,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48,49,50,51,52,53,54,55,56,57,58,59,60,61,62,63,64,65,66,67,68,69,70,71,72,73,74,75,76,77,78,79,80,81,82,83,84,85,86,87,88,89,90,91,92,93,94,95,96,97,98,99,100,101,102,103,104,105,106,107,108,109,110,111,112,113,114,115,116]. These studies were published between 1988 and 2021, with 51 (51%) of these studies published during the last ten years (2012–2021). Eighty-five of these studies (85%) applied fESWs, eleven (11%) of these studies applied rESWs, two (2%) of these studies applied both fESWs and rESWs, and in two (2%) of these studies it was not described whether fESWs or rESWs were applied. The majority of these studies (64 of 100, i.e., 64%) described animal experiments; primary or secondary cell-culture experiments were described in 23 (23%) or 7 (7%) of these studies, respectively. Three (3%) of these studies combined animal experiments with primary cell-culture experiments; one (1%) of these studies combined animal experiments with secondary cell-culture experiments, and two (2%) of these studies were conducted ex vivo without animal experiments and cell-culture experiments (details are provided in Table S1). Very different effects of ESWs on bone and cartilage tissue were addressed in these 100 studies; these effects are summarized in Table 1.

3.2. Effects of Exposure of Connective Tissue to Extracorporeal Shock Waves

Our systematic review revealed 39 studies that addressed effects of ESWs on connective tissue [117,118,119,120,121,122,123,124,125,126,127,128,129,130,131,132,133,134,135,136,137,138,139,140,141,142,143,144,145,146,147,148,149,150,151,152,153,154,155]. These studies were published between 1994 and 2021, with 18 (46.2%) of these studies published during the last ten years (2012–2021). Thirty (76.9%) of these study applied fESWs, and nine (23.1%) of these studies applied rESWs. The majority of these studies (24, i.e., 61.5%) described animal experiments; primary or secondary cell-culture experiments were described in nine (23.1%) or three (7.7%) of these studies, respectively. One study each (2.6% each) described cell-culture experiments (not further specified), experiments on fertilized chicken embryos and a human experiment (details are provided in Table S2). As in case of the exposure of bone and cartilage tissue to ESWs (Table 1), very different effects of ESWs on connective tissue were addressed in these 39 studies. These effects are summarized in Table 2.

3.3. Effects of Exposure of Muscle and Nerve Tissue to Extracorporeal Shock Waves

Our systematic review revealed 42 studies that addressed effects of ESWs on muscle and nerve tissue [156,157,158,159,160,161,162,163,164,165,166,167,168,169,170,171,172,173,174,175,176,177,178,179,180,181,182,183,184,185,186,187,188,189,190,191,192,193,194,195,196,197]. These studies were published between 1998 and 2021, with 25 (59.5%) of these studies published during the last ten years (2012–2021). Twenty-eight (66.7%) of these study applied fESWs, 10 (23.8%) of these studies applied rESWs, and in four (9.5%) of these studies it was not described whether fESWs or rESWs were applied. The vast majority (39, i.e., 92.9%) of these studies described animal experiments; two (4.8%) of these studies described primary cell-culture experiments, and one (2.4%) of these studies combined animal experiments with primary cell-culture experiments (details are provided in Table S3). As in case of the exposure of bone, cartilage tissue and connective tissue to ESWs (Table 1 and Table 2), very different effects of ESWs on muscle and nerve tissue were addressed in these 42 studies. These effects are summarized in Table 3.

4. Discussion

Based on the results summarized in Table 1, Table 2 and Table 3, we established ten take-home messages regarding the effects of exposure of musculoskeletal tissue to extracorporeal shock waves. These take-home messages are summarized in Table 4 and discussed below.
The first take-home message of this study is that compared to the effects of many other forms of therapy; the clinical benefit of extracorporeal shock wave therapy does not appear to be based on a single mechanism. Most of the basic studies on medical therapies run exactly opposite to the studies on the mode of action of ESWT. In preclinical research, mechanisms are often sought that are later clinically tested for their benefit. However, for the treatment indications of ESWT on the musculoskeletal system, mainly the clinical success is known to date, while, in contrast, the molecular and cellular causes for this success are widely unknown. Thus, studies of the mode of action of ESWT are based on rational considerations of the mechanisms by which clinical success might occur. In the numerous studies, a variety of effects were described, most of which are desirable for the respective indication. Many of these mechanisms are not causally related, so that it is obvious that the combination of different effects leads to the therapeutic success of ESWT.
The second take-home message of this study is that different tissues respond to the same mechanical stimulus in different ways. Based on many years of clinical experience and numerous clinical studies, various pathologies of the musculoskeletal system are known nowadays that can be successfully treated with ESWT [8,9,10,11]. These indications include mainly degenerations and injuries of muscle, bone and cartilage tissue. From basic research, a wide variety of effects at the molecular and cellular levels were described to date, whereby the effects of the ESWs differ in each case from the tissue treated. On the one hand, very tissue-specific reactions were observed. For example, while the enhancement of the osseous differentiation of stem cells occurred in the bone [23,48], the differentiation of stem cells into the osteocytic lineage was not observed in tendon tissue [125]. On the other hand, there are similar effects that were observed, despite the different tissues, such as an increase in the expression of vascular endothelial growth factor (VEGF) after exposure to ESWs in the bone and cartilage tissue [42,58,69,73,90,97], nerve tissue [170,175] and connective tissue [137]. This leads to the conclusion that ESWs generally promote angiogenesis, despite the fact that some studies described no effects after the exposure of tissue to ESWs on the expression of VEGF [118,119,162], or even the reduced expression of VEGF [26]. In addition, the condition of the treated tissue also seems to play a role. For example, healthy tenocytes responded to the exposure to ESWs with a different protein expression pattern than tenocytes from tendinopathic or ruptured tendon tissues [134,141]. This highlights one of the key problems in evaluating studies of the effects of ESWs on the musculoskeletal system: due to differences in design and the prevailing conditions in these studies, comparisons are sometimes difficult to make.
The third take-home message of this study is that just because a mechanism of action of extracorporeal shock wave therapy was described in a study does not automatically mean that this mechanism was relevant to the observed clinical effect. Some of the many effects described include effects that, considered in isolation, would not be desirable for the success of the therapy. However, as clinically a treatment success is mostly shown, other mechanisms must play a greater role for the effect of ESWT. One example is the increased vascularization of tendon tissue after exposure to ESWs [97,133]; although increased vascularization is usually associated with tendon inflammation [199], clinical findings were shown to improve after treatment [148]. Likewise, in the treatment of muscular spasticity by ESWT, it is unlikely that a stimulating effect of ESWs on, for example, stem cells, has anything to do with the reduced muscle tone after ESWT (e.g., [200]). Thus, when deducing the modes of action of ESWT in certain pathologies of the musculoskeletal system, one should always relate certain modes of action to the pathology under investigation in order to not obtain incorrect conclusions.
The fourth take-home message of this study is that focused and radial extracorporeal shock wave therapy seem to act in a similar way. Numerous effects were described for both fESWT and rESWT, however, more effects were described for fESWT (Table 1, Table 2 and Table 3). This may be due to the fact that fESWT was developed before rESWT [10]. From a physics point of view, these two forms of ESWT appear to differ greatly. Focused ESWs are generated by three methods that are named electrohydraulic, electromagnetic and piezoelectric [10]. Additionally, unlike rESWs, fESWs are generated in water that is inside the applicator [201]. In contrast, rESWs are generated by the acceleration of a projectile in a tube (through compressed air or a magnetic field), and the projectile hits an applicator at the end of the tube. Through contact with the skin via contact gel (to facilitate transmission), the rESWs are transmitted into the treated tissue [201]. As a result of these different mechanisms of ESW generation, rESWT has more of a superficial effect on tissues, while fESWT can also affect deeper tissues [10,201].
Some authors argued that rESWs should not be called shock waves, since they lack the characteristic physical features of true shock waves, including a short rise time in the amount of nanoseconds, a high peak pressure and non-linearity [202]. The physical definition of a “true” shock wave is as follows [203]: a high positive peak pressure (P+), sometimes more than 100 Megapascal (Mpa), but more often approximately 50 to 80 MPa; a fast initial rise in pressure (Tr) during a period of less than 10 nanoseconds (ns); a low tensile amplitude (P−, up to 10 MPa); a short life cycle (I) of approximately 10 microseconds (μs); and a broad frequency spectrum, typically in the range of 16 Hertz (Hz) to 20 MHz. It is well-known that rESWs are not “true” shock waves in the strict physical sense outlined above [202]. This is because rESWs show a lower positive peak pressure (~10 MPa) and a substantially longer rise time (~600 ns), and have thus been termed radial pressure waves by some authors [204]. However, in 2007, it was already noticed that for treatment protocols at low-energy settings, neither piezoelectric nor electromagnetic fESWT devices generated true shock waves according to the physical criteria set out above [202]. With respect to the various ESWT devices’ abilities to generate shock waves as opposed to pressure waves, the initial concept can thus be refined into a concept that considers high-energy settings as a prerequisite for the generation of true shock waves. For clinical applications of ESWT, however, a more feasible concept of therapeutic shock wave technology needs to factor in two more considerations: that biological cells and tissues can differentiate between true shock waves and pressure waves, but cannot differentiate between radial or focused wave forms. As to the former point, it is certainly reasonable to differentiate between shock waves and pressure waves in terms of the differences in positive peak pressure delivered to the pathologic site. However, the question arises whether therapy success in many pathologies of the musculoskeletal system requires “true” shock waves [205]. It appears that this is not the case. With respect to the differentiation between rESWs and fESWs, under plain geometric considerations it is highly unlikely that tissues and cells can differentiate whether they are affected by focused or by radial acoustic waves—the only difference is in the number of affected cells. In consequence, it appears that, clinically, “a wave is a wave” regardless of whether it is generated with an fESWT device or a rESWT device. Much more important is whether sufficient ESWT energy is achieved where it is needed in the body.
Cavitation can be generated only during the shock wave’s tensile phase [206]. Of note, both fESWs and rESWs can generate vaporous cavitation [206]. Vaporous cavitation is assumed to play an important role in mediating molecular and cellular mechanisms of action of ESWT in biological tissues, presumably via the mechanical activation of membrane-bound signaling molecules, which, in turn, elicit cellular responses [206]. Yet, many questions remain open concerning the therapeutic effects of vaporous cavitation during ESWT. For example, it was found that tissues exposed to ESWs show a subsequent decrease in proinflammatory neuropeptides, similar to a “wash-out” effect [193]. This correlates well with the long-term analgesic effect mediated by ESWT in tendinopathies [10]. Yet, it remains unknown which effects vaporous cavitation has on the unmyelinated terminal endings of nociceptive fibers (i.e., C fibers) in the peripheral nervous system. More generally speaking, it is still unknown as to whether the therapeutic benefits of ESWT are due mainly to the positive (i.e., shear stress) or negative (i.e., cavitation) pressures, or a combination of both, in order to optimize treatment protocols [10]. Because of the potentially deleterious side effects of vaporous cavitation on the body, it is imperative to realize that both fESWT devices and rESWT devices can in fact generate vaporous cavitation in the treated tissue.
In summary, it is reasonable to hypothesize that further research into the effects of exposure of musculoskeletal tissue to fESWs and rESWs will demonstrate more similarities than dissimilarities between these modalities. Nevertheless, due to the differing energy distribution of both treatment forms in the target tissue, different energy-dependent effects may occur (e.g., [102]).
The fifth take-home message of this study is that extracorporeal shock wave therapy stimulates both progenitor and differentiated cells and has positive effects on the pathologies of bone and cartilage. A central aspect for the treatment of degenerations and injuries of muscles, tendons, bones and cartilage using ESWT is the activation of the respective tissue-specific cells. The mechanical pressure on the cells themselves leads to an increased expression of cell-specific proteins and cell viability. In bone, for example, there are several mechanisms by which bone growth is promoted and the activity of fully differentiated cells is increased. Numerous studies showed the upregulation of bone morphogenetic protein 2 (BMP-2) after the exposure of bones to fESWs [47,67,104]. BMP-2 plays a major role in osteoblast differentiation by transforming osteoblast precursor cells into mature osteoblasts that form healthy bones [207]. On the other hand, for proteins, such as RANKL, which, in turn, plays a role in osteoclast differentiation [208], a reduced expression was found after exposure to ESWs [19,63,80]. Furthermore, cavitation induced by ESWs can cause so-called “microcracks”, which is a stimulus for bone remodeling and new bone formation [209]. It was demonstrated in the bones of horses that fESWs can induce new microcracks, and rESWs can extend the length of existing microcracks [102]. When observing the effects of ESWT on the activity of different cell types, an increase in activity in tissue-specific cells, such as fibroblasts [68,124] and osteoblasts [39,83], but, at the same time, a reduced activity of osteoclasts [19], was observed. Together with the reduced RANKL expression, this could indicate a positive effect of ESWT on bone formation, as well as an improvement of diseases affecting the skeletal system, such as osteoporosis. In fact, ESWT shows positive effects in the treatment of these indications [8,210].
The sixth take-home message of this study is that extracorporeal shock wave therapy apparently mimics the effect of capsaicin by reducing substance-P concentration. In pathologies of tendons, muscle injuries and dysfunctions, as well as in osteoarthritis, the inflammatory cycle plays a crucial role, as does nociception for the quality of life of the patients. Substance P is a neuropeptide, which, once released after the activation of the TRPV1 receptor on mainly polymodal C-fibers [211], primarily activates the neurokinin-1 receptor (NK1R) [211,212]. Substance P plays an important role in nociception and neurogenic inflammation [213] through several intracellular pathways [212]. Therefore, in recent years, special attention was paid to capsaicin, a naturally occurring alkaloid that has certain reducing effects on substance-P concentration. Specifically, after application to the peripheral nerve, one of the effects of capsaicin was shown in an activation of the TRPV1 channel, mainly in the terminal endings of nociceptive fibers (especially C fibers), which initially does not lead to a reduction in pain and inflammation as an increase in substance-P concentration is to be expected [211,214]. By releasing substance P from the nerve fibers and simultaneously blocking the axoplasmic transport [215], the terminals are then depleted of their substance-P content [211,214]. However, whether this mechanism is (in addition to reducing inflammation [216]) also responsible for the pain relief with local capsaicin application is currently highly debated [217]. With ESWT, on the other hand, there is evidence that one of the analgesic effects is due to a reduction in the substance-P concentration in the tissue under treatment [191,193], thereby removing substance P from the C fibers. The mechanism behind this is probably a detrimental effect of ESWs on the TRPV1 channel. As with capsaicin, a similar time course of alterations in the amount of substance P in the periosteum was found after exposure of the femur of healthy rabbits to fESWs [191,193]. This may break the inflammatory cycle created by substance-P release, and thus has a different mechanism than medications, such as non-steroidal anti-inflammatory drugs (NSAIDs) that inhibit cyclooxygenase [218], but still helps reduce inflammation. In addition, both substance-P and calcitonin gene-related peptide expressions were demonstrated to be reduced in dorsal root ganglia after the exposure of peripheral tissue to ESWs [184,186,192]. The effect on the local inflammatory circuit is probably additionally enhanced by this. Due to the local application of ESWs, this effect is limited to the treatment region and the affected spinal cord segments, proven at least for substance P [184]. An important result of this is that ESWT does not induce the typical adverse events of treatments with NSAIDs, such as gastrointestinal ulcers and renal damage [218].
The seventh take-home message of this study is that extracorporeal shock wave therapy apparently mimics effects of injection of Botulinum toxin A by destroying endplates in the neuromuscular junction. Botulinum toxin A (BTX-A) injections are nowadays widely used for treating spasticity, which mainly affects individual muscle groups. Examples include spasticity induced by stroke [219], spinal cord injury [220] and infantile cerebral palsy [221]. The central problem in muscle spasticity is constant overexcitation at the neuromuscular endplate. BTX-A effectively prevents the formation of a stable SNARE complex by cleaving one of its associated proteins, SNAP-25. Since the SNARE complex is essential for acetylcholine release, a block of the skeletal cholinergic neuromuscular transmission occurs [222]. As reports of potentially serious side effects of BTX-A injections for treating spasticity continue to emerge [223,224] and long-term effects of this treatment modality remain to be established, the question of new treatment options arises. Extracorporeal shock wave therapy, similar to BTX-A injection, can transiently reduce excitatory transmission at the neuromuscular endplate. In this regard, it was shown in a rat model that the exposure of muscles to rESWs reduced the compound muscle action potential while maintaining the latency [157,168]. The key mechanism of ESWs, in contrast to BTX-A, is most likely the destruction of end plates in neuromuscular junctions, whereby the damage was confined to the postsynaptic membrane [168]. In a recent randomized controlled trial, it was found that BTX-A injection was not superior to rESWT in the treatment of plantar flexor muscle spasticity in patients with cerebral palsy [200].
The eighth take-home message of this study is that extracorporeal shock wave therapy apparently imitates certain mechanisms of action of neural therapy. Neural therapy is a treatment commonly used in Europe for pain relief. Its aim is to normalize the nervous system through targeted injections of local anesthetics [225]. Local anesthetics, such as the commonly used procaine, cause a blockade of the voltage-dependent sodium channels of nerve fibers [226]. This causes a reversible blockade of excitation conduction in nerve fibers, i.e., nociceptive afferents are shut down [226]. ESWT may have a similar principle of action in order to reduce pain conditions. Specifically, it was shown that, after the exposure of the femur to fESWs, a selective destruction and decreased number of unmyelinated nerve fibers in the sciatic nerve of rabbits was induced [183]. C fibers, for example, as part of the nociceptive system, belong to the unmyelinated nerve fibers. Furthermore, ESWs were shown to induce disturbed integrity of myelin sheaths combined with reduced nerve conduction velocities in palmar digital nerves in horses [190], as well as a reduced number of epidermal nerve fibers in the skin [189]. In summary, these results suggest that ESWT can reduce peripheral nerve function and conduction, without affecting the performance of professional athletes [227]. This mechanism may be central to the reduction in pain perception following ESWT, given the possibility that the transmission of nociceptive signals via peripheral nerves is impaired. Furthermore, it cannot be excluded that ESWT influences the conduction ability of sensitive nerves through the activation of gate-control mechanisms in the spinal cord [228]. Compared to neural therapy, a recent study demonstrated that, in patients with myofascial trigger points in the upper trapezius, both the repeated injection of 1% lidocaine and rESWT resulted in reduced pain alongside improved muscle elasticity, pressure pain threshold and neck disability index [229].
The ninth take-home message is that extracorporeal shock wave therapy apparently imitates certain mechanisms of manual therapy treatments. Many manual therapy treatments, such as massages, are aimed at achieving effects, including improved blood circulation, angiogenesis and reduced lymph congestion [230]. These effects were also observed after ESWT. For example, the exposure of skin and muscle tissue to both fESWs and rESWs resulted in a significant increase in the local microcirculation [126,131,173]. A positive effect of ESWT was also described on lymphatic drainage [137], and increased angiogenesis after exposure to ESWs was found in both blood vessels [131,165] and lymph vessels [137]. In addition, ESWT has a stimulating effect on the expression of lubricin in fasciae and tendon sheaths [135]. Lubricin was shown to induce an improvement in tendon gliding in vivo, and the absence of lubricin was demonstrated to significantly limit tendon mobility [231]. Of note, tendon gliding plays a major role in the rehabilitation of tendinopathies and tendon injuries [232]. Furthermore, rESWT was shown to significantly improve immobility-related muscle contractures and muscle fibrosis [156] in a rabbit model. A possible mechanism behind this is the reduced collagen deposition that was observed after treatment. However, it is unclear whether ESWT can also improve fascial fibrosis. As this is an alteration within the collagen fiber layers due to large amounts of undirected collagen material deposition [233,234], ESWT could also have a positive effect here.
The tenth take-home message is that even the most sophisticated research into the effects of exposure of musculoskeletal tissue to extracorporeal shock waves cannot substitute clinical research in order to determine the optimum intensity, treatment frequency and localization of extracorporeal shock wave therapy. Since this study was mainly about the different mechanisms of ESWT, no optimal treatment settings can be determined from the results summarized in Table 1, Table 2 and Table 3. In several studies, certain processes at the cellular level were described at certain points in time, which even contradicted each other in part. For example, while the exposure of cells to ESWs often led to reduced cell viability shortly after exposure, an increase in cell viability was observed in the further course of observation [84]. Therefore, it is reasonable to hypothesize that some biological changes only occur at a certain time, which, however, must be carefully considered in the study protocol and the measurements. In addition, some effects of the exposure of cells and tissue to ESWs were found only at certain energy levels [105,157] and numbers of applied ESWs [42]. Some studies even showed that the exposure of musculoskeletal tissue with ESWs with increasing EFD did not necessarily lead to better outcomes [105,106]. In summary, the only way to further optimize clinical application of ESWT is to perform more and better clinical research on this fascinating treatment modality. It is obvious that the results of basic research may be inspirational in this regard.
This systematic review had three limitations. First, only PubMed and Web of Science were searched. However, considering the fact that, in this review, considerably more studies were considered than in previous reviews on the same topic [13,14,15,16,17], it is reasonable to hypothesize that, in the present investigation, the risk to overlook any relevant study on the effects of exposure of musculoskeletal tissue to extracorporeal shock waves was minimized. Second, no meta-analysis of the presented data was performed. However, as outlined, particularly in the take-home messages 1–3 and 5, this appears to not be possible. Third, this review did not address all the potential indications of ESWT, but was restricted to musculoskeletal tissue. The mechanisms of action of ESWs in the treatment of, e.g., acute and chronic soft tissue wounds (e.g., [235]) or coronary artery disease (e.g., [236]) with ESWT may or may not be the same as discussed in this investigation.

