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Int. J. Mol. Sci. 2019, 20(3), 683;

Influence of Platelet-Rich and Platelet-Poor Plasma on Endogenous Mechanisms of Skeletal Muscle Repair/Regeneration
Department of Experimental and Clinical Medicine—Section of Anatomy and Histology, University of Florence, Largo Brambilla 3, 50134 Florence, Italy
Author to whom correspondence should be addressed.
Received: 17 January 2019 / Accepted: 1 February 2019 / Published: 5 February 2019


The morpho-functional recovery of injured skeletal muscle still represents an unmet need. None of the therapeutic options so far adopted have proved to be resolutive. A current scientific challenge remains the identification of effective strategies improving the endogenous skeletal muscle regenerative program. Indeed, skeletal muscle tissue possesses an intrinsic remarkable regenerative capacity in response to injury, mainly thanks to the activity of a population of resident muscle progenitors called satellite cells, largely influenced by the dynamic interplay established with different molecular and cellular components of the surrounding niche/microenvironment. Other myogenic non-satellite cells, residing within muscle or recruited via circulation may contribute to post-natal muscle regeneration. Unfortunately, in the case of extended damage the tissue repair may become aberrant, giving rise to a maladaptive fibrotic scar or adipose tissue infiltration, mainly due to dysregulated activity of different muscle interstitial cells. In this context, plasma preparations, including Platelet-Rich Plasma (PRP) and more recently Platelet-Poor Plasma (PPP), have shown advantages and promising therapeutic perspectives. This review focuses on the contribution of these blood-derived products on repair/regeneration of damaged skeletal muscle, paying particular attention to the potential cellular targets and molecular mechanisms through which these products may exert their beneficial effects.
fibrosis; myoblasts; myofibroblasts; myogenesis; Platelet-Rich Plasma (PRP); Platelet-Poor Plasma (PPP); satellite cells; skeletal muscle regeneration; stem cell niche; regenerative medicine

1. Introduction

Skeletal muscle can be considered as the largest organ of the human body, accounting for 40–45% of the total body mass, responsible for generating forces that guarantee breathing and movement. In addition, it represents an important metabolic and endocrine organ [1,2]. The incidence of skeletal muscle injuries as a consequence of trauma (e.g., sport injuries), inherited genetic diseases (e.g., muscular dystrophies), pathology (cancer and endocrinological disorders) or systemic conditions such as aging is very high worldwide, thus representing a serious socio-economic concern with relevant Health Care System costs [2,3]. Indeed skeletal muscle damage is the most common cause of severe long-term pain and physical disability, restricting patient daily living activities and imposing lost working days.
It is well known that skeletal muscle tissue possesses an intrinsic remarkable regenerative potential, which, however, becomes compromised in the case of severe and extended damage. In particular, it has been reported that skeletal muscle tissue is able to compensate for up to 20% of muscle mass loss but beyond this threshold the restoration of the native muscle tissue structure and function cannot be achieved [2].
Traditional therapeutic options for the treatment of damaged muscles include RICE (Rest, Ice, Compression and Elevation) treatment, rehabilitation therapies, corticosteroid administration and, in the worst conditions, even reconstructive surgical intervention. The progress of scientific research, that allowed a deeper understanding of the cellular and molecular mechanisms driving skeletal muscle tissue repair/regeneration, together with the advances of regenerative medicine and biotechnology, pushed the development and application of alternative and innovative strategies to promote or improve muscle repair/regeneration, such as cell-based therapy with different kinds of stem cells [4,5,6,7,8,9,10], tissue engineering [11,12] and Low Level Laser Therapy (LLLT—more recently termed Photobiomodulation) [13,14,15].
However, none of the therapeutic options so far adopted has proved to be resolutive and satisfactory; in addition, the most innovative therapeutic strategies, despite the encouraging elicited outcomes, still bear several criticisms hindering their clinical application for regenerative purposes. For more detailed information readers are referred to recent reviews [2,9,10] focusing on this topic, which is beyond the aim of this review.
Based on these considerations, the development of new effective treatments for skeletal muscle injury represents a priority and an urgent need. A current scientific challenge remains the identification of appropriate factors capable of limiting muscle degeneration and/or potentiating the endogenous muscle regenerative program. In this context, plasma preparations, including Platelet-Rich Plasma (PRP) and more recently Platelet-Poor Plasma (PPP), have shown several advantages and promising therapeutic perspectives.
This review, besides giving an updated description of the main cell types driving skeletal muscle repair/regeneration, focuses on the contribution of PRP and PPP on repair/regeneration of damaged skeletal muscle, paying particular attention to the cellular and molecular mechanisms through which these blood-derived products may exert their beneficial effects.

2. Adult Skeletal Muscle Repair and Regeneration: Role of Satellite and Non-Satellite Cells

It has been clearly proven that adult skeletal muscle tissue possesses a remarkable ability to regenerate in response to focal injuries. This is mainly thanks to the activity of a small population of resident mononucleated myogenic precursors, called satellite cells, whose name depends on the unique anatomical location at the periphery of a skeletal myofiber, beneath the surrounding basal lamina in intimate association with the myofiber sarcolemma, and in close proximity to capillaries and neuromuscular junction [16,17,18,19,20]. In healthy skeletal muscles, satellite cells are mitotically quiescent and transcriptionally inactive; in this dormant state they express the paired box transcription factor Pax7, necessary for their survival and function, and the myogenic transcription factor Myf5 (~90% of quiescent satellite cells) but not the myogenic regulatory factors, namely MyoD or myogenin [21,22]. In injured muscles, in response to signals coming from damaged myofibers and infiltrating inflammatory cells (neutrophils and macrophages), satellite cells become activated, giving rise to a progeny of proliferating Pax7, Ki67, Myf5 and MyoD positive adult myoblasts which subsequently, down-regulating Pax7 and expressing myogenin and MRF4/Myf6, differentiate into skeletal myocytes that finally either fuse with each other, forming nascent syncytial contractile myofibers, and fuse with injured myofibers, thus repairing the damage [21]. There is evidence indicating that a small percentage of satellite cells are true stem cells, capable of self-renewal, thereby ensuring the replenishment of the basal pool of resident satellite cells that are recruitable in the case of muscle re-injury [16,23,24].
The behavior and the fate of satellite cells are largely influenced by the dynamic interplay established with components of the surrounding microenvironment, which changes under homeostatic conditions (quiescent state) and during regeneration (activated state) [25]. This microenvironment includes the so called “immediate satellite cell niche” and the microenvironment beyond the immediate niche [16]. The immediate satellite niche represents the microenvironment where the satellite cells reside. It consists of several signaling molecules diffusing between the satellite cell and the myofiber, different extracellular matrix (ECM) components, satellite cell and myofiber surface-associated receptors for mediating cell-to-cell or cell-to-ECM interactions or binding different regulatory factors, and receptors present in the basal lamina to sequester inactive growth factor precursors secreted by either satellite cells and myofibers, serving as a local reservoir to be rapidly activated during muscle injury [16,26,27]. The microenvironment beyond this niche comprises the local milieu and the systemic milieu. The local milieu may be identified by the muscle fascicle wrapped by perimysium, consisting of other myofibers surrounded and connected to each other by endomysium sheath, a heterogeneous population of interstitial cells in the stroma between the myofibers [28,29,30], blood capillaries together with their secretable factors and associated pericytes and mesoangioblasts [31,32] and motor neuron endings. The main interstitial cells types are represented by fibroblasts, mesenchymal stem/stromal cells (MSCs) including fibro-adipogenic progenitors -FAPs- [33], telocytes (CD34+/vimentin+ stromal cells with a small cell body and distinctive extremely long, thin and moniliform cytoplasmic extensions called telopodes alternating slender segments—podomeres- with dilatations-podoms) [34], Abcg2+ side population (SP) [35], skeletal muscle-derived CD34+/45− (Sk-34) myogenic endothelial progenitors [36], interstitial stem cells positive for stress mediator PW1 expression and negative for Pax7 expression termed PICs [37], integrin β4 interstitial cells.
The systemic milieu includes the entire muscle belly along with bones and surrounding skeletal muscles, the immune cells and circulating growth factors, interleukins/chemokines and hormones.
The dynamic and collaborative interaction between satellite cells and the different cell types of the surrounding microenvironment, becomes crucial for a proper execution of the essential events of repair/regeneration process. Indeed, in contrast to the quiescent conditions, in the activated ones, many cells are found close to satellite cells exerting a supportive role for their functionality. On the other hand, this cell interaction is bidirectional since satellite cells can influence the behavior of interacting cells [20,38].
The cells supporting satellite cell-mediated regeneration in the activated niche may include several cell types.
Pro-inflammatory phagocytic macrophages (M1) and anti-inflammatory pro-regenerative macrophages (M2) have been demonstrated to promote proliferation and differentiation of myogenic precursors respectively, via both paracrine and juxtacrine signaling [39,40,41,42]. The ability of macrophages to rescue myoblasts and myotubes from apoptosis has also been demonstrated [43].
Fibroblasts-myofibroblasts and FAPs are the major contributors to the deposition and remodeling of the transitional ECM after a muscle lesion, required to rapidly restore tissue integrity [44]; on the other hand the capability of fibroblasts to promote myoblast proliferation and differentiation and to enhance satellite cell renewal as well as pro-myogenic function of FAPs has been documented [38,45,46,47,48,49].
Telocytes have been supposed to play a “nursing” role in satellite cell-mediated regeneration. By means of their telopodes they connect with each other via homocellular junctions, or with neighboring cells including satellite cells via heterocellular ones, thus forming a three-dimensional network in the interstitium: telocytes might act as a guidance stromal scaffold able to carry signals over long distances, driving satellite cell proliferation, migration and differentiation after their recruitment [34]. In addition, telocytes may modulate satellite cell function in a paracrine manner by the release of extracellular vesicles containing myogenic factors (e.g., Vascular Endothelial Growth Factor, VEGF, or microRNAs) [4,34,50,51].
Capillary endothelial cells secrete different paracrine factors strongly stimulating growth of myogenic progenitors and/or protecting them from apoptosis [19,52,53], whereas periendothelial cells including pericytes are crucial for the re-entry of satellite cells into quiescence at the end of the regeneration process and myofiber growth [54,55].
In addition, motor neurons and Schwann cells secreting neurotrophic factors including Insulin Growth Factor (IGF)-1, Nerve Growth Factor (NGF), Brain-Derived Growth Factor (BDNF) and Ciliary Neurotrophic Factor (CNTF) may contribute to the modulation of satellite cell/myoblast viability, proliferation and fusion [16,20,29,56,57].
Furthermore, in regulating satellite cell quiescence, activation, proliferation and differentiation an essential role is played by ECM factors (both of basal lamina and of interstitial matrix) including specific ligands, soluble factors sequestered within the matrix, as well as by the mechanical properties of ECM itself as extensively discussed in the review by Thomas and co-workers [27].
Many works have demonstrated that, in addition to satellite cells, other cell types residing within muscle or recruited via circulation may contribute to muscle regeneration thanks to their inducible myogenic potential [58]. These so-called myogenic non-satellite cells include: the interstitial Abcg2+SP [35,59,60,61], skeletal muscle-derived CD34+/45− (Sk-34) cells (likely a subpopulation of SP with more pronounced myogenic potential) [36], PICs [37], mesoangioblasts and pericytes [31,62,63,64], integrin β4 interstitial cells, CD133+ human skeletal muscle derived and blood- derived stem cells [65,66,67]. However, if these cells represent an independent source of muscle progenitors undergoing unconventional myogenic differentiation or if they give rise to satellite cells, remains to be elucidated. Moreover, also the molecular mechanisms guiding the lineage switch of these muscle interstitial or circulating cells in the regenerating environment are still unclear [28,29]. Based on all of this evidence, it appears clear that, for an effective restoration of muscle structure and function, collaborative and temporally coordinated juxtacrine and paracrine interactions among many myogenic and non-myogenic cells, are required.
Unfortunately, in case of severe and extended damage, with an intense and persisting inflammatory reaction or in disease settings, the muscle repair may become aberrant, occurring with a maladaptive fibrotic scar or adipose tissue infiltration, or even with heterotopic ossification, mainly as a consequence of dysregulated activity and number of fibroblasts and mesenchymal progenitors [3,33,49,68,69,70], which hamper the muscle regenerative response. Moreover, a critical event that must be considered for the achievement of a regenerating functional muscle tissue after injury is the re-establishment of neuromuscular junctions for the new myofibers, which is mandatory to prevent muscle wasting [71].
On the basis of these considerations, improving the functionality of satellite and non-satellite myogenic cells either directly or by acting on their microenvironment (e.g., modulating the inflammatory response or MSC functionality thus limiting fibrosis or muscle fatty deposition) as well as nerve regeneration, could represent the final goal of effective therapeutic strategies for efficient muscle regeneration.