5. Conclusions

The complementary effects of ESWT in the treatment of musculoskeletal pathologies make it an effective form of therapy that can be used alone or in combination with other therapeutic modalities. Not to be underestimated is the possibility of using ESWT as a supportive measure for any myofascial imbalances and functional movement restrictions underlying the pathologies. This is explained by the effects of ESWT on the myofascial units, such as the reduction in muscle tone, the decreased inflammatory activity and the effect on trigger points. Further studies, especially clinical studies, are needed for the future use of ESWT. To date, there is still minimal evidence on the ideal treatment settings, intensity, duration, localization and applied energy to provide the best possible treatment.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/biomedicines10051084/s1. Table S1: Details of the studies listed in Table 1. Table S2: details of the studies listed in Table 2. Table S3: Details of the studies listed in Table 3.

Author Contributions

Conceptualization, T.W., C.S. and L.L.J.J.; methodology, T.W., C.S. and L.L.J.J.; software, T.W., C.S. and L.L.J.J.; validation, T.W., C.S. and L.L.J.J.; formal analysis, T.W., C.S. and L.L.J.J.; investigation, T.W., C.S. and L.L.J.J.; resources, T.W., C.S. and L.L.J.J.; data curation, T.W., C.S. and L.L.J.J.; writing---original draft preparation, T.W., C.S. and L.L.J.J.; writing---review and editing, T.W., C.S. and L.L.J.J.; visualization, T.W., C.S. and L.L.J.J.; supervision, C.S.; project administration, C.S.; funding acquisition, not applicable. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

All relevant data are provided in the text.

Acknowledgments

The authors are infinitely grateful to all those who made it possible for L.L.J.J. to study medicine and perform research on ESWT, despite his extreme disability (tetraplegia from C4). L.L.J.J. has been treated with rESWT because of his spasticity by C.S. and T.W., and has not needed any related medication since then, particularly no injection of BTX-A.

Conflicts of Interest

C.S. served as consultant for Electro Medical Systems (Nyon, Switzerland) (the inventor of rESWT and the manufacturer and distributor of the rESWT device, Swiss DolorClast, as well as the distributor of the fESWT device, Swiss PiezoClast) until December 2017, and received funding from Electro Medical Systems for conducting basic research into rESWT at his lab. However, Electro Medical Systems had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results. No other conflicts of interest are reported.