3. Plasma Preparations: Platelet-Rich Plasma (PRP) and Platelet-Poor Plasma (PPP)

3.1. Definition and Biological Properties

PRP and PPP can be defined as a plasma fraction with a concentration of platelets respectively above and below baseline levels in whole blood. Unfortunately, so far, no univocal guidelines are available for these plasma preparations and protocols show a high variability among authors, thus leading to plasma fractions differing in terms of concentration of blood cells (platelets and leukocytes) and content and type of cytokines and growth factors. For a detailed and updated overview of the different methods of plasma fraction collection, of the commercially available PRP systems and of PRP classification, the readers are referred to several excellent reviews [72,73,74,75]. In any case, the rationale for using plasma fractions, in particular PRP, for regenerative purposes in different areas of medicine [76,77,78,79,80,81] including musculoskeletal and sport medicine [82,83,84,85], relies on the fact that they represent a cost-effective reservoir of numerous biologically active molecules, holding a strong potential for improving tissue healing and regeneration [86,87,88,89]. Moreover, the prompt availability from whole blood of patients and thus the autologous source of these blood products, posing no risks of disease transmission or immunogenic reactions [90,91], as well as the ease of administration, represent additional advantages for their clinical use. Furthermore, recent findings demonstrating the safety and favorable outcomes of the application of allogenic PRP to treat musculoskeletal conditions have opened new perspectives for off-the-shelf PRP therapy for all patients for whom the use of autologous PRP would not be recommended [92,93].

3.2. Contribution of PRP to Skeletal Muscle Repair/Regeneration

Some studies have reported positive outcomes after administration of PRP in patients with injured skeletal muscles, without negative side effects. Indeed, patients/athletes with acute muscle strains after PRP intralesional injections combined with a rehabilitation treatment, exhibited an earlier “return to play”, faster pain relief without a significant increase of the re-injury risk in short and long term, when compared to patients undergoing a rehabilitation program only [94,95,96,97,98]. Improvement of inflammatory state, reduction of fibrotic scar size and parenchymal recovery was also demonstrated in PRP-treated muscle lesions [94,99,100,101]. However, despite the encouraging findings, these studies do not reach sufficient statistical significance to support the adoption of PRP therapy for skeletal muscle injury in clinical routine, as recently extensively discussed [83,84,102,103,104]. Therefore, further human studies, to ascertain and validate the effective therapeutic benefits of PRP for skeletal muscle regenerative purpose, are strongly required.
On the other hand, a large body of experimental evidence supporting the contribution of PRP to the morpho-functional recovery of damaged skeletal muscles comes from studies carried out in animal models. Although these studies cannot be directly validated or extrapolated to human species, they do provide a robust and instructive scientific background to design and perform clinical investigations.
In particular, PRP injections into skeletal muscles of rats or mice subjected to different traumatic injuries (incision, laceration, contusion or lengthening/eccentric contractions) or to cardiotoxin injection or into ischemic muscles, have been demonstrated to contribute to the muscle healing process: (i) by modulating the inflammatory response including the increase in M2 macrophage cell recruitment to the injury site and function [105,106,107,108]; (ii) by generating a myogenic response, as evaluated by satellite cell activation, increase of the expression of different myogenic regulatory factors, modulation of the expression of muscle specific microRNAs and activation of myogenic signaling pathways leading to myofiber formation [105,106,109,110,111,112,113]; (iii) by attenuating the impairment of myocytes mitochondrial function determined by muscle damage and improving their endogenous antioxidant defense system [114]; and (iv) by protecting cells from apoptosis [108,111].
In addition, the reduction of type-I collagen deposition and scar formation (fibrosis) [106,110,112,113,115,116,117,118] the enhancement of angiogenesis [106,110,112,116,117] and a faster functional recovery [108,109,113,118,119] have been also observed after PRP application on damaged muscles. Takase and co-workers [120] recently demonstrated also the ability of PRP to prevent fatty degenerative changes of rotator cuff muscles in a rat rotator cuff tear model, when administered into subacromial space.

3.3. Impact of PRP on Satellite Cells and on Myogenic and Non-Myogenic Interstitial Cells

The cellular and molecular mechanisms that could mediate the beneficial pro-regenerative and anti-fibrotic effects of PRP-derived growth factors on muscle tissue healing have been investigated in a growing number of in vitro studies.
Satellite cells may represent a direct target of PRP action. Indeed, the capability of PRP to positively influence the behavior of primary myoblasts—human skeletal myoblasts [116,121], human pre-plated muscle derived-progenitor cells [122], rabbit myogenic progenitor cells [123], rat intrinsic skeletal muscle cells [124]—or satellite cell-derived myoblast line namely murine C2C12 myoblasts [120,121,125,126] and human CD56+ myoblasts [127] by promoting their activation and proliferation [116,120,121,122,123,124,125,126,127] and protecting them from apoptosis [116] has been demonstrated.
Among the different growth factors within PRP, Platelet Derived Growth Factor (PDGF) [122], has been identified as a key factor mediating PRP-induced mitogenic response, whereas the involvement of others—demonstrated to be contained in PRP [86,87,88,89]—has been proposed on the basis of previous studies investigating their effects on myoblast cell line or primary muscle stem cells, such as VEGF [4,128], Hepatocyte Growth Factor (HGF) [129] or IGF-1 [130]. The involvement of PDGF and VEGF in mediating the PRP effects may be also presumed on the basis of the recent study of Scully and co-workers [131] showing that platelet releasate is capable to drive C2C12 myoblast proliferation and terminal differentiation, as well as the commitment to differentiation of myofiber-derived stem cells, at least in part via PDGF and VEGF signaling pathways.
However, it must be pointed out that the effects of the combination of different growth factors (such as PRP) may be completely different from the ones elicited by the single growth factors, based on the proved antagonistic or synergistic cross-talk between diverse growth factor-mediated signaling.
In addition, primary cultured skeletal muscle cells treated with PRP have also shown an increased motility/migratory ability associated with an up-regulation of different focal adhesion proteins and F-actin cytoskeleton remodeling [132]; these findings are of particular interest given that migration of satellite cells and of satellite derived-myoblasts is a crucial process in muscle regeneration by which the cells reach the injured site.
In line with these findings, our research group has recently shown that PRP used as single treatment, positively influenced C2C12 myoblast viability and proliferation in the same manner of standard culture media containing animal sera, by promoting the activation of AKT-mediated signaling, as well as the activation of cultured murine satellite cells isolated from single skeletal muscle fibers [133].
In this paper, we also demonstrated that PRP treatment induced C2C12 myoblasts to enter and progress into the myogenic program by stimulating MyoD, myogenin, α-sarcomeric actin expression as standard differentiation culture media containing animal sera did, accordingly to a previous study [127]. The pro-myogenic effect of PRP was also recently demonstrated by the study of McClure and co-workers [126] where C2C12 myoblasts cultured on PRP embedded ECM mimicking scaffold, exhibited a PRP-dose dependent increase in myogenin and myosin heavy chain protein expression, mediated by ERK1/2 signaling activation. In addition, our study reported that PRP promoted also the C2C12 myoblast expression of matrix metalloprotease (MMP)-2 [133] whose function has been reported to be required for satellite cell activation [134,135,136], for basal lamina degradation and, at the elongation stage of the myogenic differentiation process, for completing the successive myoblast cell migration and fusion [6,27,137,138] thus supporting the pro-myogenic effect of PRP. These latter results concerning differentiation seem to question findings in the literature showing a reduction or even inhibition of myogenic differentiation of myoblasts cultured with PRP [120,121,125]. These contradictory PRP-elicited biological responses may be attributed to different experimental settings and more likely to the heterogeneity of PRP preparation techniques and formulations used, which may contain interplaying pro-myogenic growth factors—such as IGF-1 [130], HGF [129] and β-Fibroblast Growth Factor (FGF) [125,139]—and anti-myogenic ones—such as Transforming Growth Factor (TGF)-β [140]—in different concentrations and proportions. In addition, the different PRP dosages used may account for the discrepancy in the literature on whether PRP hampers or improves differentiation, when considering the dose-dependence of some myogenic cell responses as well as the timing of PRP application [116,121,125,126,132,133].
Another very interesting finding of our recent work [133] is the ability of PRP to satisfactorily support and stimulate in vitro viability, survival and proliferation of MSCs. Taking into consideration the close morpho-functional relationship between satellite and the interstitial stromal cells, in particular the reported supportive juxtacrine and paracrine role of different stromal cell types for satellite cells, our results may suggest that the muscle resident stromal cells could also benefit from the treatment with PRP. In other words, PRP might indirectly promote satellite cell-mediated regeneration by potentiating the nursing role of interstitial MSCs.
PRP may impact indirectly on satellite cells also by favoring the appearance and functionality of pro-regenerative macrophage phenotype (M2) within the healing niche on the basis of recent in vitro studies, evaluating the effects of different PRP preparations on human monocyte-derived cells [141,142] and also being consistent with the findings of in vivo research [106].
On the other hand, PRP could also affect the myogenic potential of the so-called non-satellite myogenic cells. In this line, Li and co-workers [122] have reported that pericytes, isolated from post-mortem human skeletal muscle biopsies, exhibited an enhanced proliferative ability when cultured with PRP as compared to standard culture media while maintaining their in vitro myogenic differentiation capability and in vivo myogenic potential.
Interestingly, we have recently demonstrated the capability of PRP to prevent the transition of fibroblasts into myofibroblasts, the main drivers of tissue scarring [143] via VEGF-A/VEGF-A Receptor-1-mediated inhibition of TGF-β1/Smad3 signaling [144]. These findings, in accordance with other studies [145,146], may contribute to the identification of the cellular and molecular mechanisms responsible for the observed reduction in the fibrotic response achieved after injections of PRP in damaged skeletal muscles [106,110,112,113,115,116,117,118]. Such reduction is necessary for the recreation of a more hospitable and conductive microenvironment for muscle progenitor functionality and for axonal growth/regeneration and muscle re-innervation [147] and eventually for a complete tissue morpho-functional recovery. On the other hand, the ability of different PRP-derived factors to positively influence Schwann cell function in vitro and to assist peripheral nerve repair/regeneration in vivo has been documented [148,149,150,151], thus allowing us to include Schwann cells in the list of the potential cell targets of PRP in the skeletal muscle tissue during tissue repair/regeneration (Figure 1) and to suggest that promotion of muscle re-innervation may represent an additional benefit exerted by PRP for damaged muscles.

3.4. Impact of PPP on Skeletal Myogenesis

PPP, long regarded as the waste product of PRP and used a sham treatment [109], has recently been demonstrated to exert a beneficial effect on myogenesis. In particular, Miroshnychenko and co-workers [121] showed that PPP, differently from PRP, reduced the proliferation rate of human primary skeletal muscle myoblasts but led to a significant induction of differentiation of these cells into the myogenic pathway and myotube formation. The same cell responses were elicited by a modified preparation of PRP, by means of a second spin to remove platelets, suggesting that the beneficial effects of PRP on myogenesis can be mostly due to plasma per se. Indeed, although PPP by its definition contains a very low concentration of platelets and therefore smaller quantities of growth factors, it still represents a reservoir of bioactive molecules (such as PDGF, IGF-1) [87], which may be responsible for the observed pro-myogenic effects on myoblasts. This research is in line with previous studies showing the effectiveness of PPP in evoking biological responses from different cell types involved in the healing process of several tissues, namely fibroblast migration and ECM remodeling [152,153], periodontal ligament stem cell differentiation towards osteoblastic phenotype [154], tenocyte proliferation and collagen production [155], endothelial cell differentiation towards angiogenic cells [156] and macrophage anti-inflammatory activity [157]. Miroshnychenko and co-workers [121] conclude that PPP and platelet deprived-PRP may be more appropriate to promote skeletal muscle regeneration than the traditionally formulated PRP, probably containing a greater quantity of platelet- derived factors detrimental for myoblast differentiation such as TGFβ-1. However, this laboratory evidence does not seem to have received a large consensus from the current literature [103,109].
Therefore, although PPP may hold promise for skeletal muscle injuries, further investigations to assess the exact growth factors contained in this kind of plasma formulation and especially to validate the impact of PPP on myogenic precursors, as well as to disclose its role on skeletal muscle tissue regeneration in vivo are absolutely required. Moreover, to date, no clinical studies evaluating the effect of PPP on skeletal muscle have been conducted.

4. Conclusions and Further Directions

Plasma preparations and especially PRP have been demonstrated to hold a strong therapeutic potential for the healing of injured skeletal muscle tissue due to their ability to potentiate the endogenous mechanisms of tissue repair/regeneration while contributing to limit the aberrant responses such as fibrosis. Nevertheless, despite these encouraging outcomes, evidence from animal studies and even more from human ones, are not still sufficient to attain the effective clinical translation of these blood products for skeletal regenerative purpose. Therefore, further investigations are necessary to validate the regenerative potential of these blood derivates and ultimately drive medical decisions. On the other hand, it must be considered that some human and animal studies have reported limited effectiveness or inefficacy of a PRP therapy for damaged skeletal muscle in terms of tissue regeneration and recovery of functionality, or even an exacerbation of the fibrotic response [158,159,160,161,162,163,164,165,166,167,168]. The main reason for these conflicting results certainly could be due to individual-based variations and muscle lesion type and severity but is most likely due to the great heterogeneity of the injected available products and application timing. Therefore, standardization of PRP preparation techniques as well as of application protocols—considering the cascade of events through which skeletal muscle tissue repair/regeneration progresses—is a priority in order to perform meaningful comparative analyses, to enable reproducibility and reach reliable conclusions. Moreover, a full characterization of plasma preparations is also necessary, evaluating the quantity and the type of the contained bioactive factors, as well as a deep investigation of their effects on the main local cell types involved in the skeletal muscle tissue regeneration. This may lead to novel and optimized plasma formulations for muscle regenerative purposes, based on the selection of factors capable, for instance, of exerting a pro-regenerative and anti-fibrotic action on skeletal muscle tissue.

Author Contributions

Conceptualization, review of the literature, F.C., A.T., C.S.; writing—original draft preparation, C.S.; preparation of figure, C.S.; writing—review and editing, F.C., A.T., S.Z.-O. and C.S. All the authors have read and approved the submitted version of the review.