References

  1. Jocham, D.; Chaussy, C.; Schmiedt, E. Extracorporeal shock wave lithotripsy. Urol. Int. 1986, 41, 357–368. [Google Scholar] [CrossRef] [PubMed]
  2. Sauerbruch, T.; Delius, M.; Paumgartner, G.; Holl, J.; Wess, O.; Weber, W.; Hepp, W.; Brendel, W. Fragmentation of gallstones by extracorporeal shock waves. N. Engl. J. Med. 1986, 314, 818–822. [Google Scholar] [CrossRef] [PubMed]
  3. Sauerbruch, T.; Holl, J.; Sackmann, M.; Werner, R.; Wotzka, R.; Paumgartner, G. Disintegration of a pancreatic duct stone with extracorporeal shock waves in a patient with chronic pancreatitis. Endoscopy 1987, 19, 207–208. [Google Scholar] [CrossRef] [PubMed]
  4. Sauerbruch, T.; Stern, M. Fragmentation of bile duct stones by extracorporeal shock waves: A new approach to biliary calculi after failure of routine endoscopic measures. Gastroenterology 1989, 96, 146–152. [Google Scholar] [CrossRef]
  5. Iro, H.; Nitsche, N.; Schneider, H.T.; Ell, C. Extracorporeal shockwave lithotripsy of salivary gland stones. Lancet 1989, 2, 115. [Google Scholar] [CrossRef]
  6. Graff, J.; Richter, K.D.; Pastor, J. Effect of high energy shock waves on bony tissue. Urol. Res. 1988, 16, 252–258. [Google Scholar]
  7. Haupt, G.; Haupt, A.; Ekkernkamp, A.; Gerety, B.; Chvapil, M. Influence of shock waves on fracture healing. Urology 1992, 39, 529–532. [Google Scholar] [CrossRef]
  8. Kertzman, P.; Csaszar, N.B.M.; Furia, J.P.; Schmitz, C. Radial extracorporeal shock wave therapy is efficient and safe in the treatment of fracture nonunions of superficial bones: A retrospective case series. J. Orthop. Surg. Res. 2017, 12, 164. [Google Scholar] [CrossRef] [Green Version]
  9. Speed, C. A systematic review of shockwave therapies in soft tissue conditions: Focusing on the evidence. Br. J. Sports Med. 2014, 48, 1538–1542. [Google Scholar] [CrossRef]
  10. Schmitz, C.; Csaszar, N.B.; Milz, S.; Schieker, M.; Maffulli, N.; Rompe, J.D.; Furia, J.P. Efficacy and safety of extracorporeal shock wave therapy for orthopedic conditions: A systematic review on studies listed in the PEDro database. Br. Med. Bull. 2015, 116, 115–138. [Google Scholar] [CrossRef] [Green Version]
  11. Reilly, J.M.; Bluman, E.; Tenforde, A.S. Effect of shockwave treatment for management of upper and lower extremity musculoskeletal conditions: A narrative review. PM R 2018, 10, 1385–1403. [Google Scholar] [CrossRef] [PubMed]
  12. Hughes, J.P.; Rees, S.; Kalindjian, S.B.; Philpott, K.L. Principles of early drug discovery. Br. J. Pharmacol. 2011, 162, 1239–1249. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  13. Visco, V.; Vulpiani, M.C.; Torrisi, M.R.; Ferretti, A.; Pavan, A.; Vetrano, M. Experimental studies on the biological effects of extracorporeal shock wave therapy on tendon models. A review of the literature. Muscles Ligaments Tendons J. 2014, 4, 357–361. [Google Scholar] [CrossRef]
  14. Liu, T.; Shindel, A.W.; Lin, G.; Lue, T.F. Cellular signaling pathways modulated by low-intensity extracorporeal shock wave therapy. Int. J. Impot. Res. 2019, 31, 170–176. [Google Scholar] [CrossRef]
  15. Auersperg, V.; Trieb, K. Extracorporeal shock wave therapy: An update. EFORT Open. Rev. 2020, 5, 584–592. [Google Scholar] [CrossRef] [PubMed]
  16. Simplicio, C.L.; Purita, J.; Murrell, W.; Santos, G.S.; Dos Santos, R.G.; Lana, J. Extracorporeal shock wave therapy mechanisms in musculoskeletal regenerative medicine. J. Clin. Orthop. Trauma 2020, 11, S309–S318. [Google Scholar] [CrossRef]
  17. Rola, P.; Wlodarczak, A.; Barycki, M.; Doroszko, A. Use of the shock wave therapy in basic research and clinical applications-from bench to bedsite. Biomedicines 2022, 10, 568. [Google Scholar] [CrossRef]
  18. Li, B.; Wang, R.; Huang, X.; Ou, Y.; Jia, Z.; Lin, S.; Zhang, Y.; Xia, H.; Chen, B. Extracorporeal shock wave therapy promotes osteogenic differentiation in a rabbit osteoporosis model. Front. Endocrinol. 2021, 12, 627718. [Google Scholar] [CrossRef]
  19. Inoue, S.; Hatakeyama, J.; Aoki, H.; Kuroki, H.; Niikura, T.; Oe, K.; Fukui, T.; Kuroda, R.; Akisue, T.; Moriyama, H. Utilization of mechanical stress to treat osteoporosis: The effects of electrical stimulation, radial extracorporeal shock wave, and ultrasound on experimental osteoporosis in ovariectomized rats. Calcif. Tissue Int. 2021, 109, 215–229. [Google Scholar] [CrossRef]
  20. Inoue, S.; Hatakeyama, J.; Aoki, H.; Kuroki, H.; Niikura, T.; Oe, K.; Fukui, T.; Kuroda, R.; Akisue, T.; Moriyama, H. Effects of ultrasound, radial extracorporeal shock waves, and electrical stimulation on rat bone defect healing. Ann. N. Y. Acad. Sci. 2021, 1497, 3–14. [Google Scholar] [CrossRef]
  21. Zhao, Z.; Wang, Y.; Wang, Q.; Liang, J.; Hu, W.; Zhao, S.; Li, P.; Zhu, H.; Li, Z. Radial extracorporeal shockwave promotes subchondral bone stem/progenitor cell self-renewal by activating YAP/TAZ and facilitates cartilage repair in vivo. Stem Cell Res. Ther. 2021, 12, 19. [Google Scholar] [CrossRef] [PubMed]
  22. Kobayashi, M.; Chijimatsu, R.; Yoshikawa, H.; Yoshida, K. Extracorporeal shock wave therapy accelerates endochondral ossification and fracture healing in a rat femur delayed-union model. Biochem. Biophys. Res. Commun. 2020, 530, 632–637. [Google Scholar] [CrossRef] [PubMed]
  23. Alshihri, A.; Niu, W.; Kammerer, P.W.; Al-Askar, M.; Yamashita, A.; Kurisawa, M.; Spector, M. The effects of shock wave stimulation of mesenchymal stem cells on proliferation, migration, and differentiation in an injectable gelatin matrix for osteogenic regeneration. J. Tissue Eng. Regen. Med. 2020, 14, 1630–1640. [Google Scholar] [CrossRef] [PubMed]
  24. Hsu, S.L.; Chou, W.Y.; Hsu, C.C.; Ko, J.Y.; Jhan, S.W.; Wang, C.J.; Lee, M.S.; Hsu, T.C.; Cheng, J.H. Shockwave therapy modulates the expression of BMP2 for prevention of bone and cartilage loss in the lower limbs of postmenopausal osteoporosis rat model. Biomedicines 2020, 8, 614. [Google Scholar] [CrossRef] [PubMed]
  25. Ramesh, S.; Zaman, F.; Madhuri, V.; Savendahl, L. Radial extracorporeal shock wave treatment promotes bone growth and chondrogenesis in cultured fetal rat metatarsal bones. Clin. Orthop. Relat. Res. 2020, 478, 668–678. [Google Scholar] [CrossRef] [PubMed]
  26. Colbath, A.C.; Kisiday, J.D.; Phillips, J.N.; Goodrich, L.R. Can extracorporeal shockwave promote osteogenesis of equine bone marrow-derived mesenchymal stem cells in vitro? Stem Cells Dev. 2020, 29, 110–118. [Google Scholar] [CrossRef]
  27. Hashimoto, S.; Ichinose, T.; Ohsawa, T.; Koibuchi, N.; Chikuda, H. Extracorporeal shockwave therapy accelerates the healing of a meniscal tear in the avascular region in a rat model. Am. J. Sports Med. 2019, 47, 2937–2944. [Google Scholar] [CrossRef]
  28. Senel, E.; Ozkan, E.; Bereket, M.C.; Onger, M.E. The assessment of new bone formation induced by unfocused extracorporeal shock wave therapy applied on pre-surgical phase of distraction osteogenesis. Eur. Oral Res. 2019, 53, 125–131. [Google Scholar] [CrossRef]
  29. Kim, Y.H.; Bang, J.I.; Son, H.J.; Kim, Y.; Kim, J.H.; Bae, H.; Han, S.J.; Yoon, H.J.; Kim, B.S. Protective effects of extracorporeal shockwave on rat chondrocytes and temporomandibular joint osteoarthritis; preclinical evaluation with in vivo 99m Tc-HDP SPECT and ex vivo micro-CT. Osteoarthr. Cartil. 2019, 27, 1692–1701. [Google Scholar] [CrossRef]
  30. Buarque de Gusmao, C.V.; Batista, N.A.; Vidotto Lemes, V.T.; Maia Neto, W.L.; de Faria, L.D.; Alves, J.M.; Belangero, W.D. Effect of low-intensity pulsed ultrasound stimulation, extracorporeal shockwaves and radial pressure waves on Akt, BMP-2, ERK-2, FAK and TGF-β1 during bone healing in rat tibial defects. Ultrasound Med. Biol. 2019, 45, 2140–2161. [Google Scholar] [CrossRef]
  31. Cheng, J.H.; Wang, C.J.; Chou, W.Y.; Hsu, S.L.; Chen, J.H.; Hsu, T.C. Comparison efficacy of ESWT and Wharton’s jelly mesenchymal stem cell in early osteoarthritis of rat knee. Am. J. Transl. Res. 2019, 11, 586–598. [Google Scholar] [PubMed]
  32. Ginini, J.G.; Emodi, O.; Sabo, E.; Maor, G.; Shilo, D.; Rachmiel, A. Effects of timing of extracorporeal shock wave therapy on mandibular distraction osteogenesis: An experimental study in a rat model. J. Oral Maxillofac. Surg. 2019, 77, 629–638. [Google Scholar] [CrossRef] [PubMed]
  33. Ginini, J.G.; Maor, G.; Emodi, O.; Shilo, D.; Gabet, Y.; Aizenbud, D.; Rachmiel, A. Effects of extracorporeal shock wave therapy on distraction osteogenesis in rat mandible. Plast. Reconstr. Surg. 2018, 142, 1501–1509. [Google Scholar] [CrossRef] [PubMed]
  34. Qi, H.; Jin, S.; Yin, C.; Chen, L.; Sun, L.; Liu, Y. Radial extracorporeal shock wave therapy promotes osteochondral regeneration of knee joints in rabbits. Exp. Ther. Med. 2018, 16, 3478–3484. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Koolen, M.K.E.; Kruyt, M.C.; Zadpoor, A.A.; Oner, F.C.; Weinans, H.; van der Jagt, O.P. Optimization of screw fixation in rat bone with extracorporeal shock waves. J. Orthop. Res. 2018, 36, 76–84. [Google Scholar] [CrossRef]
  36. Mackert, G.A.; Schulte, M.; Hirche, C.; Kotsougiani, D.; Vogelpohl, J.; Hoener, B.; Fiebig, T.; Kirschner, S.; Brockmann, M.A.; Lehnhardt, M.; et al. Low-energy extracorporeal shockwave therapy (ESWT) improves metaphyseal fracture healing in an osteoporotic rat model. PLoS ONE 2017, 12, e0189356. [Google Scholar] [CrossRef] [Green Version]
  37. Tan, L.; Zhao, B.; Ge, F.T.; Sun, D.H.; Yu, T. Shockwaves inhibit chondrogenic differentiation of human mesenchymal stem cells in association with adenosine and A2B receptors. Sci. Rep. 2017, 7, 14377. [Google Scholar] [CrossRef]
  38. Hsu, S.L.; Cheng, J.H.; Wang, C.J.; Ko, J.Y.; Hsu, C.H. Extracorporeal shockwave therapy enhances expression of Pdia-3 which is a key factor of the 1alpha,25-dihydroxyvitamin D 3 rapid membrane signaling pathway in treatment of early osteoarthritis of the knee. Int. J. Med. Sci. 2017, 14, 1220–1230. [Google Scholar] [CrossRef] [Green Version]
  39. Yilmaz, V.; Karadas, O.; Dandinoglu, T.; Umay, E.; Cakci, A.; Tan, A.K. Efficacy of extracorporeal shockwave therapy and low-intensity pulsed ultrasound in a rat knee osteoarthritis model: A randomized controlled trial. Eur. J. Rheumatol. 2017, 4, 104–108. [Google Scholar] [CrossRef]
  40. Wang, C.J.; Cheng, J.H.; Huang, C.Y.; Hsu, S.L.; Lee, F.Y.; Yip, H.K. Medial tibial subchondral bone is the key target for extracorporeal shockwave therapy in early osteoarthritis of the knee. Am. J. Transl. Res. 2017, 9, 1720–1731. [Google Scholar]
  41. Chen, Y.; Xu, J.; Huang, Z.; Yu, M.; Zhang, Y.; Chen, H.; Ma, Z.; Liao, H.; Hu, J. An innovative approach for enhancing bone defect healing using PLGA scaffolds seeded with extracorporeal-shock-wave-treated bone marrow mesenchymal stem cells (BMSCs). Sci. Rep. 2017, 7, 44130. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  42. Onger, M.E.; Bereket, C.; Sener, I.; Ozkan, N.; Senel, E.; Polat, A.V. Is it possible to change of the duration of consolidation period in the distraction osteogenesis with the repetition of extracorporeal shock waves? Med. Oral. Patol. Oral. Cir. Bucal 2017, 22, e251–e257. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Wang, C.J.; Cheng, J.H.; Chou, W.Y.; Hsu, S.L.; Chen, J.H.; Huang, C.Y. Changes of articular cartilage and subchondral bone after extracorporeal shockwave therapy in osteoarthritis of the knee. Int. J. Med. Sci. 2017, 14, 213–223. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Lama, A.; Santoro, A.; Corrado, B.; Pirozzi, C.; Paciello, O.; Pagano, T.B.; Russo, S.; Calignano, A.; Mattace Raso, G.; Meli, R. Extracorporeal shock waves alone or combined with raloxifene promote bone formation and suppress resorption in ovariectomized rats. PLoS ONE 2017, 12, e0171276. [Google Scholar]
  45. Catalano, M.G.; Marano, F.; Rinella, L.; de Girolamo, L.; Bosco, O.; Fortunati, N.; Berta, L.; Frairia, R. Extracorporeal shockwaves (ESWs) enhance the osteogenic medium-induced differentiation of adipose-derived stem cells into osteoblast-like cells. J. Tissue Eng. Regen. Med. 2017, 11, 390–399. [Google Scholar] [CrossRef] [Green Version]
  46. Ma, H.Z.; Zhou, D.S.; Li, D.; Zhang, W.; Zeng, B.F. A histomorphometric study of necrotic femoral head in rabbits treated with extracorporeal shock waves. J. Phys. Ther. Sci. 2017, 29, 24–28. [Google Scholar] [CrossRef] [Green Version]
  47. Huang, H.M.; Li, X.L.; Tu, S.Q.; Chen, X.F.; Lu, C.C.; Jiang, L.H. Effects of roughly focused extracorporeal shock waves therapy on the expressions of bone morphogenetic protein-2 and osteoprotegerin in osteoporotic fracture in rats. Chin. Med. J. 2016, 129, 2567–2575. [Google Scholar] [CrossRef]
  48. Notarnicola, A.; Vicenti, G.; Maccagnano, G.; Silvestris, F.; Cafforio, P.; Moretti, B. Extracorporeal shock waves induce osteogenic differentiation of human bone-marrow stromal cells. J. Biol. Regul. Homeost. Agents 2016, 30, 139–144. [Google Scholar]
  49. Zhai, L.; Sun, N.; Zhang, B.; Liu, S.T.; Zhao, Z.; Jin, H.C.; Ma, X.L.; Xing, G.Y. Effects of focused extracorporeal shock waves on bone marrow mesenchymal stem cells in patients with avascular necrosis of the femoral head. Ultrasound Med. Biol. 2016, 42, 753–762. [Google Scholar] [CrossRef]
  50. Dias dos Santos, P.R.; De Medeiros, V.P.; Freire Martins de Moura, J.P.; da Silveira Franciozi, C.E.; Nader, H.B.; Faloppa, F. Effects of shock wave therapy on glycosaminoglycan expression during bone healing. Int. J. Surg. 2015, 24, 120–123. [Google Scholar] [CrossRef]
  51. Wang, C.J.; Huang, C.Y.; Hsu, S.L.; Chen, J.H.; Cheng, J.H. Extracorporeal shockwave therapy in osteoporotic osteoarthritis of the knee in rats: An experiment in animals. Arthritis Res. Ther. 2014, 16, R139. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  52. Muzio, G.; Martinasso, G.; Baino, F.; Frairia, R.; Vitale-Brovarone, C.; Canuto, R.A. Key role of the expression of bone morphogenetic proteins in increasing the osteogenic activity of osteoblast-like cells exposed to shock waves and seeded on bioactive glass-ceramic scaffolds for bone tissue engineering. J. Biomater. Appl. 2014, 29, 728–736. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Oktas, B.; Orhan, Z.; Erbil, B.; Degirmenci, E.; Ustundag, N. Effect of extracorporeal shock wave therapy on fracture healing in rat femural fractures with intact and excised periosteum. Eklem Hastalik. Cerrahisi 2014, 25, 158–162. [Google Scholar] [CrossRef] [PubMed]
  54. Sun, D.; Junger, W.G.; Yuan, C.; Zhang, W.; Bao, Y.; Qin, D.; Wang, C.; Tan, L.; Qi, B.; Zhu, D.; et al. Shockwaves induce osteogenic differentiation of human mesenchymal stem cells through atp release and activation of P2X7 receptors. Stem Cells 2013, 31, 1170–1180. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Suhr, F.; Delhasse, Y.; Bungartz, G.; Schmidt, A.; Pfannkuche, K.; Bloch, W. Cell biological effects of mechanical stimulations generated by focused extracorporeal shock wave applications on cultured human bone marrow stromal cells. Stem Cell Res. 2013, 11, 951–964. [Google Scholar] [CrossRef] [Green Version]
  56. Lyon, R.; Liu, X.C.; Kubin, M.; Schwab, J. Does extracorporeal shock wave therapy enhance healing of osteochondritis dissecans of the rabbit knee?: A pilot study. Clin. Orthop. Relat. Res. 2013, 471, 1159–1165. [Google Scholar] [CrossRef] [Green Version]
  57. Wang, C.J.; Sun, Y.C.; Siu, K.K.; Wu, C.T. Extracorporeal shockwave therapy shows site-specific effects in osteoarthritis of the knee in rats. J. Surg. Res. 2013, 183, 612–619. [Google Scholar] [CrossRef]
  58. Wang, C.J.; Hsu, S.L.; Weng, L.H.; Sun, Y.C.; Wang, F.S. Extracorporeal shockwave therapy shows a number of treatment related chondroprotective effect in osteoarthritis of the knee in rats. BMC Musculoskelet. Disord. 2013, 14, 44. [Google Scholar] [CrossRef] [Green Version]
  59. van der Jagt, O.P.; Waarsing, J.H.; Kops, N.; Schaden, W.; Jahr, H.; Verhaar, J.A.; Weinans, H. Unfocused extracorporeal shock waves induce anabolic effects in osteoporotic rats. J. Orthop. Res. 2013, 31, 768–775. [Google Scholar] [CrossRef] [Green Version]
  60. Oztemur, Z.; Ozturk, H.; Ozyurek, S.; Kaloglu, C.; Golge, U.H.; Bulut, O. The long-term effects of extracorporeal shock waves on the epiphysis of the adolescent rat. J. Orthop. Sci. 2013, 18, 159–164. [Google Scholar] [CrossRef]
  61. Gollwitzer, H.; Gloeck, T.; Roessner, M.; Langer, R.; Horn, C.; Gerdesmeyer, L.; Diehl, P. Radial extracorporeal shock wave therapy (reswt) induces new bone formation in vivo: Results of an animal study in rabbits. Ultrasound Med. Biol. 2013, 39, 126–133. [Google Scholar] [CrossRef] [PubMed]
  62. Altuntas, E.E.; Oztemur, Z.; Ozer, H.; Muderris, S. Effect of extracorporeal shock waves on subcondylar mandibular fractures. J. Craniofac. Surg. 2012, 23, 1645–1648. [Google Scholar] [CrossRef] [PubMed]
  63. Notarnicola, A.; Tamma, R.; Moretti, L.; Fiore, A.; Vicenti, G.; Zallone, A.; Moretti, B. Effects of radial shock waves therapy on osteoblasts activities. Musculoskelet. Surg. 2012, 96, 183–189. [Google Scholar] [CrossRef] [PubMed]