The publication of this review was supported by FFABR-MIUR 2017 (Funding Fund for Basic Research Activities - Ministry of Education, University and Research, Italy) to C.S.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Giudice, J.; Taylor, J.M. Muscle as a paracrine and endocrine organ. Curr. Opin. Pharm. 2017, 34, 49–55. [Google Scholar] [CrossRef] [PubMed]
  2. Liu, J.; Saul, D.; Böker, K.O.; Ernst, J.; Lehman, W.; Schilling, A.F. Current Methods for Skeletal Muscle Tissue Repair and Regeneration. Biomed. Res. Int. 2018, 16, 1984879. [Google Scholar] [CrossRef] [PubMed]
  3. Schaaf, G.; Sage, F.; Stok, M.; Brusse, E.; Pijnappel, W.W.M.; Reuser, A.J.; vd Ploeg, A.T. Ex-vivo Expansion of Muscle-Regenerative Cells for the Treatment of Muscle Disorders. J. Stem Cell Res. 2012, S11, 003. [Google Scholar] [CrossRef]
  4. Sassoli, C.; Pini, A.; Chellini, F.; Mazzanti, B.; Nistri, S.; Nosi, D.; Saccardi, R.; Quercioli, F.; Zecchi-Orlandini, S.; Formigli, L. Bone marrow mesenchymal stromal cells stimulate skeletal myoblast proliferation through the paracrine release of VEGF. PLoS ONE 2012, 7, e37512. [Google Scholar] [CrossRef] [PubMed]
  5. Sassoli, C.; Zecchi-Orlandini, S.; Formigli, L. Trophic actions of bone marrow-derived mesenchymal stromal cells for muscle repair/regeneration. Cells 2012, 1, 832–850. [Google Scholar] [CrossRef] [PubMed]
  6. Sassoli, C.; Nosi, D.; Tani, A.; Chellini, F.; Mazzanti, B.; Quercioli, F.; Zecchi-Orlandini, S.; Formigli, L. Defining the role of mesenchymal stromal cells on the regulation of matrix metalloproteinases in skeletal muscle cells. Exp. Cell Res. 2014, 323, 297–313. [Google Scholar] [CrossRef] [PubMed]
  7. Berry, S.E. Concise review: Mesoangioblast and mesenchymal stem cell therapy for muscular dystrophy: Progress, challenges, and future directions. Stem Cells Transl. Med. 2015, 4, 91–98. [Google Scholar] [CrossRef] [PubMed]
  8. De Albornoz, P.M.; Aicale, R.; Forriol, F.; Maffulli, N. Cell Therapies in Tendon, Ligament, and Musculoskeletal System Repair. Sports Med. Arthrosc. Rev. 2018, 26, 48–58. [Google Scholar] [CrossRef]
  9. Andia, I.; Maffulli, N. New biotechnologies for musculoskeletal injuries. Surgeon 2018. [Google Scholar] [CrossRef] [PubMed]
  10. Dunn, A.; Talovic, M.; Patel, K.; Patel, A.; Marcinczyk, M.; Garg, K. Biomaterial and stem cell-based strategies for skeletal muscle regeneration. J. Orthop. Res. 2019. [Google Scholar] [CrossRef]
  11. Han, W.M.; Jang, Y.C.; García, A.J. Engineered matrices for skeletal muscle satellite cell engraftment and function. Matrix Biol. 2017, 60–61, 96–109. [Google Scholar] [CrossRef] [PubMed]
  12. Kwee, B.J.; Mooney, D.J. Biomaterials for skeletal muscle tissue engineering. Curr. Opin. Biotechnol. 2017, 47, 16–22. [Google Scholar] [CrossRef] [PubMed]
  13. Alves, A.N.; Fernandes, K.P.; Deana, A.M.; Bussadori, S.K.; Mesquita-Ferrari, R.A. Effects of low-level laser therapy on skeletal muscle repair: A systematic review. Am. J. Phys. Med. Rehabil. 2014, 93, 1073–1085. [Google Scholar] [CrossRef] [PubMed]
  14. Sassoli, C.; Chellini, F.; Squecco, R.; Tani, A.; Idrizaj, E.; Nosi, D.; Giannelli, M.; Zecchi-Orlandini, S. Low intensity 635 nm diode laser irradiation inhibits fibroblast-myofibroblast transition reducing TRPC1 channel expression/activity: New perspectives for tissue fibrosis treatment. Lasers Surg. Med. 2016, 48, 318–332. [Google Scholar] [CrossRef] [PubMed]
  15. De Oliveira, H.A.; Antonio, E.L.; Silva, F.A.; de Carvalho, P.T.C.; Feliciano, R.; Yoshizaki, A.; Vieira, S.S.; de Melo, B.L.; Leal-Junior, E.C.P.; Labat, R.; et al. Protective effects of photobiomodulation against resistance exercise-induced muscle damage and inflammation in rats. J. Sports Sci. 2018, 36, 2349–2357. [Google Scholar] [CrossRef] [PubMed]
  16. Yin, H.; Price, F.; Rudnicki, M.A. Satellite cells and the muscle stem cell niche. Physiol. Rev. 2013, 93, 23–67. [Google Scholar] [CrossRef] [PubMed]
  17. Scicchitano, B.M.; Sica, G.; Musarò, A. Stem Cells and Tissue Niche: Two Faces of the Same Coin of Muscle Regeneration. Eur. J. Transl. Myol. 2016, 26, 6125. [Google Scholar] [CrossRef] [PubMed]
  18. Le Grand, F.; Rudnicki, M.A. Skeletal muscle satellite cells and adult myogenesis. Curr. Opin. Cell Biol. 2007, 19, 628–633. [Google Scholar] [CrossRef] [PubMed]
  19. Mounier, R.; Chrétien, F.; Chazaud, B. Blood vessels and the satellite cell niche. Curr. Top. Dev. Biol. 2011, 96, 121–138. [Google Scholar] [CrossRef] [PubMed]
  20. Dinulovic, I.; Furrer, R.; Handschin, C. Plasticity of the Muscle Stem Cell Microenvironment. Adv. Exp. Med. Biol. 2017, 1041, 141–169. [Google Scholar] [CrossRef]
  21. Dumont, N.A.; Wang, Y.X.; Rudnicki, M.A. Intrinsic and extrinsic mechanisms regulating satellite cell function. Development 2015, 142, 1572–1581. [Google Scholar] [CrossRef] [PubMed]
  22. Von Maltzahn, J.; Jones, A.E.; Parks, R.J.; Rudnicki, M.A. Pax7 is critical for the normal function of satellite cells in adult skeletal muscle. Proc. Natl. Acad. Sci. USA 2013, 110, 16474–16479. [Google Scholar] [CrossRef] [PubMed]
  23. Tierney, M.T.; Sacco, A. Satellite Cell Heterogeneity in Skeletal Muscle Homeostasis. Trends Cell Biol. 2016, 26, 434–444. [Google Scholar] [CrossRef] [PubMed]
  24. Kuang, S.; Kuroda, K.; Le Grand, F.; Rudnicki, M.A. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell 2007, 129, 999–1010. [Google Scholar] [CrossRef] [PubMed]
  25. Bentzinger, C.F.; Wang, Y.X.; Dumont, N.A.; Rudnicki, M.A. Cellular dynamics in the muscle satellite cell niche. EMBO Rep. 2013, 14, 1062–1072. [Google Scholar] [CrossRef] [PubMed]
  26. Urciuolo, A.; Quarta, M.; Morbidoni, V.; Gattazzo, F.; Molon, S.; Grumati, P.; Montemurro, F.; Tedesco, F.S.; Blaauw, B.; Cossu, G.; et al. Collagen VI regulates satellite cell self-renewal and muscle regeneration. Nat. Commun. 2013, 4, 1964. [Google Scholar] [CrossRef] [PubMed]
  27. Thomas, K.; Engler, A.J.; Meyer, G.A. Extracellular matrix regulation in the muscle satellite cell niche. Connect. Tissue Res. 2015, 56, 1–8. [Google Scholar] [CrossRef]
  28. Malecova, B.; Puri, P.L. “Mix of Mics”-Phenotypic and Biological Heterogeneity of “Multipotent” Muscle Interstitial Cells (MICs). J. Stem Cell Res. 2012, pii:004. [Google Scholar] [CrossRef]
  29. Ceafalan, L.C.; Popescu, B.O.; Hinescu, M.E. Cellular players in skeletal muscle regeneration. Biomed. Res. Int. 2014, 2014, 957014. [Google Scholar] [CrossRef]
  30. Čamernik, K.; Barlič, A.; Drobnič, M.; Marc, J.; Jeras, M.; Zupan, J. Mesenchymal Stem Cells in the Musculoskeletal System: From Animal Models to Human Tissue Regeneration? Stem Cell Rev. 2018, 14, 346–369. [Google Scholar] [CrossRef]
  31. Tonlorenzi, R.; Rossi, G.; Messina, G. Isolation and Characterization of Vessel-Associated Stem/Progenitor Cells from Skeletal Muscle. Methods Mol. Biol. 2017, 1556, 149–177. [Google Scholar] [CrossRef]
  32. Vezzani, B.; Pierantozzi, E.; Sorrentino, V. Not All Pericytes Are Born Equal: Pericytes from Human Adult Tissues Present Different Differentiation Properties. Stem Cells Dev. 2016, 25, 1549–1558. [Google Scholar] [CrossRef] [PubMed]
  33. Judson, R.N.; Low, M.; Eisner, C.; Rossi, F.M. Isolation, Culture, and Differentiation of Fibro/Adipogenic Progenitors (FAPs) from Skeletal Muscle. Methods Mol. Biol. 2017, 1668, 93–103. [Google Scholar] [CrossRef]
  34. Marini, M.; Rosa, I.; Ibba-Manneschi, L.; Manetti, M. Telocytes in skeletal, cardiac and smooth muscle interstitium: Morphological and functional aspects. Histol. Histopathol. 2018, 33, 1151–1165. [Google Scholar] [CrossRef] [PubMed]
  35. Doyle, M.J.; Zhou, S.; Tanaka, K.K.; Pisconti, A.; Farina, N.H.; Sorrentino, B.P.; Olwin, B.B. Abcg2 labels multiple cell types in skeletal muscle and participates in muscle regeneration. J. Cell Biol. 2011, 195, 147–163. [Google Scholar] [CrossRef] [PubMed]
  36. Tamaki, T.; Akatsuka, A.; Okada, Y.; Matsuzaki, Y.; Okano, H.; Kimura, M. Growth and differentiation potential of main- and side-population cells derived from murine skeletal muscle. Exp. Cell Res. 2003, 291, 83–90. [Google Scholar] [CrossRef]
  37. Mitchell, K.J.; Pannérec, A.; Cadot, B.; Parlakian, A.; Besson, V.; Gomes, E.R.; Marazzi, G.; Sassoon, D.A. Identification and characterization of a non-satellite cell muscle resident progenitor during postnatal development. Nat. Cell. Biol. 2010, 12, 257–266. [Google Scholar] [CrossRef] [PubMed]
  38. Farup, J.; Madaro, L.; Puri, P.L.; Mikkelsen, U.R. Interactions between muscle stem cells, mesenchymal-derived cells and immune cells in muscle homeostasis, regeneration and disease. Cell Death Dis. 2015, 6, e1830. [Google Scholar] [CrossRef] [PubMed]
  39. Tidball, J.G.; Villalta, S.A. Regulatory interactions between muscle and the immune system during muscle regeneration. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2010, 298, R1173–R1187. [Google Scholar] [CrossRef] [PubMed]
  40. Bencze, M.; Negroni, E.; Vallese, D.; Yacoub-Youssef, H.; Chaouch, S.; Wolff, A.; Aamiri, A.; Di Santo, J.P.; Chazaud, B.; Butler-Browne, G.; et al. Proinflammatory macrophages enhance the regenerative capacity of human myoblasts by modifying their kinetics of proliferation and differentiation. Mol. Ther. 2012, 20, 2168–2179. [Google Scholar] [CrossRef] [PubMed]
  41. Ceafalan, L.C.; Fertig, T.E.; Popescu, A.C.; Popescu, B.O.; Hinescu, M.E.; Gherghiceanu, M. Skeletal muscle regeneration involves macrophage-myoblast bonding. Cell Adhes. Migr. 2018, 12, 228–235. [Google Scholar] [CrossRef] [PubMed]
  42. Wang, X.; Zhao, W.; Ransohoff, R.M.; Zhou, L. Infiltrating macrophages are broadly activated at the early stage to support acute skeletal muscleinjury repair. J. Neuroimmunol. 2018, 317, 55–66. [Google Scholar] [CrossRef] [PubMed]
  43. Sonnet, C.; Lafuste, P.; Arnold, L.; Brigitte, M.; Poron, F.; Authier, F.J.; Chrétien, F.; Gherardi, R.K.; Chazaud, B. Human macrophages rescue myoblasts and myotubes from apoptosis through a set of adhesion molecular systems. J. Cell Sci. 2006, 119, 2497–2507. [Google Scholar] [CrossRef] [PubMed]
  44. Serrano, A.L.; Mann, C.J.; Vidal, B.; Ardite, E.; Perdiguero, E.; Muñoz-Cánoves, P. Cellular and molecular mechanisms regulating fibrosis in skeletal muscle repair and disease. Curr. Top. Dev. Biol. 2011, 96, 167–201. [Google Scholar] [CrossRef] [PubMed]
  45. Joe, A.W.; Yi, L.; Natarajan, A.; Le Grand, F.; So, L.; Wang, J.; Rudnicki, M.A.; Rossi, F.M. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nat. Cell Biol. 2010, 12, 153–163. [Google Scholar] [CrossRef] [PubMed]
  46. Murphy, M.M.; Lawson, J.A.; Mathew, S.J.; Hutcheson, D.A.; Kardon, G. Satellite cells, connective tissue fibroblasts and their interactions are crucial for muscle regeneration. Development 2011, 138, 3625–3637. [Google Scholar] [CrossRef] [PubMed]
  47. Heredia, J.E.; Mukundan, L.; Chen, F.M.; Mueller, A.A.; Deo, R.C.; Locksley, R.M.; Rando, T.A.; Chawla, A. Type 2 innate signals stimulate fibro/adipogenic progenitors to facilitate muscle regeneration. Cell 2013, 153, 376–388. [Google Scholar] [CrossRef] [PubMed]
  48. Mackey, A.L.; Magnan, M.; Chazaud, B.; Kjaer, M. Human skeletal muscle fibroblasts stimulate in vitro myogenesis and in vivo muscle regeneration. J. Physiol. 2017, 595, 5115–5127. [Google Scholar] [CrossRef] [PubMed]
  49. Malecova, B.; Gatto, S.; Etxaniz, U.; Passafaro, M.; Cortez, A.; Nicoletti, C.; Giordani, L.; Torcinaro, A.; De Bardi, M.; Bicciato, S.; et al. Dynamics of cellular states of fibro-adipogenic progenitors during myogenesis and muscular dystrophy. Nat. Commun. 2018, 9, 3670. [Google Scholar] [CrossRef] [PubMed]
  50. Cretoiu, S.M.; Popescu, L.M. Telocytes revisited. Biomol. Concepts 2014, 5, 353–369. [Google Scholar] [CrossRef] [PubMed]
  51. Nakamura, Y.; Miyaki, S.; Ishitobi, H.; Matsuyama, S.; Nakasa, T.; Kamei, N.; Akimoto, T.; Higashi, Y.; Ochi, M. Mesenchymal-stem-cell-derived exosomes accelerate skeletal muscle regeneration. FEBS Lett. 2015, 589, 1257–1265. [Google Scholar] [CrossRef] [PubMed]
  52. Christov, C.; Chrétien, F.; Abou-Khalil, R.; Bassez, G.; Vallet, G.; Authier, F.J.; Bassaglia, Y.; Shinin, V.; Tajbakhsh, S.; Chazaud, B.; et al. Muscle satellite cells and endothelial cells: Close neighbors and privileged partners. Mol. Biol. Cell 2007, 18, 1397–1409. [Google Scholar] [CrossRef] [PubMed]
  53. Verma, M.; Asakura, Y.; Murakonda, B.S.R.; Pengo, T.; Latroche, C.; Chazaud, B.; McLoon, L.K.; Asakura, A. Muscle Satellite Cell Cross-Talk with a Vascular Niche Maintains Quiescence via VEGF and Notch Signaling. Cell Stem Cell 2018, 23, 530–543.e9. [Google Scholar] [CrossRef] [PubMed]
  54. Abou-Khalil, R.; Mounier, R.; Chazaud, B. Regulation of myogenic stem cell behavior by vessel cells: The “menage a trois” of satellite cells, periendothelial cells and endothelial cells. Cell Cycle 2010, 9, 892–896. [Google Scholar] [CrossRef] [PubMed]
  55. Kostallari, E.; Baba-Amer, Y.; Alonso-Martin, S.; Ngoh, P.; Relaix, F.; Lafuste, P.; Gherardi, R.K. Pericytes in the myovascular niche promote post-natal myofiber growth and satellite cell quiescence. Development 2015, 142, 1242–1253. [Google Scholar] [CrossRef] [PubMed]
  56. Menetrey, J.; Kasemkijwattana, C.; Day, C.S.; Bosch, P.; Vogt, M.; Fu, F.H.; Moreland, M.S.; Huard, J. Growth factors improve muscle healing in vivo. J. Bone Jt. Surg. Br. 2000, 82, 131–137. [Google Scholar] [CrossRef]
  57. De Perini, A.; Dimauro, I.; Duranti, G.; Fantini, C.; Mercatelli, N.; Ceci, R.; Di Luigi, L.; Sabatini, S.; Caporossi, D. The p75NTR-mediated effect of nerve growth factor in L6C5 myogenic cells. BMC Res. Notes 2017, 10, 686. [Google Scholar] [CrossRef] [PubMed]
  58. Judson, R.N.; Zhang, R.H.; Rossi, F.M. Tissue-resident mesenchymal stem/progenitor cells in skeletal muscle: Collaborators or saboteurs? FEBS J. 2013, 280, 4100–4108. [Google Scholar] [CrossRef] [PubMed]
  59. Asakura, A.; Seale, P.; Girgis-Gabardo, A.; Rudnicki, M.A. Myogenic specification of side population cells in skeletal muscle. J. Cell Biol. 2002, 159, 123–134. [Google Scholar] [CrossRef] [PubMed]
  60. Uezumi, A.; Ojima, K.; Fukada, S.; Ikemoto, M.; Masuda, S.; Miyagoe-Suzuki, Y.; Takeda, S. Functional heterogeneity of side population cells in skeletal muscle. Biochem. Biophys. Res. Commun. 2006, 341, 864–873. [Google Scholar] [CrossRef]
  61. Tanaka, K.K.; Hall, J.K.; Troy, A.A.; Cornelison, D.D.; Majka, S.M.; Olwin, B.B. Syndecan-4-expressing muscle progenitor cells in the SP engraft as satellite cells during muscle regeneration. Cell Stem Cell 2009, 4, 217–225. [Google Scholar] [CrossRef] [PubMed]
  62. Dellavalle, A.; Maroli, G.; Covarello, D.; Azzoni, E.; Innocenzi, A.; Perani, L.; Antonini, S.; Sambasivan, R.; Brunelli, S.; Tajbakhsh, S.; et al. Pericytes resident in postnatal skeletal muscle differentiate into muscle fibres and generate satellite cells. Nat. Commun. 2011, 2, 499. [Google Scholar] [CrossRef] [PubMed]
  63. Díaz-Manera, J.; Gallardo, E.; de Luna, N.; Navas, M.; Soria, L.; Garibaldi, M.; Rojas-García, R.; Tonlorenzi, R.; Cossu, G.; Illa, I. The increase of pericyte population in human neuromuscular disorders supports their role in muscle regeneration in vivo. J. Pathol. 2012, 228, 544–553. [Google Scholar] [CrossRef] [PubMed]
  64. Birbrair, A.; Zhang, T.; Wang, Z.M.; Messi, M.L.; Enikolopov, G.N.; Mintz, A.; Delbono, O. Skeletal muscle pericyte subtypes differ in their differentiation potential. Stem Cell Res. 2013, 10, 67–84. [Google Scholar] [CrossRef] [PubMed]
  65. Benchaouir, R.; Meregalli, M.; Farini, A.; D’Antona, G.; Belicchi, M.; Goyenvalle, A.; Battistelli, M.; Bresolin, N.; Bottinelli, R.; Garcia, L.; et al. Restoration of human dystrophin following transplantation of exon-skipping-engineered DMD patient stem cells into dystrophic mice. Cell Stem Cell 2007, 1, 646–657. [Google Scholar] [CrossRef] [PubMed]
  66. Negroni, E.; Riederer, I.; Chaouch, S.; Belicchi, M.; Razini, P.; Di Santo, J.; Torrente, Y.; Butler-Browne, G.S.; Mouly, V. In vivo myogenic potential of human CD133+ muscle-derived stem cells: A quantitative study. Mol. Ther. 2009, 17, 1771–1778. [Google Scholar] [CrossRef] [PubMed]
  67. Meng, J.; Chun, S.; Asfahani, R.; Lochmüller, H.; Muntoni, F.; Morgan, J. Human skeletal muscle-derived CD133(+) cells form functional satellite cells after intramuscular transplantation in immunodeficient host mice. Mol. Ther. 2014, 22, 1008–1017. [Google Scholar] [CrossRef]
  68. Dulauroy, S.; Di Carlo, S.E.; Langa, F.; Eberl, G.; Peduto, L. Lineage tracing and genetic ablation of ADAM12(+) perivascular cells identify a major source of profibrotic cells during acute tissue injury. Nat. Med. 2012, 18, 1262–1270. [Google Scholar] [CrossRef]
  69. Uezumi, A.; Ikemoto-Uezumi, M.; Tsuchida, K. Roles of non myogenic mesenchymal progenitors in pathogenesis and regeneration of skeletal muscle. Front. Physiol. 2014, 5, 68. [Google Scholar] [CrossRef]
  70. Lemos, D.R.; Babaeijandaghi, F.; Low, M.; Chang, C.K.; Lee, S.T.; Fiore, D.; Zhang, R.H.; Natarajan, A.; Nedospasov, S.A.; Rossi, F.M. Nilotinib reduces muscle fibrosis in chronic muscle injury by promoting TNF-mediated apoptosis of fibro/adipogenic progenitors. Nat. Med. 2015, 21, 786–794. [Google Scholar] [CrossRef]
  71. Rudolf, R.; Deschenes, M.R.; Sandri, M. Neuromuscular junction degeneration in muscle wasting. Curr. Opin. Clin. Nutr. Metab. Care 2016, 19, 177–181. [Google Scholar] [CrossRef] [PubMed]
  72. DeLong, J.M.; Russell, R.P.; Mazzocca, A.D. Platelet-rich plasma: The PAW classification system. Arthroscopy 2012, 28, 998–1009. [Google Scholar] [CrossRef] [PubMed]