  64. Zhao, Z.; Ji, H.; Jing, R.; Liu, C.; Wang, M.; Zhai, L.; Bai, X.; Xing, G. Extracorporeal shock-wave therapy reduces progression of knee osteoarthritis in rabbits by reducing nitric oxide level and chondrocyte apoptosis. Arch. Orthop. Trauma Surg. 2012, 132, 1547–1553. [Google Scholar] [CrossRef] [PubMed]
  65. Kearney, C.J.; Hsu, H.P.; Spector, M. The use of extracorporeal shock wave-stimulated periosteal cells for orthotopic bone generation. Tissue Eng. Part A 2012, 18, 1500–1508. [Google Scholar] [CrossRef]
  66. Xu, J.K.; Chen, H.J.; Li, X.D.; Huang, Z.L.; Xu, H.; Yang, H.L.; Hu, J. Optimal intensity shock wave promotes the adhesion and migration of rat osteoblasts via integrin beta1-mediated expression of phosphorylated focal adhesion kinase. J. Biol. Chem. 2012, 287, 26200–26212. [Google Scholar] [CrossRef] [Green Version]
  67. Wang, C.J.; Sun, Y.C.; Wong, T.; Hsu, S.L.; Chou, W.Y.; Chang, H.W. Extracorporeal shockwave therapy shows time-dependent chondroprotective effects in osteoarthritis of the knee in rats. J. Surg. Res. 2012, 178, 196–205. [Google Scholar] [CrossRef]
  68. Erturk, C.; Altay, M.A.; Ozardali, I.; Altay, N.; Cece, H.; Isikan, U.E. The effect of extracorporeal shockwaves on cartilage end-plates in rabbits: A preliminary mri and histopathological study. Acta Orthop. Traumatol. Turc. 2012, 46, 449–454. [Google Scholar] [CrossRef]
  69. Wang, C.J.; Weng, L.H.; Ko, J.Y.; Wang, J.W.; Chen, J.M.; Sun, Y.C.; Yang, Y.J. Extracorporeal shockwave shows regression of osteoarthritis of the knee in rats. J. Surg. Res. 2011, 171, 601–608. [Google Scholar] [CrossRef]
  70. van der Jagt, O.P.; Piscaer, T.M.; Schaden, W.; Li, J.; Kops, N.; Jahr, H.; van der Linden, J.C.; Waarsing, J.H.; Verhaar, J.A.; de Jong, M.; et al. Unfocused extracorporeal shock waves induce anabolic effects in rat bone. J. Bone Jt. Surg. Am. 2011, 93, 38–48. [Google Scholar] [CrossRef] [Green Version]
  71. Notarnicola, A.; Tamma, R.; Moretti, L.; Panella, A.; Dell’endice, S.; Zallone, A.; Moretti, B. Effect of shock wave treatment on platelet-rich plasma added to osteoblast cultures. Ultrasound Med. Biol. 2011, 37, 160–168. [Google Scholar] [CrossRef] [PubMed]
  72. Hausdorf, J.; Sievers, B.; Schmitt-Sody, M.; Jansson, V.; Maier, M.; Mayer-Wagner, S. Stimulation of bone growth factor synthesis in human osteoblasts and fibroblasts after extracorporeal shock wave application. Arch. Orthop. Trauma Surg. 2011, 131, 303–309. [Google Scholar] [CrossRef] [PubMed]
  73. Wang, C.J.; Huang, K.E.; Sun, Y.C.; Yang, Y.J.; Ko, J.Y.; Weng, L.H.; Wang, F.S. VEGF modulates angiogenesis and osteogenesis in shockwave-promoted fracture healing in rabbits. J. Surg. Res. 2011, 171, 114–119. [Google Scholar] [CrossRef] [PubMed]
  74. Mayer-Wagner, S.; Ernst, J.; Maier, M.; Chiquet, M.; Joos, H.; Muller, P.E.; Jansson, V.; Sievers, B.; Hausdorf, J. The effect of high-energy extracorporeal shock waves on hyaline cartilage of adult rats in vivo. J. Orthop. Res. 2010, 28, 1050–1056. [Google Scholar] [CrossRef]
  75. Muzio, G.; Verne, E.; Canuto, R.A.; Martinasso, G.; Saracino, S.; Baino, F.; Miola, M.; Berta, L.; Frairia, R.; Vitale-Brovarone, C. Shock waves induce activity of human osteoblast-like cells in bioactive scaffolds. J. Trauma 2010, 68, 1439–1444. [Google Scholar] [CrossRef] [Green Version]
  76. Lai, J.P.; Wang, F.S.; Hung, C.M.; Wang, C.J.; Huang, C.J.; Kuo, Y.R. Extracorporeal shock wave accelerates consolidation in distraction osteogenesis of the rat mandible. J. Trauma 2010, 69, 1252–1258. [Google Scholar] [CrossRef]
  77. Qin, L.; Wang, L.; Wong, M.W.; Wen, C.; Wang, G.; Zhang, G.; Chan, K.M.; Cheung, W.H.; Leung, K.S. Osteogenesis induced by extracorporeal shockwave in treatment of delayed osteotendinous junction healing. J. Orthop. Res. 2010, 28, 70–76. [Google Scholar] [CrossRef]
  78. van der Jagt, O.P.; van der Linden, J.C.; Schaden, W.; van Schie, H.T.; Piscaer, T.M.; Verhaar, J.A.; Weinans, H.; Waarsing, J.H. Unfocused extracorporeal shock wave therapy as potential treatment for osteoporosis. J. Orthop. Res. 2009, 27, 1528–1533. [Google Scholar] [CrossRef]
  79. Iannone, F.; Moretti, B.; Notarnicola, A.; Moretti, L.; Patella, S.; Patella, V.; Lapadula, G. Extracorporeal shock waves increase interleukin-10 expression by human osteoarthritic and healthy osteoblasts in vitro. Clin. Exp. Rheumatol. 2009, 27, 794–799. [Google Scholar]
  80. Tamma, R.; dell’Endice, S.; Notarnicola, A.; Moretti, L.; Patella, S.; Patella, V.; Zallone, A.; Moretti, B. Extracorporeal shock waves stimulate osteoblast activities. Ultrasound Med. Biol. 2009, 35, 2093–2100. [Google Scholar] [CrossRef]
  81. Lee, T.C.; Yang, Y.L.; Chang, N.K.; Lin, T.S.; Lin, W.C.; Liu, Y.S.; Wang, C.J. Biomechanical testing of spinal fusion segments enhanced by extracorporeal shock wave treatment in rabbits. Chang Gung Med. J. 2009, 32, 276–282. [Google Scholar] [PubMed]
  82. Tam, K.F.; Cheung, W.H.; Lee, K.M.; Qin, L.; Leung, K.S. Shockwave exerts osteogenic effect on osteoporotic bone in an ovariectomized goat model. Ultrasound Med. Biol. 2009, 35, 1109–1118. [Google Scholar] [CrossRef] [PubMed]
  83. Hofmann, A.; Ritz, U.; Hessmann, M.H.; Alini, M.; Rommens, P.M.; Rompe, J.D. Extracorporeal shock wave-mediated changes in proliferation, differentiation, and gene expression of human osteoblasts. J. Trauma 2008, 65, 1402–1410. [Google Scholar] [CrossRef] [PubMed]
  84. Tam, K.F.; Cheung, W.H.; Lee, K.M.; Qin, L.; Leung, K.S. Osteogenic effects of low-intensity pulsed ultrasound, extracorporeal shockwaves and their combination-an in vitro comparative study on human periosteal cells. Ultrasound Med. Biol. 2008, 34, 1957–1965. [Google Scholar] [CrossRef]
  85. Lee, T.C.; Huang, H.Y.; Yang, Y.L.; Hung, K.S.; Cheng, C.H.; Lin, W.C.; Wang, C.J. Application of extracorporeal shock wave treatment to enhance spinal fusion: A rabbit experiment. Surg. Neurol. 2008, 70, 129–134. [Google Scholar] [CrossRef]
  86. Wang, C.J.; Wang, F.S.; Yang, K.D. Biological effects of extracorporeal shockwave in bone healing: A study in rabbits. Arch. Orthop. Trauma Surg. 2008, 128, 879–884. [Google Scholar] [CrossRef]
  87. Moretti, B.; Iannone, F.; Notarnicola, A.; Lapadula, G.; Moretti, L.; Patella, V.; Garofalo, R. Extracorporeal shock waves down-regulate the expression of interleukin-10 and tumor necrosis factor-alpha in osteoarthritic chondrocytes. BMC Musculoskelet. Disord. 2008, 9, 16. [Google Scholar] [CrossRef] [Green Version]
  88. Tischer, T.; Milz, S.; Weiler, C.; Pautke, C.; Hausdorf, J.; Schmitz, C.; Maier, M. Dose-dependent new bone formation by extracorporeal shock wave application on the intact femur of rabbits. Eur. Surg. Res. 2008, 41, 44–53. [Google Scholar] [CrossRef] [Green Version]
  89. Ozturk, H.; Bulut, O.; Oztemur, Z.; Kaloglu, C.; Kol, I.O. Effect of high-energy extracorporeal shock waves on the immature epiphysis in a rabbit model. Arch. Orthop. Trauma Surg. 2008, 128, 627–631. [Google Scholar] [CrossRef]
  90. Ma, H.Z.; Zeng, B.F.; Li, X.L. Upregulation of VEGF in subchondral bone of necrotic femoral heads in rabbits with use of extracorporeal shock waves. Calcif. Tissue Int. 2007, 81, 124–131. [Google Scholar] [CrossRef]
  91. Murata, R.; Nakagawa, K.; Ohtori, S.; Ochiai, N.; Arai, M.; Saisu, T.; Sasho, T.; Takahashi, K.; Moriya, H. The effects of radial shock waves on gene transfer in rabbit chondrocytes in vitro. Osteoarthr. Cartil. 2007, 15, 1275–1282. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  92. Benson, B.M.; Byron, C.R.; Pondenis, H.; Stewart, A.A. The effects of radial shock waves on the metabolism of equine cartilage explants in vitro. N. Z. Vet. J. 2007, 55, 40–44. [Google Scholar] [CrossRef] [PubMed]
  93. Martini, L.; Giavaresi, G.; Fini, M.; Borsari, V.; Torricelli, P.; Giardino, R. Early effects of extracorporeal shock wave treatment on osteoblast-like cells: A comparative study between electromagnetic and electrohydraulic devices. J. Trauma 2006, 61, 1198–1206. [Google Scholar] [CrossRef] [PubMed]
  94. Bulut, O.; Eroglu, M.; Ozturk, H.; Tezeren, G.; Bulut, S.; Koptagel, E. Extracorporeal shock wave treatment for defective nonunion of the radius: A rabbit model. J. Orthop. Surg. 2006, 14, 133–137. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Martini, L.; Giavaresi, G.; Fini, M.; Torricelli, P.; Borsari, V.; Giardino, R.; De Pretto, M.; Remondini, D.; Castellani, G.C. Shock wave therapy as an innovative technology in skeletal disorders: Study on transmembrane current in stimulated osteoblast-like cells. Int. J. Artif. Organs 2005, 28, 841–847. [Google Scholar] [CrossRef] [PubMed]
  96. Saisu, T.; Kamegaya, M.; Wada, Y.; Takahashi, K.; Mitsuhashi, S.; Moriya, H.; Maier, M. Acetabular augmentation induced by extracorporeal shock waves in rabbits. J. Pediatr. Orthop. B 2005, 14, 162–167. [Google Scholar] [CrossRef]
  97. Chen, Y.J.; Wurtz, T.; Wang, C.J.; Kuo, Y.R.; Yang, K.D.; Huang, H.C.; Wang, F.S. Recruitment of mesenchymal stem cells and expression of TGF-beta 1 and VEGF in the early stage of shock wave-promoted bone regeneration of segmental defect in rats. J. Orthop. Res. 2004, 22, 526–534. [Google Scholar] [CrossRef]
  98. Saisu, T.; Takahashi, K.; Kamegaya, M.; Mitsuhashi, S.; Wada, Y.; Moriya, H. Effects of extracorporeal shock waves on immature rabbit femurs. J. Pediatr. Orthop. B 2004, 13, 176–183. [Google Scholar]
  99. Chen, Y.J.; Kuo, Y.R.; Yang, K.D.; Wang, C.J.; Sheen Chen, S.M.; Huang, H.C.; Yang, Y.J.; Yi-Chih, S.; Wang, F.S. Activation of extracellular signal-regulated kinase (ERK) and p38 kinase in shock wave-promoted bone formation of segmental defect in rats. Bone 2004, 34, 466–477. [Google Scholar] [CrossRef]
  100. Pauwels, F.E.; McClure, S.R.; Amin, V.; Van Sickle, D.; Evans, R.B. Effects of extracorporeal shock wave therapy and radial pressure wave therapy on elasticity and microstructure of equine cortical bone. Am. J. Vet. Res. 2004, 65, 207–212. [Google Scholar] [CrossRef]
  101. Wang, F.S.; Yang, K.D.; Wang, C.J.; Huang, H.C.; Chio, C.C.; Hsu, T.Y.; Ou, C.Y. Shockwave stimulates oxygen radical-mediated osteogenesis of the mesenchymal cells from human umbilical cord blood. J. Bone Miner. Res. 2004, 19, 973–982. [Google Scholar] [CrossRef] [PubMed]
  102. Da Costa Gomez, T.M.; Radtke, C.L.; Kalscheur, V.L.; Swain, C.A.; Scollay, M.C.; Edwards, R.B.; Santschi, E.M.; Markel, M.D.; Muir, P. Effect of focused and radial extracorporeal shock wave therapy on equine bone microdamage. Vet. Surg. 2004, 33, 49–55. [Google Scholar] [CrossRef] [PubMed]
  103. Takahashi, K.; Yamazaki, M.; Saisu, T.; Nakajima, A.; Shimizu, S.; Mitsuhashi, S.; Moriya, H. Gene expression for extracellular matrix proteins in shockwave-induced osteogenesis in rats. Calcif. Tissue Int. 2004, 74, 187–193. [Google Scholar] [CrossRef] [PubMed]
  104. Chen, Y.J.; Kuo, Y.R.; Yang, K.D.; Wang, C.J.; Huang, H.C.; Wang, F.S. Shock wave application enhances pertussis toxin protein-sensitive bone formation of segmental femoral defect in rats. J. Bone Miner. Res. 2003, 18, 2169–2179. [Google Scholar] [CrossRef]
  105. Martini, L.; Fini, M.; Giavaresi, G.; Torricelli, P.; de Pretto, M.; Rimondini, L.; Giardino, R. Primary osteoblasts response to shock wave therapy using different parameters. Artif. Cells Blood Substit. Immobil. Biotechnol. 2003, 31, 449–466. [Google Scholar] [CrossRef]
  106. Martini, L.; Giavaresi, G.; Fini, M.; Torricelli, P.; de Pretto, M.; Schaden, W.; Giardino, R. Effect of extracorporeal shock wave therapy on osteoblastlike cells. Clin. Orthop. Relat. Res. 2003, 413, 269–280. [Google Scholar] [CrossRef] [PubMed]
  107. Dorotka, R.; Kubista, B.; Schatz, K.D.; Trieb, K. Effects of extracorporeal shock waves on human articular chondrocytes and ovine bone marrow stromal cells in vitro. Arch. Orthop. Trauma Surg. 2003, 123, 345–348. [Google Scholar] [CrossRef]
  108. Wang, F.S.; Yang, K.D.; Kuo, Y.R.; Wang, C.J.; Sheen-Chen, S.M.; Huang, H.C.; Chen, Y.J. Temporal and spatial expression of bone morphogenetic proteins in extracorporeal shock wave-promoted healing of segmental defect. Bone 2003, 32, 387–396. [Google Scholar] [CrossRef]
  109. Maier, M.; Milz, S.; Tischer, T.; Munzing, W.; Manthey, N.; Stabler, A.; Holzknecht, N.; Weiler, C.; Nerlich, A.; Refior, H.J.; et al. Influence of extracorporeal shock-wave application on normal bone in an animal model in vivo. Scintigraphy, MRI and histopathology. J. Bone Jt. Surg. Br. 2002, 84, 592–599. [Google Scholar] [CrossRef]
  110. Wang, F.S.; Yang, K.D.; Chen, R.F.; Wang, C.J.; Sheen-Chen, S.M. Extracorporeal shock wave promotes growth and differentiation of bone-marrow stromal cells towards osteoprogenitors associated with induction of TGF-beta1. J. Bone Jt. Surg. Br. 2002, 84, 457–461. [Google Scholar] [CrossRef]
  111. Wang, F.S.; Wang, C.J.; Huang, H.J.; Chung, H.; Chen, R.F.; Yang, K.D. Physical shock wave mediates membrane hyperpolarization and Ras activation for osteogenesis in human bone marrow stromal cells. Biochem. Biophys. Res. Commun. 2001, 287, 648–655. [Google Scholar] [CrossRef] [PubMed]
  112. Wang, C.J.; Huang, H.Y.; Chen, H.H.; Pai, C.H.; Yang, K.D. Effect of shock wave therapy on acute fractures of the tibia: A study in a dog model. Clin. Orthop. Relat. Res. 2001, 387, 112–118. [Google Scholar] [CrossRef] [PubMed]
  113. Vaterlein, N.; Lussenhop, S.; Hahn, M.; Delling, G.; Meiss, A.L. The effect of extracorporeal shock waves on joint cartilage--an in vivo study in rabbits. Arch. Orthop. Trauma Surg. 2000, 120, 403–406. [Google Scholar] [CrossRef] [PubMed]
  114. Peters, N.; Dahmen, G.; Schmidt, W.; Stein, F. Über die Auswirkungen von extrakorporalen Ultraschall-Stossenwellen auf weitentwickelte Embryonen des Knochenfisches Oryzias latipes [Effects of extracorporeal ultrasound shockwaves on the relatively mature embryos of the teleost oryzias latipes]. Ultraschall Med. 1998, 19, 52–58. (In German) [Google Scholar] [CrossRef] [PubMed]
  115. Augat, P.; Claes, L.; Suger, G. In vivo effect of shock-waves on the healing of fractured bone. Clin. Biomech. 1995, 10, 374–378. [Google Scholar] [CrossRef]
  116. Forriol, F.; Solchaga, L.; Moreno, J.L.; Canadell, J. The effect of shockwaves on mature and healing cortical bone. Int. Orthop. 1994, 18, 325–329. [Google Scholar] [CrossRef] [Green Version]
  117. Haberal, B.; Simsek, E.K.; Akpinar, K.; Turkbey Simsek, D.; Sahinturk, F. Impact of radial extracorporeal shock wave therapy in post-laminectomy epidural fibrosis in a rat model. Jt. Dis. Relat. Surg. 2021, 32, 162–169. [Google Scholar] [CrossRef]
  118. Heimes, D.; Wiesmann, N.; Eckrich, J.; Brieger, J.; Mattyasovszky, S.; Proff, P.; Weber, M.; Deschner, J.; Al-Nawas, B.; Kammerer, P.W. In vivo modulation of angiogenesis and immune response on a collagen matrix via extracorporeal shockwaves. Int. J. Mol. Sci. 2020, 21, 7574. [Google Scholar] [CrossRef]
  119. Lu, C.C.; Chou, S.H.; Shen, P.C.; Chou, P.H.; Ho, M.L.; Tien, Y.C. Extracorporeal shock wave promotes activation of anterior cruciate ligament remnant cells and their paracrine regulation of bone marrow stromal cells’ proliferation, migration, collagen synthesis, and differentiation. Bone Jt. Res. 2020, 9, 458–468. [Google Scholar] [CrossRef]
  120. Basoli, V.; Chaudary, S.; Cruciani, S.; Santaniello, S.; Balzano, F.; Ventura, C.; Redl, H.; Dungel, P.; Maioli, M. Mechanical stimulation of fibroblasts by extracorporeal shock waves: Modulation of cell activation and proliferation through a transient proinflammatory milieu. Cell Transplant. 2020, 29, 963689720916175. [Google Scholar] [CrossRef] [Green Version]
  121. Schnurrer-Luke-Vrbanic, T.; Avancini-Dobrovic, V.; Sosa, I.; Cvijanovic, O.; Bobinac, D. VEGF-A expression in soft tissues repaired by shockwave therapy: Differences between modalities. J. Biol. Regul. Homeost. Agents 2018, 32, 583–588. [Google Scholar] [PubMed]
  122. Cui, H.S.; Hong, A.R.; Kim, J.B.; Yu, J.H.; Cho, Y.S.; Joo, S.Y.; Seo, C.H. Extracorporeal shock wave therapy alters the expression of fibrosis-related molecules in fibroblast derived from human hypertrophic scar. Int. J. Mol. Sci. 2018, 19, 124. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  123. Cai, Z.; Falkensammer, F.; Andrukhov, O.; Chen, J.; Mittermayr, R.; Rausch-Fan, X. Effects of shock waves on expression of IL-6, IL-8, MCP-1, and TNF-alpha expression by human periodontal ligament fibroblasts: An in vitro study. Med. Sci. Monit. 2016, 22, 914–921. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  124. Hochstrasser, T.; Frank, H.G.; Schmitz, C. Dose-dependent and cell type-specific cell death and proliferation following in vitro exposure to radial extracorporeal shock waves. Sci. Rep. 2016, 6, 30637. [Google Scholar] [CrossRef] [Green Version]
  125. Leone, L.; Raffa, S.; Vetrano, M.; Ranieri, D.; Malisan, F.; Scrofani, C.; Vulpiani, M.C.; Ferretti, A.; Torrisi, M.R.; Visco, V. Extracorporeal shock wave treatment (ESWT) enhances the in vitro-induced differentiation of human tendon-derived stem/progenitor cells (hTSPCs). Oncotarget 2016, 7, 6410–6423. [Google Scholar] [CrossRef] [Green Version]
  126. Kisch, T.; Sorg, H.; Forstmeier, V.; Knobloch, K.; Liodaki, E.; Stang, F.; Mailander, P.; Kramer, R. Remote effects of extracorporeal shock wave therapy on cutaneous microcirculation. J. Tissue Viability 2015, 24, 140–145. [Google Scholar] [CrossRef]