  73. De Pascale, M.R.; Sommese, L.; Casamassimi, A.; Napoli, C. Platelet derivatives in regenerative medicine: An update. Transfus. Med. Rev. 2015, 29, 52–61. [Google Scholar] [CrossRef] [PubMed]
  74. Mautner, K.; Malanga, G.A.; Smith, J.; Shiple, B.; Ibrahim, V.; Sampson, S.; Bowen, J.E. A call for a standard classification system for future biologic research: The rationale for new PRP nomenclature. PM R 2015, 7, S53–S59. [Google Scholar] [CrossRef] [PubMed]
  75. Le, A.D.K.; Enweze, L.; DeBaun, M.R.; Dragoo, J.L. Platelet-Rich Plasma. Clin. Sports Med. 2019, 38, 17–44. [Google Scholar] [CrossRef] [PubMed]
  76. Cervelli, V.; Gentile, P.; Scioli, M.G.; Grimaldi, M.; Casciani, C.U.; Spagnoli, L.G.; Orlandi, A. Application of platelet-rich plasma in plastic surgery: Clinical and in vitro evaluation. Tissue Eng. Part C Methods 2009, 15, 625–634. [Google Scholar] [CrossRef] [PubMed]
  77. Agrawal, A.A. Evolution, current status and advances in application of platelet concentrate in periodontics and implantology. World J. Clin. Cases 2017, 5, 159–171. [Google Scholar] [CrossRef]
  78. Giannaccare, G.; Versura, P.; Buzzi, M.; Primavera, L.; Pellegrini, M.; Campos, E.C. Blood derived eye drops for the treatment of cornea and ocular surface diseases. Transfus. Apher. Sci. 2017, 56, 595–604. [Google Scholar] [CrossRef]
  79. Tandulwadkar, S.R.; Naralkar, M.V.; Surana, A.D.; Selvakarthick, M.; Kharat, A.H. Autologous intrauterine platelet-rich plasma instillation for suboptimal endometrium in frozen embryo transfer cycles: A pilot study. J. Hum. Reprod. Sci. 2017, 10, 208–212. [Google Scholar] [CrossRef]
  80. Santos, S.C.N.D.S.; Sigurjonsson, O.E.; Custodio, C.A.; Mano, J.F.C.D.L. Blood plasma derivatives for tissue engineering and regenerative medicine therapies. Tissue Eng. Part B Rev. 2018, 24, 454–462. [Google Scholar] [CrossRef]
  81. Chicharro-Alcántara, D.; Rubio-Zaragoza, M.; Damiá-Giménez, E.; Carrillo-Poveda, J.M.; Cuervo-Serrato, B.; Peláez-Gorrea, P.; Sopena-Juncosa, J.J. Platelet rich plasma: New insights for cutaneous wound healing management. J. Funct. Biomater. 2018, 9, 10. [Google Scholar] [CrossRef] [PubMed]
  82. Nguyen, R.T.; Borg-Stein, J.; McInnis, K. Applications of platelet-rich plasma in musculoskeletal and sports medicine: An evidence-based approach. PM R 2011, 3, 226–250. [Google Scholar] [CrossRef] [PubMed]
  83. Andia, I.; Abate, M. Platelet-rich plasma in the treatment of skeletal muscle injuries. Expert. Opin. Biol. 2015, 15, 987–999. [Google Scholar] [CrossRef] [PubMed]
  84. Andia, I.; Abate, M. Platelet-rich plasma: Combinational treatment modalities for musculoskeletal conditions. Front. Med. 2018, 12, 139–152. [Google Scholar] [CrossRef] [PubMed]
  85. Andia, I.; Martin, J.I.; Maffulli, N. Advances with platelet rich plasma therapies for tendon regeneration. Expert Opin. Biol. 2018, 18, 389–398. [Google Scholar] [CrossRef] [PubMed]
  86. Amable, P.R.; Carias, R.B.; Teixeira, M.V.; da Cruz Pacheco, I.; Corrêa do Amaral, R.J.; Granjeiro, J.M.; Borojevic, R. Platelet-rich plasma preparation for regenerative medicine: Optimization and quantification of cytokines and growth factors. Stem Cell Res. 2013, 4, 67. [Google Scholar] [CrossRef] [PubMed]
  87. Martínez, C.E.; Smith, P.C.; Palma Alvarado, V.A. The influence of platelet-derived products on angiogenesis and tissue repair: A concise update. Front. Physiol. 2015, 6, 290. [Google Scholar] [CrossRef] [PubMed]
  88. Pochini, A.C.; Antonioli, E.; Bucci, D.Z.; Sardinha, L.R.; Andreoli, C.V.; Ferretti, M.; Ejnisman, B.; Goldberg, A.C.; Cohen, M. Analysis of cytokine profile and growth factors in platelet-rich plasma obtained by open systems and commercial columns. Einstein 2016, 14, 391–397. [Google Scholar] [CrossRef] [PubMed]
  89. Qiao, J.; An, N.; Ouyang, X. Quantification of growth factors in different platelet concentrates. Platelets 2017, 28, 774–778. [Google Scholar] [CrossRef]
  90. San Sebastian, K.M.; Lobato, I.; Hernández, I.; Burgos-Alonso, N.; Gomez-Fernandez, M.C.; López, J.L.; Rodríguez, B.; March, A.G.; Grandes, G.; Andia, I. Efficacy and safety of autologous platelet rich plasma for the treatment of vascular ulcers in primary care: Phase III study. BMC Fam. Pract. 2014, 15, 211. [Google Scholar] [CrossRef]
  91. Suthar, M.; Gupta, S.; Bukhari, S.; Ponemone, V. Treatment of chronic non-healing ulcers using autologous platelet rich plasma: A case series. J. Biomed. Sci. 2017, 24, 16. [Google Scholar] [CrossRef] [PubMed]
  92. Bottegoni, C.; Farinelli, L.; Aquili, A.; Chiurazzi, E.; Gigante, A. Homologous platelet-rich plasma for the treatment of knee involvement in primary Sjögren’s syndrome. J. Biol. Regul. Homeost. Agents 2016, 30, 63–67. [Google Scholar] [PubMed]
  93. Anitua, E.; Prado, R.; Orive, G. Allogeneic Platelet-Rich Plasma: At the Dawn of an Off-the-Shelf Therapy? Trends Biotechnol. 2017, 35, 91–93. [Google Scholar] [CrossRef] [PubMed]
  94. Bubnov, R.; Yevseenko, V.; Semeniv, I. Ultrasound guided injections of platelets rich plasma for muscle injury in professional athletes. Comparative study. Med. Ultrason. 2013, 15, 101–105. [Google Scholar] [CrossRef] [PubMed]
  95. Wetzel, R.J.; Patel, R.M.; Terry, M.A. Platelet-rich plasma as an effective treatment for proximal hamstring injuries. Orthopedics 2013, 36, e64–e70. [Google Scholar] [CrossRef] [PubMed]
  96. A Hamid, M.S.; Mohamed Ali, M.R.; Yusof, A.; George, J.; Lee, L.P. Platelet-rich plasma injections for the treatment of hamstring injuries: A randomized controlled trial. Am. J. Sports Med. 2014, 42, 2410–2418. [Google Scholar] [CrossRef] [PubMed]
  97. Rossi, L.A.; Molina Rómoli, A.R.; Bertona Altieri, B.A.; Burgos Flor, J.A.; Scordo, W.E.; Elizondo, C.M. Does platelet-rich plasma decrease time to return to sports in acute muscle tear? A randomized controlled trial. Knee Surg. Sports Traumatol. Arthrosc. 2017, 25, 3319–3325. [Google Scholar] [CrossRef]
  98. Borrione, P.; Fossati, C.; Pereira, M.T.; Giannini, S.; Davico, M.; Minganti, C.; Pigozzi, F. The use of platelet-rich plasma (PRP) in the treatment of gastrocnemius strains: A retrospective observational study. Platelets 2018, 29, 596–601. [Google Scholar] [CrossRef]
  99. Bernuzzi, G.; Petraglia, F.; Pedrini, M.F.; De Filippo, M.; Pogliacomi, F.; Verdano, M.A.; Costantino, C. Use of platelet-rich plasma in the care of sports injuries: Our experience with ultrasound-guided injection. Blood Transfus. 2014, 12, s229–s234. [Google Scholar] [CrossRef]
  100. Zanon, G.; Combi, F.; Combi, A.; Perticarini, L.; Sammarchi, L.; Benazzo, F. Platelet-rich plasma in the treatment of acute hamstring injuries in professional football players. Joints 2016, 4, 17–23. [Google Scholar] [CrossRef]
  101. Punduk, Z.; Oral, O.; Ozkayin, N.; Rahman, K.; Varol, R. Single dose of intra-muscular platelet rich plasma reverses the increase in plasma iron levels in exercise-induced muscle damage: A pilot study. J. Sport Health Sci. 2016, 5, 109–114. [Google Scholar] [CrossRef] [PubMed]
  102. Sheth, U.; Dwyer, T.; Smith, I.; Wasserstein, D.; Theodoropoulos, J.; Takhar, S.; Chahal, J. Does Platelet-Rich Plasma Lead to Earlier Return to Sport When Compared With Conservative Treatment in Acute Muscle Injuries? A Systematic Review and Meta-analysis. Arthroscopy 2018, 34, 281–288.e1. [Google Scholar] [CrossRef] [PubMed]
  103. Scully, D.; Naseem, K.M.; Matsakas, A. Platelet biology in regenerative medicine of skeletal muscle. Acta Physiol. (Oxf.) 2018, 223, e13071. [Google Scholar] [CrossRef] [PubMed]
  104. Grassi, A.; Napoli, F.; Romandini, I.; Samuelsson, K.; Zaffagnini, S.; Candrian, C.; Filardo, G. Is Platelet-Rich Plasma (PRP) Effective in the Treatment of Acute Muscle Injuries? A Systematic Review and Meta-Analysis. Sports Med. 2018, 48, 971–989. [Google Scholar] [CrossRef] [PubMed]
  105. Borrione, P.; Grasso, L.; Chierto, E.; Geuna, S.; Racca, S.; Abbadessa, G.; Ronchi, G.; Faiola, F.; Di Gianfrancesco, A.; Pigozzi, F. Experimental model for the study of the effects of platelet-rich plasma on the early phases of muscle healing. Blood Transfus. 2014, 12, s221–s228. [Google Scholar] [CrossRef] [PubMed]
  106. Li, H.; Hicks, J.J.; Wang, L.; Oyster, N.; Philippon, M.J.; Hurwitz, S.; Hogan, M.V.; Huard, J. Customized platelet-rich plasma with transforming growth factor β1 neutralization antibody to reduce fibrosis in skeletal muscle. Biomaterials 2016, 87, 147–156. [Google Scholar] [CrossRef] [PubMed]
  107. Garcia, T.A.; Camargo, R.C.T.; Koike, T.E.; Ozaki, G.A.T.; Castoldi, R.C.; Camargo Filho, J.C.S. Histological analysis of the association of low level laser therapy and platelet-rich plasma in regeneration of muscle injury in rats. Braz. J. Phys. 2017, 21, 425–433. [Google Scholar] [CrossRef] [PubMed]