  127. Waugh, C.M.; Morrissey, D.; Jones, E.; Riley, G.P.; Langberg, H.; Screen, H.R. In vivo biological response to extracorporeal shockwave therapy in human tendinopathy. Eur. Cell Mater. 2015, 29, 268–280. [Google Scholar] [CrossRef]
  128. de Girolamo, L.; Stanco, D.; Galliera, E.; Vigano, M.; Lovati, A.B.; Marazzi, M.G.; Romeo, P.; Sansone, V. Soft-focused extracorporeal shock waves increase the expression of tendon-specific markers and the release of anti-inflammatory cytokines in an adherent culture model of primary human tendon cells. Ultrasound. Med. Biol. 2014, 40, 1204–1215. [Google Scholar] [CrossRef]
  129. Chow, D.H.; Suen, P.K.; Huang, L.; Cheung, W.H.; Leung, K.S.; Ng, C.; Shi, S.Q.; Wong, M.W.; Qin, L. Extracorporeal shockwave enhanced regeneration of fibrocartilage in a delayed tendon-bone insertion repair model. J. Orthop. Res. 2014, 32, 507–514. [Google Scholar] [CrossRef] [Green Version]
  130. Cinar, B.M.; Circi, E.; Balcik, C.; Guven, G.; Akpinar, S.; Derincek, A. The effects of extracorporeal shock waves on carrageenan-induced achilles tendinitis in rats: A biomechanical and histological analysis. Acta Orthop. Traumatol. Turc. 2013, 47, 266–272. [Google Scholar] [CrossRef]
  131. Contaldo, C.; Hogger, D.C.; Khorrami Borozadi, M.; Stotz, M.; Platz, U.; Forster, N.; Lindenblatt, N.; Giovanoli, P. Radial pressure waves mediate apoptosis and functional angiogenesis during wound repair in apoe deficient mice. Microvasc. Res. 2012, 84, 24–33. [Google Scholar] [CrossRef] [PubMed]
  132. Chow, D.H.; Suen, P.K.; Fu, L.H.; Cheung, W.H.; Leung, K.S.; Wong, M.W.; Qin, L. Extracorporeal shockwave therapy for treatment of delayed tendon-bone insertion healing in a rabbit model: A dose-response study. Am. J. Sports Med. 2012, 40, 2862–2871. [Google Scholar] [CrossRef] [PubMed]
  133. Yoo, S.D.; Choi, S.; Lee, G.J.; Chon, J.; Jeong, Y.S.; Park, H.K.; Kim, H.S. Effects of extracorporeal shockwave therapy on nanostructural and biomechanical responses in the collagenase-induced achilles tendinitis animal model. Lasers Med. Sci. 2012, 27, 1195–1204. [Google Scholar] [CrossRef] [PubMed]
  134. Leone, L.; Vetrano, M.; Ranieri, D.; Raffa, S.; Vulpiani, M.C.; Ferretti, A.; Torrisi, M.R.; Visco, V. Extracorporeal shock wave treatment (ESWT) improves in vitro functional activities of ruptured human tendon-derived tenocytes. PLoS ONE 2012, 7, e49759. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  135. Zhang, D.; Kearney, C.J.; Cheriyan, T.; Schmid, T.M.; Spector, M. Extracorporeal shockwave-induced expression of lubricin in tendons and septa. Cell Tissue Res. 2011, 346, 255–262. [Google Scholar] [CrossRef] [PubMed]
  136. Penteado, F.T.; Faloppa, F.; Giusti, G.; Moraes, V.Y.; Belloti, J.C.; Santos, J.B. High-energy extracorporeal shockwave therapy in a patellar tendon animal model: A vascularization focused study. Clinics 2011, 66, 1611–1614. [Google Scholar] [CrossRef] [Green Version]
  137. Kubo, M.; Li, T.S.; Kamota, T.; Ohshima, M.; Shirasawa, B.; Hamano, K. Extracorporeal shock wave therapy ameliorates secondary lymphedema by promoting lymphangiogenesis. J. Vasc. Surg. 2010, 52, 429–434. [Google Scholar] [CrossRef]
  138. Sugioka, K.; Nakagawa, K.; Murata, R.; Ochiai, N.; Sasho, T.; Arai, M.; Tsuruoka, H.; Ohtori, S.; Saisu, T.; Gemba, T.; et al. Radial shock waves effectively introduced NF-kappa b decoy into rat achilles tendon cells in vitro. J. Orthop. Res. 2010, 28, 1078–1083. [Google Scholar] [CrossRef]
  139. Berta, L.; Fazzari, A.; Ficco, A.M.; Enrica, P.M.; Catalano, M.G.; Frairia, R. Extracorporeal shock waves enhance normal fibroblast proliferation in vitro and activate mRNA expression for TGF-beta1 and for collagen types I and III. Acta Orthop. 2009, 80, 612–617. [Google Scholar] [CrossRef]
  140. Bosch, G.; de Mos, M.; van Binsbergen, R.; van Schie, H.T.; van de Lest, C.H.; van Weeren, P.R. The effect of focused extracorporeal shock wave therapy on collagen matrix and gene expression in normal tendons and ligaments. Equine Vet. J. 2009, 41, 335–341. [Google Scholar] [CrossRef]
  141. Han, S.H.; Lee, J.W.; Guyton, G.P.; Parks, B.G.; Courneya, J.P.; Schon, L.C.J. Leonard Goldner award 2008. Effect of extracorporeal shock wave therapy on cultured tenocytes. Foot Ankle Int. 2009, 30, 93–98. [Google Scholar] [CrossRef] [PubMed]
  142. Byron, C.; Stewart, A.; Benson, B.; Tennent-Brown, B.; Foreman, J. Effects of radial extracorporeal shock wave therapy on radiographic and scintigraphic outcomes in horses with palmar heel pain. Vet. Comp. Orthop. Traumatol. 2009, 22, 113–118. [Google Scholar] [PubMed]
  143. Chao, Y.H.; Tsuang, Y.H.; Sun, J.S.; Chen, L.T.; Chiang, Y.F.; Wang, C.C.; Chen, M.H. Effects of shock waves on tenocyte proliferation and extracellular matrix metabolism. Ultrasound Med. Biol. 2008, 34, 841–852. [Google Scholar] [CrossRef] [PubMed]
  144. Wang, L.; Qin, L.; Lu, H.B.; Cheung, W.H.; Yang, H.; Wong, W.N.; Chan, K.M.; Leung, K.S. Extracorporeal shock wave therapy in treatment of delayed bone-tendon healing. Am. J. Sports Med. 2008, 36, 340–347. [Google Scholar] [CrossRef] [PubMed]
  145. Bosch, G.; Lin, Y.L.; van Schie, H.T.; van De Lest, C.H.; Barneveld, A.; van Weeren, P.R. Effect of extracorporeal shock wave therapy on the biochemical composition and metabolic activity of tenocytes in normal tendinous structures in ponies. Equine Vet. J. 2007, 39, 226–231. [Google Scholar] [CrossRef] [PubMed]
  146. Kersh, K.D.; McClure, S.R.; Van Sickle, D.; Evans, R.B. The evaluation of extracorporeal shock wave therapy on collagenase induced superficial digital flexor tendonitis. Vet. Comp. Orthop. Traumatol. 2006, 19, 99–105. [Google Scholar] [CrossRef] [PubMed]
  147. Wang, C.J.; Wang, F.S.; Yang, K.D.; Weng, L.H.; Sun, Y.C.; Yang, Y.J. The effect of shock wave treatment at the tendon-bone interface-an histomorphological and biomechanical study in rabbits. J. Orthop. Res. 2005, 23, 274–280. [Google Scholar] [CrossRef]
  148. Chen, Y.J.; Wang, C.J.; Yang, K.D.; Kuo, Y.R.; Huang, H.C.; Huang, Y.T.; Sun, Y.C.; Wang, F.S. Extracorporeal shock waves promote healing of collagenase-induced achilles tendinitis and increase TGF-beta1 and IGF-I expression. J. Orthop. Res. 2004, 22, 854–861. [Google Scholar] [CrossRef]
  149. Orhan, Z.; Ozturan, K.; Guven, A.; Cam, K. The effect of extracorporeal shock waves on a rat model of injury to tendo achillis. A histological and biomechanical study. J. Bone Jt. Surg. Br. 2004, 86, 613–618. [Google Scholar] [CrossRef] [Green Version]
  150. Hsu, R.W.W.; Hsu, W.H.; Tai, C.L.; Lee, K.F. Effect of shock-wave therapy on patellar tendinopathy in a rabbit model. J. Orthop. Res. 2004, 22, 221–227. [Google Scholar] [CrossRef]
  151. Orhan, Z.; Cam, K.; Alper, M.; Ozturan, K. The effects of extracorporeal shock waves on the rat achilles tendon: Is there a critical dose for tissue injury? Arch. Orthop. Trauma Surg. 2004, 124, 631–635. [Google Scholar] [CrossRef] [PubMed]
  152. Wang, C.-J.; Wang, F.-S.; Yang, K.D.; Weng, L.-H.; Hsu, C.-C.; Huang, C.-S.; Yang, L.-C. Shock wave therapy induces neovascularization at the tendon–bone junction. A study in rabbits. J. Orthop. Res. 2003, 21, 984–989. [Google Scholar] [CrossRef]
  153. Maier, M.; Tischer, T.; Milz, S.; Weiler, C.; Nerlich, A.; Pellengahr, C.; Schmitz, C.; Refior, H.J. Dose-related effects of extracorporeal shock waves on rabbit quadriceps tendon integrity. Arch. Orthop. Trauma Surg. 2002, 122, 436–441. [Google Scholar] [CrossRef] [PubMed]
  154. Wang, C.-J.; Huang, H.-Y.; Pai, C.-H. Shock wave-enhanced neovascularization at the tendon-bone junction: An experiment in dogs. J. Foot Ankle Surg. 2002, 41, 16–22. [Google Scholar] [CrossRef]
  155. Johannes, E.J.; Kaulesar Sukul, D.M.; Bijma, A.M.; Mulder, P.G. Effects of high-energy shockwaves on normal human fibroblasts in suspension. J. Surg. Res. 1994, 57, 677–681. [Google Scholar] [CrossRef]
  156. Huang, P.P.; Zhang, Q.B.; Zhou, Y.; Liu, A.Y.; Wang, F.; Xu, Q.Y.; Yang, F. Effect of radial extracorporeal shock wave combined with ultrashort wave diathermy on fibrosis and contracture of muscle. Am. J. Phys. Med. Rehabil. 2021, 100, 643–650. [Google Scholar] [CrossRef]
  157. Kenmoku, T.; Iwakura, N.; Ochiai, N.; Saisu, T.; Ohtori, S.; Takahashi, K.; Nakazawa, T.; Fukuda, M.; Takaso, M. Influence of different energy patterns on efficacy of radial shock wave therapy. J. Orthop. Sci. 2021, 26, 698–703. [Google Scholar] [CrossRef]
  158. Park, H.J.; Hong, J.; Piao, Y.; Shin, H.J.; Lee, S.J.; Rhyu, I.J.; Yi, M.H.; Kim, J.; Kim, D.W.; Beom, J. Extracorporeal shockwave therapy enhances peripheral nerve remyelination and gait function in a crush model. Adv. Clin. Exp. Med. 2020, 29, 819–824. [Google Scholar] [CrossRef]
  159. Matsuda, M.; Kanno, H.; Sugaya, T.; Yamaya, S.; Yahata, K.; Handa, K.; Shindo, T.; Shimokawa, H.; Ozawa, H.; Itoi, E. Low-energy extracorporeal shock wave therapy promotes BDNF expression and improves functional recovery after spinal cord injury in rats. Exp. Neurol. 2020, 328, 113251. [Google Scholar] [CrossRef]
  160. Langendorf, E.K.; Klein, A.; Drees, P.; Rommens, P.M.; Mattyasovszky, S.G.; Ritz, U. Exposure to radial extracorporeal shockwaves induces muscle regeneration after muscle injury in a surgical rat model. J. Orthop. Res. 2020, 38, 1386–1397. [Google Scholar] [CrossRef]
  161. Sagir, D.; Bereket, C.; Onger, M.E.; Bakhit, N.; Keskin, M.; Ozkan, E. Efficacy of extracorporeal shockwaves therapy on peripheral nerve regeneration. J. Craniofac. Surg. 2019, 30, 2635–2639. [Google Scholar] [CrossRef] [PubMed]
  162. Feichtinger, X.; Monforte, X.; Keibl, C.; Hercher, D.; Schanda, J.; Teuschl, A.H.; Muschitz, C.; Redl, H.; Fialka, C.; Mittermayr, R. Substantial biomechanical improvement by extracorporeal shockwave therapy after surgical repair of rodent chronic rotator cuff tears. Am. J. Sports Med. 2019, 47, 2158–2166. [Google Scholar] [CrossRef] [PubMed]
  163. Yang, C.H.; Yip, H.K.; Chen, H.F.; Yin, T.C.; Chiang, J.Y.; Sung, P.H.; Lin, K.C.; Tsou, Y.H.; Chen, Y.L.; Li, Y.C.; et al. Long-term therapeutic effects of extracorporeal shock wave-assisted melatonin therapy on mononeuropathic pain in rats. Neurochem. Res. 2019, 44, 796–810. [Google Scholar] [CrossRef] [PubMed]
  164. Mattyasovszky, S.G.; Langendorf, E.K.; Ritz, U.; Schmitz, C.; Schmidtmann, I.; Nowak, T.E.; Wagner, D.; Hofmann, A.; Rommens, P.M.; Drees, P. Exposure to radial extracorporeal shock waves modulates viability and gene expression of human skeletal muscle cells: A controlled in vitro study. J. Orthop. Surg. Res. 2018, 13, 75. [Google Scholar] [CrossRef] [Green Version]
  165. Yin, T.C.; Wu, R.W.; Sheu, J.J.; Sung, P.H.; Chen, K.H.; Chiang, J.Y.; Hsueh, S.K.; Chung, W.J.; Lin, P.Y.; Hsu, S.L.; et al. Combined therapy with extracorporeal shock wave and adipose-derived mesenchymal stem cells remarkably improved acute ischemia-reperfusion injury of quadriceps muscle. Oxid. Med. Cell. Longev. 2018, 2018, 6012636. [Google Scholar] [CrossRef] [PubMed]
  166. Shin, D.C.; Ha, K.Y.; Kim, Y.H.; Kim, J.W.; Cho, Y.K.; Kim, S.I. Induction of endogenous neural stem cells by extracorporeal shock waves after spinal cord injury. Spine 2018, 43, E200–E207. [Google Scholar] [CrossRef] [PubMed]
  167. Luh, J.J.; Huang, W.T.; Lin, K.H.; Huang, Y.Y.; Kuo, P.L.; Chen, W.S. Effects of extracorporeal shock wave-mediated transdermal local anesthetic drug delivery on rat caudal nerves. Ultrasound. Med. Biol. 2018, 44, 214–222. [Google Scholar] [CrossRef]
  168. Kenmoku, T.; Nemoto, N.; Iwakura, N.; Ochiai, N.; Uchida, K.; Saisu, T.; Ohtori, S.; Nakagawa, K.; Sasho, T.; Takaso, M. Extracorporeal shock wave treatment can selectively destroy end plates in neuromuscular junctions. Muscle Nerve 2018, 57, 466–472. [Google Scholar] [CrossRef]
  169. Chen, K.H.; Yang, C.H.; Wallace, C.G.; Lin, C.R.; Liu, C.K.; Yin, T.C.; Huang, T.H.; Chen, Y.L.; Sun, C.K.; Yip, H.K. Combination therapy with extracorporeal shock wave and melatonin markedly attenuated neuropathic pain in rat. Am. J. Transl. Res. 2017, 9, 4593–4606. [Google Scholar]
  170. Yahata, K.; Kanno, H.; Ozawa, H.; Yamaya, S.; Tateda, S.; Ito, K.; Shimokawa, H.; Itoi, E. Low-energy extracorporeal shock wave therapy for promotion of vascular endothelial growth factor expression and angiogenesis and improvement of locomotor and sensory functions after spinal cord injury. J. Neurosurg. Spine 2016, 25, 745–755. [Google Scholar] [CrossRef] [Green Version]
  171. Schuh, C.M.; Hercher, D.; Stainer, M.; Hopf, R.; Teuschl, A.H.; Schmidhammer, R.; Redl, H. Extracorporeal shockwave treatment: A novel tool to improve schwann cell isolation and culture. Cytotherapy 2016, 18, 760–770. [Google Scholar] [CrossRef] [PubMed]
  172. Lee, J.H. Knee joint angle of intracerebral hemorrhage-induced rats after extracorporeal shock wave therapy. J. Phys. Ther. Sci. 2016, 28, 3122–3124. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  173. Kisch, T.; Wuerfel, W.; Forstmeier, V.; Liodaki, E.; Stang, F.H.; Knobloch, K.; Mailaender, P.; Kraemer, R. Repetitive shock wave therapy improves muscular microcirculation. J. Surg. Res. 2016, 201, 440–445. [Google Scholar] [CrossRef] [PubMed]
  174. Lee, J.H.; Kim, S.G. Effects of extracorporeal shock wave therapy on functional recovery and neurotrophin-3 expression in the spinal cord after crushed sciatic nerve injury in rats. Ultrasound Med. Biol. 2015, 41, 790–796. [Google Scholar] [CrossRef]
  175. Yamaya, S.; Ozawa, H.; Kanno, H.; Kishimoto, K.N.; Sekiguchi, A.; Tateda, S.; Yahata, K.; Ito, K.; Shimokawa, H.; Itoi, E. Low-energy extracorporeal shock wave therapy promotes vascular endothelial growth factor expression and improves locomotor recovery after spinal cord injury. J. Neurosurg. 2014, 121, 1514–1525. [Google Scholar] [CrossRef] [Green Version]
  176. Fu, M.; Cheng, H.; Li, D.; Yu, X.; Ji, N.; Luo, F. Radial shock wave therapy in the treatment of chronic constriction injury model in rats: A preliminary study. Chin. Med. J. 2014, 127, 830–834. [Google Scholar]
  177. Ishikawa, T.; Miyagi, M.; Yamashita, M.; Kamoda, H.; Eguchi, Y.; Arai, G.; Suzuki, M.; Sakuma, Y.; Oikawa, Y.; Orita, S.; et al. In-vivo transfection of the proopiomelanocortin gene, precursor of endogenous endorphin, by use of radial shock waves alleviates neuropathic pain. J. Orthop. Sci. 2013, 18, 636–645. [Google Scholar] [CrossRef]
  178. Mense, S.; Hoheisel, U. Shock wave treatment improves nerve regeneration in the rat. Muscle Nerve 2013, 47, 702–710. [Google Scholar] [CrossRef]
  179. Hausner, T.; Pajer, K.; Halat, G.; Hopf, R.; Schmidhammer, R.; Redl, H.; Nogradi, A. Improved rate of peripheral nerve regeneration induced by extracorporeal shock wave treatment in the rat. Exp. Neurol. 2012, 236, 363–370. [Google Scholar] [CrossRef] [Green Version]
  180. Kenmoku, T.; Ochiai, N.; Ohtori, S.; Saisu, T.; Sasho, T.; Nakagawa, K.; Iwakura, N.; Miyagi, M.; Ishikawa, T.; Tatsuoka, H.; et al. Degeneration and recovery of the neuromuscular junction after application of extracorporeal shock wave therapy. J. Orthop. Res. 2012, 30, 1660–1665. [Google Scholar] [CrossRef]
  181. Yamashita, M.; Yamauchi, K.; Suzuki, M.; Eguchi, Y.; Orita, S.; Endo, M.; Yamashita, T.; Takahashi, K.; Ohtori, S. Transfection of rat cells with proopiomeranocortin gene, precursor of endogenous endorphin, using radial shock waves suppresses inflammatory pain. Spine 2009, 34, 2270–2277. [Google Scholar] [CrossRef] [PubMed]
  182. Wu, Y.H.; Liang, H.W.; Chen, W.S.; Lai, J.S.; Luh, J.J.; Chong, F.C. Electrophysiological and functional effects of shock waves on the sciatic nerve of rats. Ultrasound Med. Biol. 2008, 34, 1688–1696. [Google Scholar] [CrossRef] [PubMed]
  183. Hausdorf, J.; Lemmens, M.A.; Heck, K.D.; Grolms, N.; Korr, H.; Kertschanska, S.; Steinbusch, H.W.; Schmitz, C.; Maier, M. Selective loss of unmyelinated nerve fibers after extracorporeal shockwave application to the musculoskeletal system. Neuroscience 2008, 155, 138–144. [Google Scholar] [CrossRef] [PubMed]
  184. Hausdorf, J.; Lemmens, M.A.; Kaplan, S.; Marangoz, C.; Milz, S.; Odaci, E.; Korr, H.; Schmitz, C.; Maier, M. Extracorporeal shockwave application to the distal femur of rabbits diminishes the number of neurons immunoreactive for substance P in dorsal root ganglia L5. Brain Res. 2008, 1207, 96–101. [Google Scholar] [CrossRef] [PubMed]
  185. Lee, T.C.; Huang, H.Y.; Yang, Y.L.; Hung, K.S.; Cheng, C.H.; Chang, N.K.; Chung, Y.H.; Hu, M.S.; Wang, C.J. Vulnerability of the spinal cord to injury from extracorporeal shock waves in rabbits. J. Clin. Neurosci. 2007, 14, 873–878. [Google Scholar] [CrossRef] [PubMed]
  186. Ochiai, N.; Ohtori, S.; Sasho, T.; Nakagawa, K.; Takahashi, K.; Takahashi, N.; Murata, R.; Takahashi, K.; Moriya, H.; Wada, Y.; et al. Extracorporeal shock wave therapy improves motor dysfunction and pain originating from knee osteoarthritis in rats. Osteoarthr. Cartil. 2007, 15, 1093–1096. [Google Scholar] [CrossRef] [Green Version]
  187. Wu, Y.H.; Lun, J.J.; Chen, W.S.; Chong, F.C. The electrophysiological and functional effect of shock wave on peripheral nerves. In Proceedings of the Annual International Conference of the IEEE Engineering in Medicine and Biology Society, Lyon, France, 22–26 August 2007; Volume 2007, pp. 2369–2372. [Google Scholar]
  188. Murata, R.; Ohtori, S.; Ochiai, N.; Takahashi, N.; Saisu, T.; Moriya, H.; Takahashi, K.; Wada, Y. Extracorporeal shockwaves induce the expression of ATF3 and GAP-43 in rat dorsal root ganglion neurons. Auton. Neurosci. 2006, 128, 96–100. [Google Scholar] [CrossRef]
  189. Takahashi, N.; Ohtori, S.; Saisu, T.; Moriya, H.; Wada, Y. Second application of low-energy shock waves has a cumulative effect on free nerve endings. Clin. Orthop. Relat. Res. 2006, 443, 315–319. [Google Scholar] [CrossRef]
  190. Bolt, D.M.; Burba, D.J.; Hubert, J.D.; Strain, G.M.; Hosgood, G.L.; Henk, W.G.; Cho, D.Y. Determination of functional and morphologic changes in palmar digital nerves after nonfocused extracorporeal shock wave treatment in horses. Am. J. Vet. Res. 2004, 65, 1714–1718. [Google Scholar] [CrossRef]
  191. Hausdorf, J.; Schmitz, C.; Averbeck, B.; Maier, M. Molekulare Grundlagen zur schmerzvermittelnden Wirkung extrakorporaler Stosswellen [Molecular basis for pain mediating properties of extracorporeal shock waves]. Schmerz 2004, 18, 492–497. (In German) [Google Scholar] [CrossRef]
  192. Takahashi, N.; Wada, Y.; Ohtori, S.; Saisu, T.; Moriya, H. Application of shock waves to rat skin decreases calcitonin gene-related peptide immunoreactivity in dorsal root ganglion neurons. Auton. Neurosci. 2003, 107, 81–84. [Google Scholar] [CrossRef]
  193. Maier, M.; Averbeck, B.; Milz, S.; Refior, H.J.; Schmitz, C. Substance P and prostaglandin E2 release after shock wave application to the rabbit femur. Clin. Orthop. Relat. Res. 2003, 406, 237–245. [Google Scholar] [CrossRef]
  194. Haake, M.; Thon, A.; Bette, M. Unchanged c-Fos expression after extracorporeal shock wave therapy: An experimental investigation in rats. Arch. Orthop. Trauma Surg. 2002, 122, 518–521. [Google Scholar] [CrossRef] [PubMed]
  195. Ohtori, S.; Inoue, G.; Mannoji, C.; Saisu, T.; Takahashi, K.; Mitsuhashi, S.; Wada, Y.; Takahashi, K.; Yamagata, M.; Moriya, H. Shock wave application to rat skin induces degeneration and reinnervation of sensory nerve fibres. Neurosci. Lett. 2001, 315, 57–60. [Google Scholar] [CrossRef]
  196. Haake, M.; Thon, A.; Bette, M. Absence of spinal response to extracorporeal shock waves on the endogenous opioid systems in the rat. Ultrasound Med. Biol. 2001, 27, 279–284. [Google Scholar] [CrossRef]
  197. Rompe, J.D.; Bohl, J.; Riehle, H.M.; Schwitalle, M.; Krischek, O. Überprüfung der Läsionsgefahr des N. ischiadicus des Kaninchens durch die Applikation niedrig- und mittelenergetischer extrakorporaler Stosswellen [Evaluating the risk of sciatic nerve damage in the rabbit by administration of low and intermediate energy extracorporeal shock waves]. Z. Orthop. Ihre Grenzgeb. 1998, 136, 407–411. (In German) [Google Scholar] [PubMed]
  198. Liberati, A.; Altman, D.G.; Tetzlaff, J.; Mulrow, C.; Gotzsche, P.C.; Ioannidis, J.P.; Clarke, M.; Devereaux, P.J.; Kleijnen, J.; Moher, D. The PRISMA statement for reporting systematic reviews and meta-analyses of studies that evaluate healthcare interventions: Explanation and elaboration. BMJ 2009, 339, b2700. [Google Scholar] [CrossRef] [Green Version]
  199. Dakin, S.G.; Newton, J.; Martinez, F.O.; Hedley, R.; Gwilym, S.; Jones, N.; Reid, H.A.B.; Wood, S.; Wells, G.; Appleton, L.; et al. Chronic inflammation is a feature of achilles tendinopathy and rupture. Br. J. Sports Med. 2018, 52, 359–367. [Google Scholar] [CrossRef]
  200. Vidal, X.; Marti-Fabregas, J.; Canet, O.; Roque, M.; Morral, A.; Tur, M.; Schmitz, C.; Sitja-Rabert, M. Efficacy of radial extracorporeal shock wave therapy compared with botulinum toxin type a injection in treatment of lower extremity spasticity in subjects with cerebral palsy: A randomized, controlled, cross-over study. J. Rehabil. Med. 2020, 52, jrm00076. [Google Scholar] [CrossRef]
  201. van der Worp, H.; van den Akker-Scheek, I.; van Schie, H.; Zwerver, J. ESWT for tendinopathy: Technology and clinical implications. Knee Surg. Sports Traumatol. Arthrosc. 2013, 21, 1451–1458. [Google Scholar] [CrossRef] [Green Version]
  202. Cleveland, R.O.; Chitnis, P.V.; McClure, S.R. Acoustic field of a ballistic shock wave therapy device. Ultrasound Med. Biol. 2007, 33, 1327–1335. [Google Scholar] [CrossRef] [PubMed]
  203. Ogden, J.A.; Toth-Kischkat, A.; Schultheiss, R. Principles of shock wave therapy. Clin. Orthop. Relat. Res. 2001, 387, 8–17. [Google Scholar] [CrossRef] [PubMed]
  204. McClure, S.; Dorfmüller, C. Extracorporeal shock wave therapy: Theory and equipment. Clin. Techn. Equine Pract. 2003, 2, 348–357. [Google Scholar] [CrossRef] [Green Version]
  205. Maier, M.; Schmitz, C. Shock wave therapy: What really matters. Ultrasound Med. Biol. 2008, 34, 1868–1869. [Google Scholar] [CrossRef] [PubMed]
  206. Csaszar, N.B.; Angstman, N.B.; Milz, S.; Sprecher, C.M.; Kobel, P.; Farhat, M.; Furia, J.P.; Schmitz, C. Radial shock wave devices generate cavitation. PLoS ONE 2015, 10, e0140541. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  207. Mandal, C.C.; Ganapathy, S.; Gorin, Y.; Mahadev, K.; Block, K.; Abboud, H.E.; Harris, S.E.; Ghosh-Choudhury, G.; Ghosh-Choudhury, N. Reactive oxygen species derived from Nox4 mediate BMP2 gene transcription and osteoblast differentiation. Biochem. J. 2011, 433, 393–402. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  208. Wright, H.L.; McCarthy, H.S.; Middleton, J.; Marshall, M.J. RANK, RANKL and osteoprotegerin in bone biology and disease. Curr. Rev. Musculoskelet. Med. 2009, 2, 56–64. [Google Scholar] [CrossRef] [Green Version]
  209. Lee, T.C.; Staines, A.; Taylor, D. Bone adaptation to load: Microdamage as a stimulus for bone remodelling. J. Anat. 2002, 201, 437–446. [Google Scholar] [CrossRef]
  210. Shi, L.; Gao, F.; Sun, W.; Wang, B.; Guo, W.; Cheng, L.; Li, Z.; Wang, W. Short-term effects of extracorporeal shock wave therapy on bone mineral density in postmenopausal osteoporotic patients. Osteoporos. Int. 2017, 28, 2945–2953. [Google Scholar] [CrossRef]
  211. Snijdelaar, D.G.; Dirksen, R.; Slappendel, R.; Crul, B.J. Substance P. Eur. J. Pain 2000, 4, 121–135. [Google Scholar] [CrossRef]
  212. Mashaghi, A.; Marmalidou, A.; Tehrani, M.; Grace, P.M.; Pothoulakis, C.; Dana, R. Neuropeptide substance P and the immune response. Cell. Mol. Life Sci. 2016, 73, 4249–4264. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  213. Cao, Y.Q.; Mantyh, P.W.; Carlson, E.J.; Gillespie, A.M.; Epstein, C.J.; Basbaum, A.I. Primary afferent tachykinins are required to experience moderate to intense pain. Nature 1998, 392, 390–394. [Google Scholar] [CrossRef] [PubMed]
  214. Frias, B.; Merighi, A. Capsaicin, nociception and pain. Molecules 2016, 21, 797. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  215. Gamse, R.; Petsche, U.; Lembeck, F.; Jancso, G. Capsaicin applied to peripheral nerve inhibits axoplasmic transport of substance p and somatostatin. Brain Res. 1982, 239, 447–462. [Google Scholar] [CrossRef]
  216. Lam, F.Y.; Ferrell, W.R. Capsaicin suppresses substance p-induced joint inflammation in the rat. Neurosci. Lett. 1989, 105, 155–158. [Google Scholar] [CrossRef]
  217. Anand, P.; Bley, K. Topical capsaicin for pain management: Therapeutic potential and mechanisms of action of the new high-concentration capsaicin 8% patch. Br. J. Anaesth. 2011, 107, 490–502. [Google Scholar] [CrossRef] [Green Version]
  218. Jones, R. Nonsteroidal anti-inflammatory drug prescribing: Past, present, and future. Am. J. Med. 2001, 110, 4S–7S. [Google Scholar] [CrossRef]
  219. Santamato, A.; Cinone, N.; Panza, F.; Letizia, S.; Santoro, L.; Lozupone, M.; Daniele, A.; Picelli, A.; Baricich, A.; Intiso, D.; et al. Botulinum toxin type a for the treatment of lower limb spasticity after stroke. Drugs 2019, 79, 143–160. [Google Scholar] [CrossRef]
  220. Palazon-Garcia, R.; Alcobendas-Maestro, M.; Esclarin-de Ruz, A.; Benavente-Valdepenas, A.M. Treatment of spasticity in spinal cord injury with botulinum toxin. J. Spinal Cord. Med. 2019, 42, 281–287. [Google Scholar] [CrossRef]
  221. Quality Standards Subcommittee of the American Academy of Neurology and the Practice Committee of the Child Neurology Society; Delgado, M.R.; Hirtz, D.; Aisen, M.; Ashwal, S.; Fehlings, D.L.; McLaughlin, J.; Morrison, L.A.; Shrader, M.W.; Tilton, A.; et al. Practice parameter: Pharmacologic treatment of spasticity in children and adolescents with cerebral palsy (an evidence-based review): Report of the Quality Standards Subcommittee of the American Academy of Neurology and the Practice Committee of the Child Neurology Society. Neurology 2010, 74, 336–343. [Google Scholar]
  222. Pirazzini, M.; Rossetto, O.; Eleopra, R.; Montecucco, C. Botulinum neurotoxins: Biology, pharmacology, and toxicology. Pharmacol. Rev. 2017, 69, 200–235. [Google Scholar] [CrossRef] [PubMed]
  223. Cote, T.R.; Mohan, A.K.; Polder, J.A.; Walton, M.K.; Braun, M.M. Botulinum toxin type a injections: Adverse events reported to the US food and drug administration in therapeutic and cosmetic cases. J. Am. Acad. Dermatol. 2005, 53, 407–415. [Google Scholar] [CrossRef] [PubMed]
  224. Paget, S.P.; Swinney, C.M.; Burton, K.L.O.; Bau, K.; O’Flaherty, S.J. Systemic adverse events after botulinum neurotoxin a injections in children with cerebral palsy. Dev. Med. Child Neurol. 2018, 60, 1172–1177. [Google Scholar] [CrossRef] [PubMed]
  225. Harris, G.R. Effective treatment of chronic pain by the integration of neural therapy and prolotherapy. J. Prolother. 2010, 2, 377–386. [Google Scholar]
  226. Dullenkopf, A.; Borgeat, A. Lokalanästhetika. Unterschiede und Gemeinsamkeiten der “-caine” [Local anesthetics. Differences and similarities in the “-cains”]. Anaesthesist 2003, 52, 329–340. (In German) [Google Scholar] [CrossRef]
  227. Morgan, J.P.M.; Hamm, M.; Schmitz, C.; Brem, M.H. Return to play after treating acute muscle injuries in elite football players with radial extracorporeal shock wave therapy. J. Orthop. Surg. Res. 2021, 16, 708. [Google Scholar] [CrossRef]
  228. Melzack, R.; Wall, P.D. Pain mechanisms: A new theory. Science 1965, 150, 971–979. [Google Scholar] [CrossRef]
  229. Suputtitada, A.; Chen, C.P.C.; Ngamrungsiri, N.; Schmitz, C. Effects of repeated injection of 1% lidocaine vs. radial extra-corporeal shock wave therapy for treating myofascial trigger points: A randomized controlled trial. Medicina 2022, 58, 479. [Google Scholar] [CrossRef]
  230. Goats, G.C. Massage--the scientific basis of an ancient art: Part 2. Physiological and therapeutic effects. Br. J. Sports Med. 1994, 28, 153–156. [Google Scholar] [CrossRef] [Green Version]
  231. Kohrs, R.T.; Zhao, C.; Sun, Y.L.; Jay, G.D.; Zhang, L.; Warman, M.L.; An, K.N.; Amadio, P.C. Tendon fascicle gliding in wild type, heterozygous, and lubricin knockout mice. J. Orthop. Res. 2011, 29, 384–389. [Google Scholar] [CrossRef]
  232. Willkomm, L.M.; Bickert, B.; Wendt, H.; Kneser, U.; Harhaus, L. Weiterbehandlung und Rehabilitation nach Beugesehnenverletzungen [Postoperative treatment and rehabilitation following flexor tendon injuries]. Unfallchirurg 2020, 123, 126–133. (In German) [Google Scholar] [CrossRef] [PubMed]
  233. Pavan, P.G.; Stecco, A.; Stern, R.; Stecco, C. Painful connections: Densification versus fibrosis of fascia. Curr. Pain Headache Rep. 2014, 18, 441. [Google Scholar] [CrossRef] [PubMed]
  234. von Heymann, W.; Stecco, C. Fasziale Dysfunktionen [Fascial dysfunction]. Man. Med. 2016, 54, 303–306. (In German) [Google Scholar] [CrossRef]
  235. Zhang, L.; Fu, X.B.; Chen, S.; Zhao, Z.B.; Schmitz, C.; Wen, C.S. Efficacy and safety of extracorporeal shock wave therapy for acute and chronic soft tissue wounds: A systematic review and meta-analysis. Int. Wound J. 2018, 15, 590–599. [Google Scholar] [CrossRef] [PubMed]
  236. Burneikaitė, G.; Shkolnik, E.; Čelutkienė, J.; Zuozienė, G.; Butkuvienė, I.; Petrauskienė, B.; Šerpytis, P.; Laucevičius, A.; Lerman, A. Cardiac shock-wave therapy in the treatment of coronary artery disease: Systematic review and meta-analysis. Cardiovasc. Ultrasound 2017, 15, 11. [Google Scholar] [CrossRef]
Biomedicines 10 01084 g001 550
Figure 1. Systematic review flowchart of the literature search regarding studies on the effects of exposure of musculoskeletal tissue to extracorporeal shock waves performed according to the PRISMA guidelines [198] on 30 September 2021.
Figure 1. Systematic review flowchart of the literature search regarding studies on the effects of exposure of musculoskeletal tissue to extracorporeal shock waves performed according to the PRISMA guidelines [198] on 30 September 2021.
Biomedicines 10 01084 g001
Table
Table 1. Effects of the exposure of bone and cartilage tissue to extracorporeal shock waves (more details of the studies listed in this table are provided in Table S1).
Table 1. Effects of the exposure of bone and cartilage tissue to extracorporeal shock waves (more details of the studies listed in this table are provided in Table S1).
Ref.First
Author		YearMMorphological, Functional and Radiological Findings
    Findings of Molecular Biological Investigations
        Findings of Histological Investigations
[18]Li2021FIncreased mineral apposition rates, trabecular bone volume, number, thickness; decreased trabecular separation
    Increased expressions of ALP, OCN, RUNX2, OPG, SMAD2
[19]Inoue2021RIncreased trabecular bone microarchitecture and bone strength
    Decreased RANKL
[20]Inoue2021RIncreased bone/tissue volumes
        Increased osteoblast surface, decreased number of sclerostin-positive osteocytes
[21]Zhao2021R     Unaltered expressions of OCN, RUNX2, COL2, SOX9; decreased expressions of CEBPα and PPARγ; increased expression of YAP
        Increased proliferation
[22]Kobayashi2020FIncreased bone union rate, radiographic score
        Increased enchondral ossification, chondrogenic differentiation without inhibited proliferation
[23]Alshihri2020F         Unaltered cell migration; increased proliferation and osteogenic differentiation
[24]Hsu2020FIncreased bone strength, bone mineral density, trabecular thickness, bone /tissue volumes, porosity
        Increased expressions of BMP2, BMP4 and Wnt3a signaling; unaltered expression of IGF1
[25]Ramesh2020RIncreased bone length
        Increased number of proliferative chondrocytes of growth plate’s cartilage and diameter of hypertrophic chondrocytes; activation of IGF1 and NFkb; increased levels of BCL2 and BCL-xL
[26]Colbath2020F     Increased expression of ALP, decreased expressions of TGFb and VEGF
[27]Hashimoto2019F     Increased expressions of COL2a1, ACAN, CCN2, SOX9
        Increased meniscal healing score and BrdU/CCN2 ratio
[28]Senel2019FBone mineral density, bone mineral content
[29]Kim2019FIncreased structure and bone quality
    Decreased expressions of TNFa, IL1b, IL6, MMP3, MMP13, BMP7
        Increased cell viability; decreased number of apoptotic cells and pro-inflammatory, cartilage degradation markers
[30]Buarque de Gusmao2019F/R     F: increased Akt and FAK activity and TGFb1 expression
    R: increased FAK activity, decreased Akt expression
[31]Cheng2019FEnhanced bone volume and trabecular thickness
        Reduced synovitis and cartilage damage; decreased expression of MMP-13; enhanced expressions of RUNX2, SOX-9 and COL10A1; enhanced expressions of IGF1, TGFb1 and COL2 and decreased TUNEL activity
[32]Ginini2019FIncreased mineral density, enhanced bone formation
        Higher collagen orientation index, increased expressions of COL1 and OCN
[33]Ginini2018FHigher degree of bone formation and mature bone; increased bone mineral density, bone volume fraction, and trabecular thickness
        Enhanced expressions of BMP2, VEGF and PCNA
[34]Qi2018RImproved International Cartilage Repair Society (ICRS) score and macroscopic osteochondral appearance
[35]Koolen2018FCortical screws: increased bone formation and screw fixation. Cancellous screws: no alterations
[36]Mackert2017FImproved average stiffness and yield load
    Increased expressions of COL1a1, NR3A1, IGF1, OCN, TRAP
        Improved average ventral, dorsal and endosteal callus formation
[37]Tan2017F     ESWT alone: increased levels of A2B receptors; ESWT in combination with adenosine and A2BR agonists downregulated ACAN, COL1A2, COL2A1, SOX9 and SOX6
        ESWT + adenosine and A2BR agonists: inhibited chondrogenic differentiation
[38]Hsu2017n.s.     Increased expressions of ERK1, OPG, ALP, MMP13; potential activation of the 1α,25-Dihydroxyvitamin D3 Rapid Membrane Signaling Pathway
        Increased expression of PDIA3
[39]Yilmaz2017FIncreased osteoblastic activity, improved pain score
        Lower modified Mankin score
[40]Wang2017FImproved OARSI score and gross pathological changes, less cartilage defects, higher bone mineral density and bone volume, improved bone porosity and yield stress
        Increased expressions PCNA and OCN, decreased expression of TUNEL