  108. Tsai, W.C.; Yu, T.Y.; Chang, G.J.; Lin, L.P.; Lin, M.S.; Pang, J.S. Platelet-Rich Plasma Releasate Promotes Regeneration and Decreases Inflammation and Apoptosis of Injured Skeletal Muscle. Am. J. Sports Med. 2018, 46, 1980–1986. [Google Scholar] [CrossRef] [PubMed]
  109. Hammond, J.W.; Hinton, R.Y.; Curl, L.A.; Muriel, J.M.; Lovering, R.M. Use of autologous platelet-rich plasma to treat muscle strain injuries. Am. J. Sports Med. 2009, 37, 1135–1142. [Google Scholar] [CrossRef] [PubMed]
  110. Gigante, A.; Del Torto, M.; Manzotti, S.; Cianforlini, M.; Busilacchi, A.; Davidson, P.A.; Greco, F.; Mattioli-Belmonte, M. Platelet rich fibrin matrix effects on skeletal muscle lesions: An experimental study. J. Biol. Regul. Homeost. Agents 2012, 26, 475–484. [Google Scholar] [PubMed]
  111. Dimauro, I.; Grasso, L.; Fittipaldi, S.; Fantini, C.; Mercatelli, N.; Racca, S.; Geuna, S.; Di Gianfrancesco, A.; Caporossi, D.; Pigozzi, F.; et al. Platelet-rich plasma and skeletal muscle healing: A molecular analysis of the early phases of the regeneration process in an experimental animal model. PLoS ONE 2014, 9, e102993. [Google Scholar] [CrossRef] [PubMed]
  112. Cianforlini, M.; Mattioli-Belmonte, M.; Manzotti, S.; Chiurazzi, E.; Piani, M.; Orlando, F.; Provinciali, M.; Gigante, A. Effect of platelet rich plasma concentration on skeletal muscle regeneration: An experimental study. J. Biol. Regul. Homeost. Agents 2015, 29, 47–55. [Google Scholar] [PubMed]
  113. Contreras-Muñoz, P.; Torrella, J.R.; Serres, X.; Rizo-Roca, D.; De la Varga, M.; Viscor, G.; Martínez-Ibáñez, V.; Peiró, J.L.; Järvinen, T.A.H.; Rodas, G.; et al. Postinjury Exercise and Platelet-Rich Plasma Therapies Improve Skeletal Muscle Healing in Rats But Are Not Synergistic When Combined. Am. J. Sports Med. 2017, 45, 2131–2141. [Google Scholar] [CrossRef] [PubMed]
  114. Martins, R.P.; Hartmann, D.D.; de Moraes, J.P.; Soares, F.A.; Puntel, G.O. Platelet-rich plasma reduces the oxidative damage determined by a skeletal muscle contusion in rats. Platelets 2016, 27, 784–790. [Google Scholar] [CrossRef] [PubMed]
  115. Cunha, R.C.; Francisco, J.C.; Cardoso, M.A.; Matos, L.F.; Lino, D.; Simeoni, R.B.; Pereira, G.; Irioda, A.C.; Simeoni, P.R.; Guarita-Souza, L.C.; et al. Effect of platelet-rich plasma therapy associated with exercise training in musculoskeletal healing in rats. Transpl. Proc. 2014, 46, 1879–1881. [Google Scholar] [CrossRef] [PubMed]
  116. Anitua, E.; Pelacho, B.; Prado, R.; Aguirre, J.J.; Sánchez, M.; Padilla, S.; Aranguren, X.L.; Abizanda, G.; Collantes, M.; Hernandez, M.; et al. Infiltration of plasma rich in growth factors enhances in vivo angiogenesis and improves reperfusion and tissue remodeling after severe hind limb ischemia. J. Control. Release 2015, 202, 31–39. [Google Scholar] [CrossRef] [PubMed]
  117. Terada, S.; Ota, S.; Kobayashi, M.; Kobayashi, T.; Mifune, Y.; Takayama, K.; Witt, M.; Vadalà, G.; Oyster, N.; Otsuka, T.; et al. Use of an antifibrotic agent improves the effect of platelet-rich plasma on muscle healing after injury. J. Bone Jt. Surg. Am. 2013, 95, 980–988. [Google Scholar] [CrossRef] [PubMed]
  118. Denapoli, P.M.; Stilhano, R.S.; Ingham, S.J.; Han, S.W.; Abdalla, R.J. Platelet-Rich Plasma in a Murine Model: Leukocytes, Growth Factors, Flt-1, and Muscle Healing. Am. J. Sports Med. 2016, 44, 1962–1971. [Google Scholar] [CrossRef]
  119. Pinheiro, C.L.; Peixinho, C.C.; Esposito, C.C.; Manso, J.E.; Machado, J.C. Ultrasound biomicroscopy and claudication test for in vivo follow-up of muscle repair enhancement based on platelet-rich plasma therapy in a rat model of gastrocnemius laceration. Acta Cir. Bras. 2016, 31, 103–110. [Google Scholar] [CrossRef] [PubMed]
  120. Takase, F.; Inui, A.; Mifune, Y.; Sakata, R.; Muto, T.; Harada, Y.; Ueda, Y.; Kokubu, T.; Kurosaka, M. Effect of platelet-rich plasma on degeneration change of rotator cuff muscles: In vitro and in vivo evaluations. J. Orthop. Res. 2017, 35, 1806–1815. [Google Scholar] [CrossRef] [PubMed]
  121. Miroshnychenko, O.; Chang, W.T.; Dragoo, J.L. The Use of Platelet-Rich and Platelet-Poor Plasma to Enhance Differentiation of Skeletal Myoblasts: Implications for the Use of Autologous Blood Products for Muscle Regeneration. Am. J. Sports Med. 2017, 45, 945–953. [Google Scholar] [CrossRef] [PubMed]
  122. Li, H.; Usas, A.; Poddar, M.; Chen, C.W.; Thompson, S.; Ahani, B.; Cummins, J.; Lavasani, M.; Huard, J. Platelet-rich plasma promotes the proliferation of human muscle derived progenitor cells and maintains their stemness. PLoS ONE 2013, 8, e64923. [Google Scholar] [CrossRef] [PubMed]
  123. Im, W.; Ban, J.J.; Lim, J.; Lee, M.; Chung, J.Y.; Bhattacharya, R.; Kim, S.H. Adipose-derived stem cells extract has a proliferative effect on myogenic progenitors. In Vitro Cell. Dev. Biol. Anim. 2014, 50, 740–766. [Google Scholar] [CrossRef] [PubMed]
  124. Tsai, W.C.; Yu, T.Y.; Lin, L.P.; Lin, M.S.; Wu, Y.C.; Liao, C.H.; Pang, J.S. Platelet rich plasma releasate promotes proliferation of skeletal muscle cells in association with upregulation of PCNA, cyclins and cyclin dependent kinases. Platelets 2017, 28, 491–497. [Google Scholar] [CrossRef] [PubMed]
  125. McClure, M.J.; Garg, K.; Simpson, D.G.; Ryan, J.J.; Sell, S.A.; Bowlin, G.L.; Ericksen, J.J. The influence of platelet-rich plasma on myogenic differentiation. J. Tissue Eng. Regen. Med. 2016, 10, E239–E249. [Google Scholar] [CrossRef] [PubMed]
  126. McClure, M.J.; Clark, N.M.; Schwartz, Z.; Boyan, B.D. Platelet-rich plasma and alignment enhance myogenin via ERK mitogen activated protein kinase signaling. Biomed. Mater. 2018, 13, 055009. [Google Scholar] [CrossRef] [PubMed]
  127. Kelc, R.; Trapecar, M.; Gradisnik, L.; Rupnik, M.S.; Vogrin, M. Platelet-rich plasma, especially when combined with a TGF-β inhibitor promotes proliferation, viability and myogenic differentiation of myoblasts in vitro. PLoS ONE 2015, 10, e0117302. [Google Scholar] [CrossRef]
  128. Deasy, B.M.; Feduska, J.M.; Payne, T.R.; Li, Y.; Ambrosio, F.; Huard, J. Effect of VEGF on the regenerative capacity of muscle stem cells in dystrophic skeletal muscle. Mol. Ther. 2009, 17, 1788–1798. [Google Scholar] [CrossRef]
  129. Walker, N.; Kahamba, T.; Woudberg, N.; Goetsch, K.; Niesler, C. Dose-dependent modulation of myogenesis by HGF: Implications for c-Met expression and downstream signalling pathways. Growth Factors 2015, 33, 229–241. [Google Scholar] [CrossRef]
  130. Duan, C.; Ren, H.; Gao, S. Insulin-like growth factors (IGFs), IGF receptors, and IGF-binding proteins: Roles in skeletal muscle growth and differentiation. Gen. Comp. Endocrinol. 2010, 167, 344–351. [Google Scholar] [CrossRef]
  131. Scully, D.; Sfyri, P.; Verpoorten, S.; Papadopoulos, P.; Muñoz-Turrillas, M.C.; Mitchell, R.; Aburima, A.; Patel, K.; Gutiérrez, L.; Naseem, K.M.; et al. Platelet releasate promotes skeletal myogenesis by increasing muscle stem cell commitment to differentiation and accelerates muscle regeneration following acute injury. Acta Physiol. (Oxf.) 2018, 19, e13207. [Google Scholar] [CrossRef] [PubMed]
  132. Tsai, W.C.; Yu, T.Y.; Lin, L.P.; Lin, M.S.; Tsai, T.T.; Pang, J.S. Platelet rich plasma promotes skeletal muscle cell migration in association with up-regulation of FAK, paxillin, and F-Actin formation. J. Orthop. Res. 2017, 35, 2506–2512. [Google Scholar] [CrossRef] [PubMed]
  133. Sassoli, C.; Vallone, L.; Tani, A.; Chellini, F.; Nosi, D.; Zecchi-Orlandini, S. Combined use of bone marrow-derived mesenchymal stromal cells (BM-MSCs) and platelet rich plasma (PRP) stimulates proliferation and differentiation of myoblasts in vitro: New therapeutic perspectives for skeletal muscle repair/regeneration. Cell Tissue Res. 2018, 372, 549–570. [Google Scholar] [CrossRef] [PubMed]
  134. Yamada, M.; Sankoda, Y.; Tatsumi, R.; Mizunoya, W.; Ikeuchi, Y.; Sunagawa, K.; Allen, R.E. Matrix metalloproteinase-2 mediates stretch-induced activation of skeletal muscle satellite cells in a nitric oxide-dependent manner. Int. J. Biochem. Cell Biol. 2008, 40, 2183–2191. [Google Scholar] [CrossRef] [PubMed]
  135. Pallafacchina, G.; François, S.; Regnault, B.; Czarny, B.; Dive, V.; Cumano, A.; Montarras, D.; Buckingham, M. An adult tissue-specific stem cell in its niche: A gene profiling analysis of in vivo quiescent and activated muscle satellite cells. Stem Cell Res. 2010, 4, 77–91. [Google Scholar] [CrossRef] [PubMed]
  136. Bellayr, I.; Holden, K.; Mu, X.; Pan, H.; Li, Y. Matrix metalloproteinase inhibition negatively affects muscle stem cell behavior. Int. J. Clin. Exp. Pathol. 2013, 6, 124–141. [Google Scholar] [PubMed]
  137. Chen, X.; Li, Y. Role of matrix metalloproteinases in skeletal muscle: Migration, differentiation, regeneration and fibrosis. Cell Adhes. Migr. 2009, 3, 337–341. [Google Scholar] [CrossRef]
  138. Miyazaki, D.; Nakamura, A.; Fukushima, K.; Yoshida, K.; Takeda, S.; Ikeda, S. Matrix metalloproteinase-2 ablation in dystrophin-deficient mdx muscles reduces angiogenesis resulting in impaired growth of regenerated muscle fibers. Hum. Mol. Genet. 2011, 20, 1787–1799. [Google Scholar] [CrossRef] [PubMed]
  139. Cassano, M.; Dellavalle, A.; Tedesco, F.S.; Quattrocelli, M.; Crippa, S.; Ronzoni, F.; Salvade, A.; Berardi, E.; Torrente, Y.; Cossu, G.; et al. Alpha sarcoglycan is required for FGF-dependent myogenic progenitor cell proliferation in vitro and in vivo. Development 2011, 138, 4523–4533. [Google Scholar] [CrossRef]
  140. Sartori, R.; Gregorevic, P.; Sandri, M. TGFβ and BMP signaling in skeletal muscle: Potential significance for muscle-related disease. Trends Endocrinol. Metab. 2014, 25, 464–471. [Google Scholar] [CrossRef] [PubMed]
  141. Escobar, G.; Escobar, A.; Ascui, G.; Tempio, F.I.; Ortiz, M.C.; Pérez, C.A.; López, M.N. Pure platelet-rich plasma and supernatant of calcium-activated P-PRP induce different phenotypes of human macrophages. Regen. Med. 2018, 13, 427–441. [Google Scholar] [CrossRef] [PubMed]
  142. Papait, A.; Cancedda, R.; Mastrogiacomo, M.; Poggi, A. Allogeneic platelet-rich plasma affects monocyte differentiation to dendritic cells causing an anti-inflammatory microenvironment, putatively fostering wound healing. J. Tissue Eng. Regen. Med. 2018, 12, 30–43. [Google Scholar] [CrossRef] [PubMed]
  143. Pakshir, P.; Hinz, B. The big five in fibrosis: Macrophages, myofibroblasts, matrix, mechanics, and miscommunication. Matrix Biol. 2018, 68–69, 81–93. [Google Scholar] [CrossRef] [PubMed]
  144. Chellini, F.; Tani, A.; Vallone, L.; Nosi, D.; Pavan, P.; Bambi, F.; Zecchi Orlandini, S.; Sassoli, C. Platelet-Rich Plasma Prevents In Vitro Transforming Growth Factor-β1-Induced Fibroblast to Myofibroblast Transition: Involvement of Vascular Endothelial Growth Factor (VEGF)-A/VEGF Receptor-1-Mediated Signaling. Cells 2018, 7, 142. [Google Scholar] [CrossRef] [PubMed]
  145. Anitua, E.; Troya, M.; Orive, G. Plasma rich in growth factors promote gingival tissue regeneration by stimulating fibroblast proliferation and migration and by blocking transforming growth factor-β1-induced myodifferentiation. J. Periodontol. 2012, 83, 1028–1037. [Google Scholar] [CrossRef] [PubMed]
  146. Anitua, E.; de la Fuente, M.; Muruzabal, F.; Riestra, A.; Merayo-Lloves, J.; Orive, G. Plasma rich in growth factors (PRGF) eye drops stimulates scarless regeneration compared to autologous serum in the ocular surface stromal fibroblasts. Exp. Eye Res. 2015, 135, 118–126. [Google Scholar] [CrossRef] [PubMed]
  147. Wang, M.L.; Rivlin, M.; Graham, J.G.; Beredjiklian, P.K. Peripheral nerve injury, scarring, and recovery. Connect. Tissue Res. 2018, 6, 1–7. [Google Scholar] [CrossRef] [PubMed]
  148. Giannessi, E.; Coli, A.; Stornelli, M.R.; Miragliotta, V.; Pirone, A.; Lenzi, C.; Burchielli, S.; Vozzi, G.; De Maria, C.; Giorgetti, M. An autologously generated platelet-rich plasma suturable membrane may enhance peripheral nerve regeneration after neurorraphy in an acute injury model of sciatic nerve neurotmesis. J. Reconstr. Microsurg. 2014, 30, 617–626. [Google Scholar] [CrossRef]
  149. Zheng, C.; Zhu, Q.; Liu, X.; Huang, X.; He, C.; Jiang, L.; Quan, D.; Zhou, X.; Zhu, Z. Effect of platelet-rich plasma (PRP) concentration on proliferation, neurotrophic function and migration of Schwann cells in vitro. J. Tissue Eng. Regen. Med. 2016, 10, 428–436. [Google Scholar] [CrossRef]
  150. Sánchez, M.; Garate, A.; Delgado, D.; Padilla, S. Platelet-rich plasma, an adjuvant biological therapy to assist peripheral nerve repair. Neural Regen. Res. 2017, 12, 47–52. [Google Scholar] [CrossRef]
  151. Teymur, H.; Tiftikcioglu, Y.O.; Cavusoglu, T.; Tiftikcioglu, B.I.; Erbas, O.; Yigitturk, G.; Uyanikgil, Y. Effect of platelet-rich plasma on reconstruction with nerve autografts. Kaohsiung J. Med. Sci. 2017, 33, 69–77. [Google Scholar] [CrossRef] [PubMed]
  152. Cáceres, M.; Martínez, C.; Martínez, J.; Smith, P.C. Effects of platelet-rich and -poor plasma on the reparative response of gingival fibroblasts. Clin. Oral Implant. Res. 2012, 23, 1104–1111. [Google Scholar] [CrossRef] [PubMed]
  153. Creeper, F.; Ivanovski, S. Effect of autologous and allogenic platelet-rich plasma on human gingival fibroblast function. Oral Dis. 2012, 18, 494–500. [Google Scholar] [CrossRef] [PubMed]
  154. Martínez, C.E.; González, S.A.; Palma, V.; Smith, P.C. Platelet-Poor and Platelet-Rich Plasma Stimulate Bone Lineage Differentiation in Periodontal Ligament Stem Cells. J. Periodontol. 2016, 87, e18–e26. [Google Scholar] [CrossRef] [PubMed]
  155. de Mos, M.; van der Windt, A.E.; Jahr, H.; van Schie, H.T.; Weinans, H.; Verhaar, J.A.; van Osch, G.J. Can platelet-rich plasma enhance tendon repair? A cell culture study. Am. J. Sports Med. 2008, 36, 1171–1178. [Google Scholar] [CrossRef] [PubMed]
  156. Shahidi, M.; Vatanmakanian, M.; Arami, M.K.; Sadeghi Shirazi, F.; Esmaeili, N.; Hydarporian, S.; Jafari, S. A comparative study between platelet-rich plasma and platelet-poor plasma effects on angiogenesis. Med. Mol. Morphol. 2018, 51, 21–31. [Google Scholar] [CrossRef] [PubMed]
  157. Renn, T.Y.; Kao, Y.H.; Wang, C.C.; Burnouf, T. Anti-inflammatory effects of platelet biomaterials in a macrophage cellular model. Vox Sang. 2015, 109, 138–147. [Google Scholar] [CrossRef] [PubMed]
  158. Rettig, A.C.; Meyer, S.; Bhadra, A.K. Platelet-Rich Plasma in Addition to Rehabilitation for Acute Hamstring Injuries in NFL Players: Clinical Effects and Time to Return to Play. Orthop. J. Sports Med. 2013, 1, 2325967113494354. [Google Scholar] [CrossRef] [PubMed]
  159. Delos, D.; Leineweber, M.J.; Chaudhury, S.; Alzoobaee, S.; Gao, Y.; Rodeo, S.A. The effect of platelet-rich plasma on muscle contusion healing in a rat model. Am. J. Sports Med. 2014, 42, 2067–2074. [Google Scholar] [CrossRef] [PubMed]
  160. Hamid, M.S.; Yusof, A.; Mohamed Ali, M.R. Platelet-rich plasma (PRP) for acute muscle injury: A systematic review. PLoS ONE 2014, 9, e90538. [Google Scholar] [CrossRef]
  161. Hamilton, B.; Tol, J.L.; Almusa, E.; Boukarroum, S.; Eirale, C.; Farooq, A.; Whiteley, R.; Chalabi, H. Platelet-rich plasma does not enhance return to play in hamstring injuries: A randomised controlled trial. Br. J. Sports Med. 2015, 49, 943–950. [Google Scholar] [CrossRef] [PubMed]
  162. Reurink, G.; Goudswaard, G.J.; Moen, M.H.; Weir, A.; Verhaar, J.A.; Bierma-Zeinstra, S.M.; Maas, M.; Tol, J.L. Dutch HIT-study Investigators. Rationale, secondary outcome scores and 1-year follow-up of a randomised trial of platelet-rich plasma injections in acute hamstring muscle injury: The Dutch Hamstring Injection Therapy study. Br. J. Sports Med. 2015, 49, 1206–1212. [Google Scholar] [CrossRef] [PubMed]
  163. Kelc, R.; Vogrin, M. Concerns about fibrosis development after scaffolded PRP therapy of muscle injuries: Commentary on an article by Sanchez et al.: Muscle repair: Platelet-rich plasma derivates as a bridge from spontaneity to intervention. Injury 2015, 46, 428. [Google Scholar] [CrossRef] [PubMed]
  164. Martinez-Zapata, M.J.; Orozco, L.; Balius, R.; Soler, R.; Bosch, A.; Rodas, G.; Til, L.; Peirau, X.; Urrútia, G.; Gich, I.; et al. PRP-RICE group. Efficacy of autologous platelet-rich plasma for the treatment of muscle rupture with haematoma: A multicentre, randomised, double-blind, placebo-controlled clinical trial. Blood Transfus. 2016, 14, 245–254. [Google Scholar] [CrossRef] [PubMed]
  165. Guillodo, Y.; Madouas, G.; Simon, T.; Le Dauphin, H.; Saraux, A. Platelet-rich plasma (PRP) treatment of sports-related severe acute hamstring injuries. Muscles Ligaments Tendons J. 2016, 5, 284–288. [Google Scholar] [CrossRef] [PubMed]
  166. Navani, A.; Li, G.; Chrystal, J. Platelet rich plasma in musculoskeletal pathology: A necessary rescue or a lost cause? Pain Physician 2017, 20, E345–E356. [Google Scholar] [PubMed]
  167. Tonogai, I.; Hayashi, F.; Iwame, T.; Takasago, T.; Matsuura, T.; Sairyo, K. Platelet-rich plasma does not reduce skeletal muscle fibrosis after distraction osteogenesis. J. Exp. Orthop. 2018, 5, 26. [Google Scholar] [CrossRef]
  168. Manduca, M.L.; Straub, S.J. Effectiveness of PRP Injection in Reducing Recovery Time of Acute Hamstring Injury: A Critically Appraised Topic. J. Sport Rehabil. 2018, 27, 480–484. [Google Scholar] [CrossRef]
Figure 1. Schematic drawing representing the potential cell targets of PRP during skeletal muscle repair/regeneration, which may mediate its beneficial effects.
Figure 1. Schematic drawing representing the potential cell targets of PRP during skeletal muscle repair/regeneration, which may mediate its beneficial effects.
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