[41]Chen2017FIn vivo: improved bone volume, trabecular volume, BV/TV, bone thickness and bone mineral density
    In vitro: increased expressions of COL1, RUNX2, OSX and ALP
        In vitro: enhanced proliferation and osteogenic differentiation; in vivo: increased bone formation and expressions of RUNX2 and OSX
[42]Onger2017F500 impulses per treatment: unaltered bone volume/bone density
1000 impulses per treatment: enhanced bone volume/bone density
        500 impulses per treatment: enhanced capillary volume, decreased connective tissue volume
1000 impulses per treatment: enhanced capillary volume; more positive areas of staining with VEGF, collagen antibody, BMP7 compared to control, but decreased capillary volume compared to 500 impulses; unaltered connective tissue volume
[43]Wang2017FImproved OARSI score and gross pathological changes, less cartilage defects, improved BV/TV ratio, improved bone porosity and trabecular thickness
        Decreased expression of TUNEL; higher amount of PCNA-positive cells and increased vascular density; increased cartilage thickness and sectional cartilage area; decreased modified Mankin score
[44]Lama2017FPrevention of bone-weight reduction and trabecular microarchitecture deterioration; restored serum parameters of ALP, RANKL, OPG and PTH due to illness
    Reduced cathepsin k, TNF-α levels, PPARγ and adiponectin transcription; increased RUNX2 and BMP2 expressions
[45]Catalano2017F     Increased ERK phosphorylation, ROS formation, RUNX2, ALP, BMP2
[46]Ma2017FHigher bone volume per tissue volume, trabecular thickness, trabecular number, osteoblast surface/bone surface, osteoid surface/bone surface, osteoid thickness, mineralizing surface/bone surface, mineralizing apposition rate and bone formation rate as well as a reduced trabecular separation
[47]Huang2016F     Increased expressions of OPG and BMP-2
[48]Notarnicola2016F     Increased expressions of BMP, ALP, OCN, COL1A1 and RUNX2
        Enhanced cell adhesion and proliferation
[49]Zhai2016F     Increased expression of OCN, core-binding factor α1 and decreased PPARγ
        Increased ALP content
[50]Dias dos Santos2015F     Increased contents of sulfated glycosaminoglycans and hyaluronic acid
[51]Wang2014FReduced arthritic area of injury joint, enhanced bone mineral density and bone strength, improved subchondral plate thickness and bone porosity, reduced cartilage damage
        Increased Mankin and Safranin O scores, improved alterations of the molecular levels due to the illness of Dickkopf-1, PCNA, VEGF and BMP-2
[52]Muzio2014F     Decreased ALP and OCN
        Increased cell growth
        Increased SMAD phosphorylation
[53]Oktas2014FNo radiologic differences
        Excised periosteum group: positive effect on bone healing
[54]Sun2013F     Shockwave-dependent ATP release that activated P2X7 receptors and downstream signaling events, which induced the differentiation
[55]Suhr2013F         Extended growth rate, proliferation, migration, cell tracking and wound healing; ameliorated cell migration meditated by active remodeling of the actin cytoskeleton as indicated by increased directed stress fiber formations
[56]Lyon2013FIncreased bony density
        More mature bone formation, better healing, higher density of the cartilage
[57]Wang2013FIncreased bone mineral density
        Improved Mankin and Safranin O scores; increased COL2; decreased MMP13
[58]Wang2013F         Treatment 1–2 times per week: improved Makin and Safranin O scores; increased COL2; decreased MMP13; increased vWF, VEGF, BMP-2 and osteocalcin; deteriorated effects after 3 treatments per week
[59]van der Jagt2013FIncreased cortical volume (CtV), higher trabecular connectivity and more plate-like and thicker trabeculae, increased trabecular bone volume fraction
[60]Oztemur2013RNo changes in bone length
        Increased blood vessel density, highly basophilic matrix and abundance of the differentiating chondrocytes
[61]Gollwitzer2013RNew bone formation
[62]Altuntas2012R         Higher specimens’ mean scores in bone fracture healing
[63]Notarnicola2012F     Reduction in COL1, OSX, bone sialoprotein and RANKL expressions, OCN and osteopontin; in summary: inhibiting effect on osteoclastogenesis
[64]Zhao2012RDecreased NO level, and severity of cartilage lesions
        Decreased chondrocyte apoptosis, enhanced Mankin score
[65]Kearney2012F         Increased cambium cell number, cambium cell thickness, osseous tissue and callus area, larger amount of osteoprogenitor tissue; improved results in combination with a bioactive scaffold
[66]Xu2012F     Promotion of Integrin alpha-5 and beta-1 expressions; induction of phosphorylation of FAK, which led to increased adhesion and migration of osteoblasts
[67]Wang2012F         Improved Mankin and Safranin O scores, increased COL2, VEGF, BMP2 and OCN expressions
[68]Erturk2012FNo alterations in MRI
        Edema, increased fibroblastic activity, neovascularization
[69]Wang2011FIncreased BMD, bone strength, modulus of elasticity
        Decreased Mankin score; improved Safranin O staining results; increased expressions of VWF, VEGF, BMP2, OCN and ALP; decreased expression of CTXII, cartilage oligomeric matrix protein
[70]van der Jagt2011FIncreased 99mTc-MDP uptake, increased trabecular and cortical bone volume, higher bone stiffness; no alterations in microcrack analysis
        Soft tissue damage, no periosteal damage, de novo bone with active osteoblasts and osteoids
[71]Notarnicola2011F     Increased expressions of RUNX2, COL1, OCN, IGF1, IGFBP3; decreased expressions of IGFBP-4 and -5
[72]Hausdorf2011F     Increased basic fibroblast growth factor; no significant alterations in TGFb
[73]Wang2011FIncreased bone mineral content
        Increased bone tissue; decreased fibrous tissue; increased expressions of VEGF, VWF, PCNA, OCN and BMP2; decreased expression of TUNEL
[74]Mayer-Wagner2010F Increased COL2A1 expression
        Ultrastructural expansion of the rough-surfaced endoplasmatic reticulum, detachment of the cell membrane and necrotic chondrocytes; increased tenascin-C and Chitinase-3-like protein 1; no alterations in Mankin score
[75]Muzio2010F Increased expressions of ALP, COL1, BMP-4, OCN
        Increased osteoblast activity as well as number and size of calcium deposits
[76]Lai2010FTreatment with 14kV: increased mineral density, biomechanical bone strength, intense osteoblastic cell recruitment, new bone formation
        Treatment with 14kV: intense osteoblastic cell recruitment, new bone formation, neovascularization, increased PCNA, VEGF, BMP-2; opposite effects after treatment with 21kV
[77]Qin2010FHigher fraction of new bone
        Increased VEGF expression in hypertrophic chondrocytes, promotion of regeneration of the fibrocartilage zone
[78]van der Jagt2009FDiminished bone loss, higher trabecular bone-volume fraction
        No differences in mineralization or osteoid appearance
[79]Iannone2009F     Increased expression of IL10; no alterations in TGFa, CD29 and CD105 expressions
[80]Tamma2009F     Increased expressions of BCL-2-associated X protein, RUNX2, OPN, bone sialoprotein, OCN and COL1; decreased RANKL/OPG ratio suggesting inhibition of osteoclastogenesis
[81]Lee2009FIncreased callus formation and both extension and flexion stiffness
[82]Tam2009FEnhanced trabecular bone mineral density, trabecular bone-volume fraction, trabecular thickness
        Increased mineral apposition rate
[83]Hofmann2008F     Altered expression of several genes involved in bone formation, osteoblast differentiation and skeletal development; no alterations in RUNX2, OSX, osteopontin, osteonectin, OC, TGFb1 expressions
        Enhanced mineralization and number of ALP-positive osteoblasts
[84]Tam2008F     Decreased cell viability 6 days after treatment; increased viability 18 days after treatment; increased cell proliferation 18 days after treatment
        Enhanced mineralization 35 days after treatment and AP activity 18 days after treatment
[85]Lee2008FNew bone formation
        Superior fusion mass
[86]Wang2008FIncreased bone strength
        Increased cortical bone formation; higher number of newly formed vessels; increased expression of VEGF, nitric oxide synthase 3, PCNA and BMP-2
[87]Moretti2008F     Decreased expression of IL10 and TNFa in both groups; no alteration in b1-integrin expression
[88]Tischer2008FDose-dependent new bone formation
        Dose-dependent new bone formation
[89]Ozturk2008F         Increased epiphyseal plaque thickness and number of chondrocytes
[90]Ma2007F     Increased VEGF expression
        Increased bone and osteoblast number; increased VEGF expression and microvessel density
[91]Murata2007R     Augmented uniform gene transfection and increased activity of vector-expressed genes
[92]Benson2007R     Decreased synthesis of GAG; no alterations in NO or Prostaglandin E2 synthesis
[93]Martini2006F     Dose- and device-dependent cell viability and expression of ALP, Capicua Transcriptional Repressor Pseudogene, OCN and TGFb
[94]Bulut2006FIncreased callus volume
Advanced bone healing
[95]Martini2005F        Enhanced transmembrane current and voltage dependence of Ca-activated/K channels
[96]Saisu2005FIncreased breadth of the acetabular roof and transient woven bone formation on the lateral margin
[97]Chen2004F     Increased TGFb1 and VEGF-A expressions
        Increased cell density and cell number of RP59-positive mesenchymal stem cells, subsequently enhanced differentiation into chondrocytes and osteocytes
[98]Saisu2004FEnhanced bone mineral content, long-bone length and width
[99]Chen2004F     Increased ALPase, COL1, COL2 and OCN expressions and [3H]-thymidine uptake; increased expressions and phosphorylations of ERK and p38
        Activated ERK and p38 expressions
[100]Pauwels2004FNo alterations in bone elasticity
[101]Wang2004n.s.     Induced superoxide production; enhanced TGFb1, RUNX2, OCN and COL1 expressions; increased bone alkaline phosphatase activity
        Increase in bone nodule formations, promotion of the CFU-stroma formation but not CFU-mix formation
[102]da Costa Gomez2004F/R         R: increased microcrack length, fESWT: increased microcrack density
[103]Takahashi2004FIncreased cortical thickening, bone mineral density, bone mineral content
        Enhanced expressions of COL1A1, COL2A1, OC and OPN; no alterations in expression of COL10A1
[104]Chen2003FIncreased callus size and calcium content, bone mineral density
    Increased ALP activity, OCN production, PCNA, TGFb1 and BMP-2 expressions
        Increased bone-tissue formation, progressive mesenchymal aggregation, enchondral ossification and hard callus formation
[105]Martini2003F     High intensity treatment (28 kV): decreased viability; reduced cell respiration; depressed ALP and NO synthesis; decreased expressions of OCN, TGFb and Procollagen-type I carboxy-terminal propeptide (PICP); low intensity treatment (14 kV) showed contrary effects with increased viability and cell respiration, increased ALP and NO synthesis as well as OCN and PICP expressions; generally negative affection of PICP production
[106]Martini2003F     Increased NO, OCN and TGFb1 production after low energy application (14kV); decreased cell viability and expression of all examined proteins at high application intensities (28 kV)
[107]Dorotka2003F     Increased cytotoxity in both chondrocytes and BMSCs at high application intensities (0.17mJ/mm2), compared to lower energy levels and control; unaltered cell proliferation at all energy levels
[108]Wang2003F     Increased expressions of BMP2, BMP3, BMP4 and BMP7
        Intensive mesenchymal cell aggregation, hypertrophic chondrogenesis and endochondral/intramembrane ossification; increased levels of PCNA, BMP2, BMP3 and BMP4
[109]Maier2002FDecreased bone metabolism after 10 days (detected by scintigraphy), but increased metabolism after 28 days; signs of soft-tissue oedema, epiperiosteal fluid and bone marrow oedema on MRI
        Epiperiostal deposits of hemosiderin
[110]Wang2002F     Increased ALP activity and TGFb1 expression
        Promotion of bone marrow stromal, but not hematopoietic cell growth; dose-dependent effect on formation of CFU osteoprogenitors
[111]Wang2001F     Induction of cell membrane hyperpolarization and consecutive Ras activation; induction of RUNX2; increased activity of bone ALP; increased expressions of OCN and COL1
        Increased bone-nodule formations
[112]Wang2001FMore callus formations
        More cortical bone and thicker, denser and heavier bone tissues
[113]Vaterlein2000FNeither macroscopic nor radiological alterations after high-intensity treatments
        No histological alterations after high-intensity treatments
[114]Peters1998F         Several damages to tissues after low-intensity treatment
[115]Augat1995FNeither alterations in biomechanical outcomes nor altered radiological results; tendency to deterioration of facture healing with increasing application intensities
[116]Forriol1994FNo effect on the periosteal surface of mature cortical bone, but on the endosteal surface induction of some new trabecular bone; delayed bone healing
[6]Graff1988FSoft-tissue bleeding
        Bone marrow hemorrhage and osteocyte damage 48 h after ESWT; increased callus and bone formation, focal regeneration, apposition of new bone, bone remodeling
Abbreviations: ACAN, aggrecan; Akt, protein kinase B; ALP, alkaline phosphatase; ATP, adenosine triphosphate; BCL, B-cell lymphoma; BMP, bone morphogenetic protein; BMSCs, bone marrow mesenchymal stem cells; BrdU, bromodeoxyuridine; CCN2, connective tissue growth factor; CEBPα, CAAT/enhancer binding protein; CFU, colony forming unit; COL, collagen; CTXII, C-telopeptide of collagen alpha-1(II) chain; ERK, extracellular signal-regulated kinases; F, focused extracorporeal shock waves; FAK, focal adhesion kinase; GAG, glycosaminoglycans; IGF, insulin-like growth factor; IL, interleukin; MMP, matrix metalloproteinase; NFkb, nuclear factor kappa-light-chain-enhancer of activated B cells; NO, nitric oxide; ns, not specified; NR3A1, estrogen-receptor alpha; OCN, osteocalcin; OPG, osteoprotegerin; OSX, osterix; PCNA, proliferating cell nuclear antigen; PDIA, protein disulfide-isomerase A; PPARγ, peroxisome proliferator-activated receptor gamma; PTH, parathyroid hormone; R, radial extracorporeal shock waves; RANKL, receptor activator of nuclear factor kappa-Β ligand; Ref, reference; ROS, reactive oxygen species; RUNX2, runt-related transcription factor 2; SMAD2, mothers against decapentaplegic homolog 2; T, type of extracorporeal shock waves; TGF, transforming growth factor; TNF, tumor necrosis factor; TRAP, tartrate-resistant acid phosphatase; TUNEL, terminal deoxynucleotidyl transferase dUTP nick end labeling; VEGF, vascular endothelial growth factor; vWF, von Willebrand factor; YAP, yes-associated protein.
Table
Table 2. Effects of the exposure of connective tissue to extracorporeal shock waves (more details of the studies listed in this table are provided in Table S2).
Table 2. Effects of the exposure of connective tissue to extracorporeal shock waves (more details of the studies listed in this table are provided in Table S2).
Ref.First
Author		YearMMorphological, Functional and Radiological Findings
    Findings of Molecular Biological Examinations
        Findings of Histological Examinations
[117]Haberal2021R         Decreased epidural fibrosis; unaltered acute/chronic inflammation and vascular proliferation
[118]Heimes2020R     Increased expression of MMP-9; decreased expression of MMP-13; unaltered expression of inducible nitric oxide synthase 2, HIF1α, VEGF
        Increased coverage of the transplant by vasculature, percentage of the vascularized area, increase in the vascularized area and number of vessel junctions
[119]Lu2020F     Increased ACL remnant cell viability; BMSC: increased expressions of Ki67, COL1 and COL3; unaltered expressions of TGFb and VEGF
        ACL cells: increased expression of COL1A1, TGFb and VEGF; BMSC: increased migration and expression of 5-Ethynyl-2’-deoxyuridine, COL1 and COL3; unaltered expression of VEGF and TGFb
[120]Basoli2020F     Increased proliferation, ATP release, ROS production, expressions of IL8, MCP1, HSP90 and HSP27; unaltered expression of IL6
[121]Schnurrer-Luke-Vrbanić2018R         Higher multiplication of collagen fibers; faster organization of muscle fibers and vascularization by treatment with radial shockwaves
[122]Cui2018F     Decreased expression of TGFb, a-SMA, vimentin, COL1A1, N-CAD and twist; increased expression of DNA-binding protein inhibitor ID1/2, E-CAD and FN after 24 h, but decreased expression of FN after 72 h
        Decreased cell migration
[123]Cai2016F     Initially decreased expression of IL6, IL8, MCP1 and TNFa; after 4 and more hours: increased expression of IL6 and IL8, unaltered expression of MCP1 and TNFa
[124]Hoch-strasser2016R     Induced mechanical cell destruction, dose-dependent decreased cell viability, increased growth potential of fibroblasts (not of JEG-3 cells), shift in proportion from G0/G1 to G2/M phase in fibroblasts (not in JEG-3 cells)
        Cellular detachments, holes in monolayers, disruption of actin filaments
[125]Leone2016F     Increased expressions of COL2A, SOX9, ALP and PPARy; unaltered expressions of OCN and RUNX2
        Increased expression of differentiation markers in cells grown in specific differentiation media
[126]Kisch2015FIncreased capillary blood velocity; unaltered postcapillary venous filling pressure
[127]Waugh2015R     Increased expressions of IL6, IL8, MMP2 complex and ProMMP9; unaltered expressions of IL1b, IL2, IL4, IL10, IL12p70, IL17A, VEGF, interferon-γ, active MMP9, ProMMP2 and active MMP2
[128]de Girolamo 2014F     Increased expressions of SCX, IL1b, IL6, IL10, TGFb and VEGF; unaltered expressions of MMP3, MMP13, COL1A1, COL3A1 and TNFa; reduced NO synthesis
[129]Chow2014F         Increased fibrocartilage area and thickness, proteoglycan deposition, expression of SOX9 and COLII and Vickers hardness; unaltered expression of COL1
[130]Cinar2013RDecreased load to failure
        Decreased collagen fiber density
[131]Contaldo2012R         Enhanced expressions of caspase-3, PCNA and eNOS; increase in functional angiogenetic density and total wound score
[132]Chow2012FIncreased load to failure, new bone area and new bone volume
        Increased fibrocartilage zone and ratio of bone forming
[133]Yoo2012F         Increased fibrillary diameter, vascularity, fibroblast activity, lymphocyte and plasma cell infiltration, dense histocytes; transient disorganization of collagen fibers
[134]Leone2012F     Ruptured tenocytes: decreased expressions of COL1 and SCX; unaltered COL3, tenomodulin, tenascin-C
        Healthy tenocytes: increased cell proliferation and migration
[135]Zhang2011F         Increased lubricine expression
[136]Penteado2011F         Unaltered blood-vessel number
[137]Kubo2010FReduced ear thickness
    Increased expressions of VEGF-C and VEGF-R3
        Increased density of lymphatic vessels
[138]Sugioka2010R         Increased introduction of NFkb decoy-FITC, activation of NFkb; decreased activation of NFkb after pretreatment with ESW+NFkb decoy-FITC
[139]Berta2009F     Decreased viability; increased expression of TGFb1; increase in COL1 and COL3 expressions after 6 days after a primary decreased expression
[140]Bosch2009F     Increased expressions of COL1 and MMP14; decreased expression of MMP3
        Unaltered total collagen content, disorganization of normal collagen structure; decreased percentage of degraded collagen 6 weeks after treatment after an increase 3 h after treatment
[141]Han2009F     Healthy: increased expression of IL1; unaltered expressions of MMP1, MMP2, MMP9, MMP13, IL6 and IL13
Diseased: decreased expressions of MMP1, MMP13 and IL6; unaltered expressions of MMP2, MMP9, IL1 and IL13
        Decreased cell viability
[142]Byron2009RRadiographic scores, scintigraphic navicular pool phase, delayed-phase region of interest density ratios
[143]Chao2008F     Increased total collagen concentration, NO production, expressions of PCNA, COL1, COL3 and TGFb
        Decreased cell viability; increased cell proliferation
[144]Wang2008FIncreased new bone formation, bone mineral status, tensile load and strength
        Increased remodeling/alignment of collagen fibers, thicker and mature regenerated fibrocartilage zone
[145]Bosch2007F     Unaltered DNA content, 3 h after treatment: increased GAG, total protein synthesis; 6 weeks after treatment: decreased GAG, collagen synthesis, noncollagenous protein synthesis, total protein synthesis
        Unaltered total collagen content, disorganization of normal collagen structure; decreased percentage of degraded collagen 6 weeks after treatment after an increase 3 h after treatment
[146]Kersh2006F         Unaltered percentage lesion, percentage disruption and gray scale, external width, fibroblast and tenocyte number, increased capillary density
[147]Wang2005FIncreased trabecular bone around the tendons and tensile strength of tendon/bone interface, better bone/tendon contact
[148]Chen2004FIncreased load to failure
        Decreased edema, swelling, inflammatory cell infiltration; increased expressions of TGFb, IGF1, tenocyte proliferation, neovascularization and progressive tendon tissue regeneration
[149]Orhan2004FHigher force to rupture
        Less adhesion formation, increased number of capillaries
[150]Hsu2004FIncreased ultimate tensile load
        Increased hydroxyproline concentration; decreased pyridinoline concentration; unaltered number of blast-like tenocytes (4 weeks); increased number of mature tenocytes (16 weeks)
[151]Orhan2004F         Disorganization of collagen fibers
[152]Wang2003F         Increased number of neo-vessels and expressions of eNOS, VEGF and PCNA
[153]Maier2002F         Exposure of tendons with high intensity ESWT: increased staining affinity, nuclear and fibrillar appearance paratendon: increased thickness, edema, capillary density
[154]Wang2002F         New capillary and muscularized vessels, newly appeared myofibroblasts; no alterations in bone matrix, bone vascularization and osteocyte activity
[155]Johannes1994F         Decreased cell viability, no alterations in cell growth
Abbreviations: a-SMA, alpha smooth muscle actin; ACL, anterior cruciate ligament; ALP, alkaline phosphatase; ATP, adenosine triphosphate; BMSCs, bone marrow mesenchymal stem cells; COL, collagen; F, focused extracorporeal shock waves; FITC, fluorescein isothiocyanate; FN, fibronectin; GAG, glycosaminoglycans; HIF, hypoxia-inducible factor; HSP, heat shock protein; IGF, insulin-like growth factor; IL, interleukin; MCP, monocyte chemoattractant protein; MMP, matrix metalloproteinase; NFkb, nuclear factor kappa-light-chain-enhancer of activated B cells; NO, nitric oxide; OCN, osteocalcin; PCNA, proliferating cell nuclear antigen; PPARγ, peroxisome proliferator-activated receptor gamma; R, radial extracorporeal shock waves; Ref, reference; ROS, reactive oxygen species; RUNX2, runt-related transcription factor 2; SCX, scleraxis; T, type of extracorporeal shock waves; TGF, transforming growth factor; TNF, tumor necrosis factor; VEGF, vascular endothelial growth factor.
Table
Table 3. Effects of the exposure of muscle and nerve tissue to extracorporeal shock waves (more details of the studies listed in this table are provided in Table S3).
Table 3. Effects of the exposure of muscle and nerve tissue to extracorporeal shock waves (more details of the studies listed in this table are provided in Table S3).
RefFirst
Author		YearMMorphological, Functional and Radiological Findings
    Findings of Molecular Biological Examinations
        Findings of Histological Examinations
[156]Huang2021RDecreased total contracture angle, muscle contracture angle
    Decreased expressions of TGFb and HIF1a
        Decreased proportion of collagen fiber area
[157]Kenmoku2021REnergy flux density- and total energy-dependent decrease in CMAP, unaltered CMAP latency
[158]Park2020FIncreased print width, print area
    Tendential increased expression of myelin basic protein
[159]Matsuda2020FImproved BBB locomotor function, increased withdrawal threshold, abbreviated latency of MEPs, no alterations in MEP amplitude
    Increased expressions of BDNF and TRKB
        Increased expression of BDNF, reduced myelin damage and oligodendrocyte loss, decreased axonal damage
[160]Langendorf2020R     Increased expressions of MyoD and myosin
        Initially higher amount of mononucleated cells; at day 7, newly formed muscle fibers with less MNCs; unaltered number of cells immunopositive for CD31
[161]Sagir2019FDecreased EMG amplitude, increased EMG latency, improved sciatic functional index
        Decreased myelin thickness, axon area and number
[162]Feichtinger2019FImproved load-to-failure testing results, intensity measurements in functional gait analysis
    Unaltered expressions of stromal cell-derived factor 1, TGFb1, TGFb3 and VEGFR2
[163]Yang2019n.s.Improved mechanical paw withdrawal threshold and thermal paw withdrawal latency
    Decreased TNFa, NFkb, MMP9, IL1b, NOX1, NOX2, NOX4, oxidized protein, cleaved caspase 3, cleaved PARP, γ-H2AX, (p)-p38, p-JNK, p-ERK1/2, Nav.1.3, Nav.1.8 and Nav.1.9
[164]Mattya-szovszky2018R     Dose-dependent increase in myogenic factor 5, MyoD, PAX7 and NCAM; downregulation of these proteins at double exposure of the highest energy flux density
        Increased cell viability at low energy flux densities, no alterations at higher energy flux densities
[165]Yin2018FIncreased angiogenesis, decreased serum myoglobin/creatine phosphokinase
    Decreased NOX1, NOX2, cleaved caspase 3, cleaved PARP, TGFb,
(p-)SMAD3, ICAM1, MMP9, TNFa, NFkb, chemokine (C-C motif) ligand 5, TLR2, TLR4, IL1b, cytosolic cytochrome C, γ-H2AX; increased Bcl-2, p-SMAD1/5, BMP-2, mitochondrial cytochrome C
        Decreased muscle-damaged/fibrosis/collagen-deposition areas
[166]Shin2018R     Increased expressions of DCX, SOX2, GAP43 and MAP2
        Increased expressions of DCX, SOX2, GAP43 and MAP-2
[167]Luh2018FEnhanced amplitude and latency of sensory nerve action potentials in combination with EMLA, compared to single EMLA and ultrasound+EMLA application
[168]Kenmoku2018RDecreased CMAP amplitude, unaltered CMAP latency
        Irregular end plates, unchanged axon terminals and muscle fibers, increased mean interjunctional fold interval
[169]Chen2017n.s.Improved mechanical paw withdrawal threshold and thermal paw withdrawal latency
    Decreased expressions of TNFa, NFkb, MMP9, IL1b, GFAP, ox42, NOX1, NOX2, NOX4, oxidized protein, γ-H2AX, cytosolic mitochondria, cleaved capase-3, PARP, p-P38, p-JNK, p-ERK1/2, Nav.1.3, Nav.1.8 and Nav.1.9
        Decreased expressions of p-P38+, peripherin+ cells, P38+ and NF200+ cells
[170]Yahata2016FImproved BBB locomotor score, withdrawal latency, 50% withdrawal threshold
        Increased expressions of VEGF, CD31, a-SMA and 5-HT; increased area of spared white matter; decreased number of TUNEL-positive cells
[171]Schuh2016F     Increased cell yield, BrdU assays, population doublings, S100b, c-Jun, GFAP and P75 expression; decreased P0 and P16 expressions, increased extracellular ATP levels immediately after application
[172]Lee2016n.s.Decreased knee-joint angle
[173]Kisch2016FIncreased muscular blood flow
[174]Lee2015n.s.Increased ankle angles (toe off + foot contact), improved sciatic functional index
    Increased expression of NT3
[175]Yamaya2014FImproved BBB locomotor score
    Increased expressions of VEGF and VEGF-receptor 1
        Increased NeuN-positive cells, VEGF staining
[176]Fu2014FImproved mechanical withdrawal threshold, thermal withdrawal latency
[177]Ishikawa2013RTransfection of POMC gene
[178]Mense2013FDecreased pressure pain threshold, improved locomotor activity
        Increased number of PGP 9.5-IR nerve fibers
[179]Hausner2012FIncreased amplitude, CMAP area
        Increased number of myelinated axons, unaltered number of endoneural vessels
[180]Kenmoku2012RDecreased amplitude, unaltered CMAP latency
        Decreased number of acetylcholine receptors
[181]Yamashita2009RDecreased mechanical allodynia
        Increased ratio of β-endorphin-IR muscle cells and number of β-endorphin-IR muscle fibers; decreased number of CGRP-IR DRG neurons
[182]Wu2008FDecreased motor nerve conduction velocity; unaltered sciatic functional index and withdrawal reflex latency
        Damage to the myelin sheath of large-diameter myelinated fibers
[183]Hausdorf2008F         Decreased number of unmyelinated nerve fibers of femoral nerve; unaltered number of unmyelinated nerve fibers of sciatic nerve; unaltered size, number and myelin sheet of myelinated nerve fibers
[184]Hausdorf2008F         Decreased number of neurons immunoreactive for substance P
[185]Lee2007FNo changes in motor and vegetative functions
        Decreased number of neurons during high-intensity treatment, dose-dependent myelin damage
[186]Ochiai2007FIncreased walking duration
        Decreased ratio of CGRP-positive dorsal root ganglion neurons
[187]Wu2007FDecreased motor nerve conduction velocity, unaltered sciatic functional index
[188]Murata2006F         Increased number of ATF3 and ATF-3/GAP-43 dual-IR neurons
[189]Takahashi2006F         Decreased number of epidermal nerve fibers
[190]Bolt2004RDecreased sensory nerve conduction velocity
        Disruption of myelin sheet
[191]Hausdorf2004F     Increased substance-P release 6 and 24 h after treatment, decreased substance-P release 6 weeks after treatment; unaltered prostaglandin-E2 release
[192]Takahashi2003F         Decreased percentage of CGRP-immunoreactive dorsal root ganglion neurons
[193]Maier2003F         Increased substance-P release after 6 and 24 h; decreased SP release after 6 weeks; no alterations in prostaglandin-E2 release
[194]Haake2002F     Unaltered c Fos expression
        Unaltered c Fos expression
[195]Ohtori2001F         Decreased number of nerve fibers immunoreactive for PGP 9.5 and CGRP
[196]Haake2001F         Unaltered expressions of met-enkephalin and dynorphin
[197]Rompe1998F         Vacuolic swelling of axons, no disruption of nerve’s continuity
Abbreviations: a-SMA, alpha smooth muscle actin; ATF, activating transcription factor; ATP, adenosine triphosphate; BCL, B-cell lymphoma; BDNF, brain-derived neurotrophic factor; BMP, bone morphogenetic protein; BrdU, bromodeoxyuridine; CFU, colony forming unit; CGRP, calcitonin gene-related peptide; CMAP, compound muscle action potential; DCX, doublecortin; DRG, dorsal root ganglion; EMG, electromyography; EMLA, eutectic mixture of local anesthetics; ERK, extracellular signal-regulated kinases; Ff, focused extracorporeal shock waves; GAG, glycosaminoglycans; GAP, growth associated protein; GFAP, glial fibrillary acidic protein; HIF, hypoxia-inducible factor; ICAM, intercellular adhesion molecule; IL, interleukin; IR, immunoreactive; JNK, jun N-terminal kinases; MAP, microtubule-associated protein; MEP, motor evoked potentials; MMP, matrix metalloproteinase; MNC, mononucleated cells; MyoD, myoblast determination protein 1; Nav, sodium channel, voltage-gated; NCAM, neural cell adhesion molecule; NeuN, hexaribonucleotide binding protein-3; NFkb, nuclear factor kappa-light-chain-enhancer of activated B cells; NOX, NADPH oxidase; NT, neurotrophin; PARP, poly (ADP-ribose) polymerase; PAX, paired box protein; PGP, protein gene product; POMC, proopiomelanocortin; R, radial extracorporeal shock waves; Ref, reference; T, type of extracorporeal shock waves; TGF, transforming growth factor; TLR, Toll-like receptor; TNF, tumor necrosis factor; TRKB, tropomyosin receptor kinase B; TUNEL, terminal deoxynucleotidyl transferase dUTP nick end labeling; VEGF, vascular endothelial growth factor; 5-HT, serotonin.
Table
Table 4. Take-home messages regarding the effects of exposure of musculoskeletal tissue to extracorporeal shock waves.
Table 4. Take-home messages regarding the effects of exposure of musculoskeletal tissue to extracorporeal shock waves.
No.Take-Home Message
1Compared to the effects of many other forms of therapy, the clinical benefit of extracorporeal shock wave therapy does not appear to be based on a single mechanism.
2Different tissues respond to the same mechanical stimulus in different ways.
3Just because a mechanism of action of extracorporeal shock wave therapy was described in a study does not automatically mean that this mechanism was relevant to the observed clinical effect.
4Focused and radial extracorporeal shock wave therapy seem to act in a similar way.
5Extracorporeal shock wave therapy stimulates both progenitor and differentiated cells, and has positive effects on pathologies of bone and cartilage.
6Extracorporeal shock wave therapy apparently mimics the effect of capsaicin by reducing substance-P concentration.
7Extracorporeal shock wave therapy apparently mimics effects of injection of Botulinum toxin A by destroying endplates in the neuromuscular junction.
8Extracorporeal shock wave therapy apparently imitates certain mechanisms of action of neural therapy.
9Extracorporeal shock wave therapy apparently imitates certain mechanisms of manual therapy treatments.
10Even the most sophisticated research into the effects of exposure of musculoskeletal tissue to extracorporeal shock waves cannot substitute clinical research in order to determine the optimum intensity, treatment frequency and localization of extracorporeal shock wave therapy.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Wuerfel, T.; Schmitz, C.; Jokinen, L.L.J. The Effects of the Exposure of Musculoskeletal Tissue to Extracorporeal Shock Waves. Biomedicines 2022, 10, 1084. https://doi.org/10.3390/biomedicines10051084

AMA Style

Wuerfel T, Schmitz C, Jokinen LLJ. The Effects of the Exposure of Musculoskeletal Tissue to Extracorporeal Shock Waves. Biomedicines. 2022; 10(5):1084. https://doi.org/10.3390/biomedicines10051084

Chicago/Turabian Style

Wuerfel, Tobias, Christoph Schmitz, and Leon L. J. Jokinen. 2022. "The Effects of the Exposure of Musculoskeletal Tissue to Extracorporeal Shock Waves" Biomedicines 10, no. 5: 1084. https://doi.org/10.3390/biomedicines10051084

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop