Skeletal muscle is one of the largest and most vital organs in the body, and a plastic organ maintained by multiple pathways regulating cell and protein turnover, and also a secretory organ secreting a variety of compounds including myokines, metabolites, microRNAs, and exosomes so on (1,2). There are a variety of pathophysiologic conditions, such as denervation, ageing, immobilization, sepsis and cancer, inducing skeletal muscle atrophy (3). Skeletal muscle atrophy occurs when the rates of protein degradation exceed the rates of protein synthesis, and involves in multiple distinct signaling pathways (4), and it is also associated with poor prognosis and the decline of life quality. However, the treatment of skeletal muscle atrophy remains an unresolved challenge to this day (5). Therefore, improving our understanding of the mechanisms responsible for skeletal muscle atrophy is very important to develop new clinical therapeutic strategies to prevent skeletal muscle atrophy.
Skeletal muscle atrophy characterized by loss of skeletal muscle mass, mitochondrial dysfunction, muscle fiber type transition, and a reduced physical functional capacity. Increasing evidence showed that inflammation was an important contributor and driver to skeletal muscle atrophy (6,7). Systemic inflammation could promote muscle atrophy via increased ubiquitin-proteasome system (UPS) and the autophagy-lysosome system (ALS) mediated protein degradation (8). Hindlimb immobilization model suggested the presence of an early inflammatory process, as evidenced by elevated IL-6, IL-1β and TNF-α, in skeletal muscles (9), and the proinflammatory cytokines (IL-6, IL-1β and TNF-α) could induce muscle proteolysis (10). IL-6 infusion resulted in muscle atrophy, and inhibition of IL-6 receptor reduced MuRF1 expression and ameliorated tail suspension-induced skeletal muscle atrophy (4,11-13). Inhibition of IL-1β could prevent inflammation-induced muscle atrophy in critically ill patients (14,15). TNF-α could induce C2C12 myotube atrophy through contributing to both hyperexpression of MAFbx and MuRF1 (16,17). Moreover, these inflammatory factors are involved in many signaling pathways, and these pathways collaborate together to trigger muscle atrophy.
Emerging evidence supports a key role for the Janus kinase/signal transduction and activator of transcription (Jak/STAT) signaling pathway in inflammation, and the Jak/STAT pathway is now recognized as a trigger of muscle protein loss (18,19). Silva et al. found that the activation of Jak/STAT3 pathway was significantly induced in muscle in mice with cachexia, and the inhibition of p-STAT3 in muscle decreased the expression of MAFbx and myostatin, alleviated muscle atrophy, and increased body weight and grip strength (19). Smith et al. demonstrated that controlled mechanical ventilation (CMV)-induced diaphragm atrophy accompanied with marked increases in STAT3 phosphorylation. Inhibition of Jak signaling could prevent the activation of STAT3 during CMV, reverse the pathologic alterations in the diaphragm, and maintain normal diaphragm contractility (20). Jak2/STAT3 pathway was also activated in muscles of immobilized intensive care unit (ICU) patients, and Jak2/STAT3 inhibitors could effectively alleviate muscle atrophy, and may provide an attractive pharmacological intervention strategy in immobilized ICU patients (21). However, the role of Jak2/STAT3 pathway in denervated muscle atrophy remains unclear.
Nuclear factor κB (NF-κB) is best known for its pro-inflammatory action, and emerging evidence suggests that the inflammation-sensitive NF-κB pathway may contribute to muscle atrophy (8,22). NF-κB activation (phospho-p65) is required for C2C12 myotube atrophy induced exogenous TNF-α, IL-1α or IL-1β, and inhibition of p65 could reverse C2C12 myotube atrophy due to TNF-α, IL-1α or IL-1β (23). NF-κB activation (phospho-p65) was markedly induced in muscle in mouse with food deprivation, and was critical for the subsequent muscle atrophy. While, NF-κB inhibition in muscle could prevent muscle atrophy induced by fasting (24). Cachectic factors TNF-α could activate NF-κB transcription pathway and p38 MAPK pathway, inducing skeletal muscle atrophy via upregulating MuRF1 (25,26), and p38 MAPK pathway is a trigger of the upregulation of MuRF1 (25). The activation of p38 MAPK and JNK were markedly induced by cigarette smoke extract in myotubes, and the p38 MAPK inhibitor significantly reduced USP-19 gene expression and alleviated muscle atrophy (27). Schellino et al. demonstrated that the activation of JNK in spinal muscular atrophy (SMA) mouse model was remarkably induced, and JNK-inhibitor could improve the trophism of SMA muscular fibers and motor performances (28). The inhibition of the JNK signaling pathway could also prevent lipopolysaccharide (LPS)-mediated Atrogin-1/MAFbx expression in C2C12 myotubes (29). Additionally, p38 MAPK and JNK were also activated, and augmented the expression of MAFbx in skeletal muscles suffered from limb immobilization and denervation (30,31). However, the exact molecular mechanisms still need to be further explored.
TGF-β1 is a multifunctional cytokine involving in the activation of satellite cells, the formation of connective tissue and the regulation of immune response intensity (32). In skeletal muscle, TGF-β1 could induce skeletal muscle atrophy, requiring Smad3 and JNK activation, and inhibition of Smad3 and JNK1/2 activation could reverse the atrophic effect of TGF-β (33,34). TGF-β1 and p-Smad3 showed significantly increase in murine model of amyotrophic lateral sclerosis, and targeting TGF-β signaling pathway might be a therapeutic approach to improve muscle function in several degenerative diseases (35). TGF-β1 was also induced in denervated muscle and promoted skeletal muscle atrophy (36). Smad3 was an important downstream signaling molecule for TGF-β, and Smad3 was significantly activated in hindlimb unloading (HU)-induced muscle atrophy (37). Other study showed that inhibition of Smad3 could suppress STAT3 signaling (38).
In summary, these signaling pathways, such as Jak2/STAT3, TGF-β1/Smad3, JNK/p38 MAPK and NF-κB signaling pathways, are all involved in muscle atrophy, and there are interactions or networks between these pathways to regulate muscle atrophy, which makes the process of muscle atrophy very complicated. Thus, if we only intervene in one pathway to protect muscle atrophy, we may not achieve the satisfactory effect.
Pyrroloquinoline quinone (PQQ), an aromatic tricyclic o-quinone, becomes an attractive bio-agent due to its multifunction, including anti-inflammatory property (39,40). Liu et al. found that PQQ could reduce the production of proinflammatory cytokines (TNF-α and IL-6), and inhibit the phosphorylation level of p65, p38 and JNK in IL-1β-stimulated human synovial sarcoma SW982 cells (40). PQQ could inhibit the activation of TGF-β1/Smad signaling in Bmi-1 knocked down human renal proximal tubular epithelial (HK2) cells (41). As reported above, PQQ can inhibit proinflammatory cytokines and their downstream signaling pathways. However, it is not clear whether PQQ could protect against denervated skeletal muscle atrophy through alleviating inflammation. Hence, the aim of current study is to investigate the influence of PQQ on skeletal muscle atrophy, and to explore the underlying molecular mechanism.
All animal procedures in this study were carried out in accordance with the recommendations of the Institutional Animal Care and Use Committee of Nantong University, and were approved by the Institutional Animal Care and Use Committee of Nantong University. Animals were subjected to unilateral sciatic nerve transection as described previously (42), followed by intraperitoneal injection of saline (100 µL) (Den group), or PQQ (5 mg/kg/day) in 100 µL saline (PQQ group) every day. The choice of PQQ concentration is based on our previous study (42). Intraperitoneal injection of 100 µL saline was administered to the animals received sham-operation (Ctrl group). After 14 days, mice were anesthetized and muscle tissues were obtained, weighed, and frozen in liquid nitrogen for further investigation.
Quantitative real-time polymerase chain reaction (qRT-PCR)
Total RNA was extracted from muscles with TRIZOL reagent (Invitrogen, Carlsbad, CA, USA), subsequently the cDNA was synthesized. The qRT-PCR was performed with the iTaq Fast SYBR Green Supermix (Bio-Rad, Hercules, CA, USA) under the manufacturer’s instructions. Quantitative data were acquired and analyzed under the Applied Biosystems 7500 RT-PCR system (Applied Biosystems, Foster City, CA, USA). The RT-PCR conditions were as follows: 42 °C for 20 min and then 40 cycles at 95 °C for 5 min, 94 °C for 20 s, and 72 °C for 42 s (4). The primers were as follows: mouse IL-6 F: CTGCAAGAGACTTCCATCCAG, R: AGTGGTATAGACAGGTCTGTTGG; mouse IL-1β F: GAAATGCCACCTTTTGACAGTG, R: TGGATGCTCTCATCAGGACAG; mouse TNF-α F: CAGGCGGTGCCTATGTCTC, R: CGATCACCCCGAAGTTCAGTAG; mouse β-actin F: GCTTCTAGGCGGACTGTTACT, R: GCCTTCACCGTTCCAGTTTTT. The relative mRNA expression was measured through the 2−ΔΔCt method (43).
Western blot analysis
Western blot analysis was used to determine the protein expression as described previously (4). The total proteins were extracted from muscles with RIPA Lysis Buffer, after centrifugation, soluble proteins were quantified with BCA Kit (Beyotime, Nantong, China) and separated on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). Subsequently, the proteins in the gel were transferred to polyvinylidene difluoride (PVDF) membranes (Millipore Corp, Billerica, MA, USA). The PVDF membranes were blocked with 5% nonfat dry milk in Tris-buffer saline, then the PVDF membranes were incubated with primary antibodies: mouse anti-MHC polyclonal antibody (R&D Systems, Minneapolis, MN, USA), rabbit anti-MuRF1, MAFbx (Fbx32), LC3B, PINK1, BNIP3, ATG7, TGF-β1, Smad3, p-Smad3, p65, p-p65, p38, p-p38, p-JNK, JNK, PGC-1α, Sirt1 and beta tubulin antibodies (Abcam, Cambridge, UK), Jak, STAT3 and p-STAT3 (CST, USA), Troponin I-FS and Troponin I-SS (Santa Cruz, USA), at 4 °C overnight. Then, the PVDF membranes were incubated with horseradish peroxidase (HRP) conjugated corresponding IgG antibodies for 2 h. The target proteins on the PVDF membranes were visualized by enhanced chemiluminescence (Thermo Scientific, Park Ellisville, MO, USA), and the intensity values were obtained for further normalization against loading control.
Muscle fiber cross-sectional area (CSA) size
The fiber CSA of skeletal muscles was determined by laminin staining. Briefly, muscles were fixed, flash-frozen, and sectioned on cryostat with 10 µm thickness. The sections were incubated with anti-laminin antibody (1:200) (Abcam, Cambridge, UK) at 4 °C overnight. The following day, sections were washed and incubated with the Alexa Fluor secondary antibody (1:400) (Invitrogen Antibodies, Waltham, MA, USA) for 30 min at room temperature. Then, the sections were imaged by fluorescence microscopy (Zeiss, Germany) and the CSA of myofibers was obtained with ImageJ software (NIH, Bethesda, MD, USA).
Muscle fiber type transition analysis
The fiber type transition is observed in the soleus muscles due to that the change of soleus muscle fiber type was significant after denervation (44). The sample processing method was similar to the CSA analysis. Tissue sections were incubated with Anti-Fast Myosin Skeletal Heavy chain antibody (1:200) (Abcam, Cambridge, UK) for 12 hours at 4 °C. Subsequently, the sections washed and incubated with the Alexa Fluor secondary antibody (1:400) (Invitrogen Antibodies, Waltham, MA, USA) for 30 min at room temperature. Then, the sections were imaged by fluorescence microscopy (Zeiss, Germany).
Enzyme-linked immunosorbent assay (ELISA)
The content of IL-1β, IL-6 and TNF-α was measured using appropriate commercial ELISA analysis (4). Briefly, ELISA plates (Beyotime, Haimen, China) were washed and incubated with 100 µL muscle lysates from the TA muscles at 37 °C for 2 h. Following, ELISA plates were washed and incubated with biotinylated polyclonal anti-IL-1β, anti-IL-6 or anti-TNF-α antibodies at 37 °C for 1 h. Subsequently, ELISA plates were washed and incubated with HRP-Streptavidin at 37 °C for 20 min in dark. Finally, enzyme activity was read at 450 nm in a microplate spectrophotometer.
Transmission electron microscopy (TEM) analysis
TEM analysis was used to observe the changes of mitochondria in skeletal muscles. The detailed procedures of TEM for muscle were previously reported (45). Briefly, muscles were first fixed in 2.5% glutaraldehyde, subsequently were fixed in 1% osmium tetroxide. Finally, muscle sections were analyzed by TEM (HT7700, Hitachi, Tokyo, Japan).
All data in this study were analyzed with one-way analysis of variance (ANOVA), followed by the Tukey’s multiple comparisons test to detect differences between groups. All statistical analyses in current study were performed with GraphPad Prism software (version 7.0) (San Diego, CA, USA). P<0.05 was regarded as statistically significant.
PQQ inhibited proinflammatory cytokines in denervated skeletal muscles
After denervation, mice were injected intraperitoneally with saline vehicle plus PQQ (5 mg/kg/d) or saline vehicle only (Den) for 14 days. After sham operation, mice were injected intraperitoneally with saline vehicle (Ctrl) for 14 days. Then, the inflammatory cytokines were determined through ELISA and qRT-PCR analyses. The present study showed that inflammation was significantly induced in the tibialis anterior (TA) muscles after sciatic nerve transection, as evidenced by the increased proinflammatory cytokines, such as IL-6, IL-1β and TNF-α in the denervated TA muscles. Interestingly, PQQ could decrease the production of IL-6, IL-1β and TNF-α in the denervated TA muscles, as shown in Figure 1. These findings suggested that PQQ could inhibit inflammation through decreasing the proinflammatory cytokines in denervated skeletal muscles.
PQQ blocked the activation of inflammatory signaling pathway in denervated skeletal muscles
The signaling pathways involved in inflammatory factors IL-6, IL-1β and TNF-α mainly include Jak2/ STAT3, TGF-β1/Smad3, JNK/p38 MAPK and NF-κB signaling pathways. In this study, we found that Jak2/STAT3, TGF-β1/Smad3, JNK/p38 MAPK and NF-κB signaling pathways were significantly activated in denervated TA muscles, as evidenced by the increased levels of Jak2, TGF-β1, and phosphorylated Stat3, Smad3, JNK, p38 and p65 in the TA muscles after sciatic nerve transection, as illustrated in Figure 2, which indicated that these signaling pathways involved in inflammation were remarkably induced after denervation in skeletal muscles. Intriguingly, PQQ could decrease the levels of Jak2, TGF-β1, and phosphorylated STAT3, Smad3, JNK, p38, p65 in the TA muscles after sciatic nerve transection, as illustrated in Figure 2, which suggested that PQQ could block the activation of inflammatory signaling pathway in denervated skeletal muscles.
PQQ attenuated denervation-induced skeletal muscle atrophy
In order to explore the effects of PQQ on muscle atrophy induced by denervation, the muscle mass and myofiber CSA of TA muscles were determined. In current study, we found that denervation-induced the loss of muscle mass and the decreased average myofiber CSA of TA muscles was significantly reversed by PQQ administration, which was illustrated by increased myofiber CSA and TA muscle mass, as shown in Figure 3. Additionally, we found that PQQ could inhibit denervation-induced decrease of MHC in TA muscles through suppressing the ubiquitin proteasome system proteolytic activity, as evidenced by the decreased levels of muscle-specific E3 ubiquitin ligases Atrogin-1/MAFbx and MuRF1 in denervated skeletal muscles, as illustrated in Figure 3. These observations suggested that PQQ attenuated denervation-induced skeletal muscle atrophy through, at least in part, inhibiting ubiquitin proteasome proteolytic pathway.
PQQ suppressed denervation-induced mitophagy in skeletal muscles
Mitochondrial dysfunction is associated with skeletal muscle atrophy (46). Therefore, we investigated the effects of PQQ on autophagosomes or autophagic vacuoles and autophagy-related molecules (ATG7, PINK1, BNIP3 and LC3B) in skeletal muscle upon denervation. Our data showed that mitophagy was considerably activated in the TA muscles after sciatic nerve transection, as illustrated by the increased autophagosomes or autophagic vacuoles and the elevated autophagy-related molecules, including ATG7, PINK1, BNIP3 and LC3B (Figure 4). Interestingly, the autophagosomes or autophagic vacuoles were significantly reduced, accompanied by the decreased autophagy-related molecules, in the muscles of denervated mice treated with PQQ compared with that treated with saline (Figure 4). Collectively, these findings indicated that PQQ suppressed denervation-induced mitophagy and the expression of autophagy-related molecules in denervated skeletal muscles.
PQQ suppressed slow-to-fast myofiber-type transition in skeletal muscles after denervation
The myofiber-type transition is one of important features of skeletal muscle atrophy (47). The soleus muscles were chosen to further study in this observation due to that the change of soleus muscle fiber type was significant after denervation (44). To investigate the effects of PQQ on myofiber-type transition induced by denervation, the soleus muscles were harvested to undergo MyHC II staining analysis after sciatic nerve transection. MyHC II staining analysis showed that the percentage of MyHC II positive fibers was markedly increased in the soleus muscles of mice after denervation compared with sham-operated mice (Figure 5A,B), which suggested that denervation induced the slow-to-fast myofiber-type transition in soleus muscles. Interestingly, the denervation-induced slow-to-fast myofiber-type transition was rescued in soleus muscles of the mice treated with PQQ, as evidenced by the decreased percentage of MyHC II positive fibers (Figure 5A,B). Additionally, Troponin I-SS and Troponin I-FS antibodies were used to distinguish the slow and fast skeletal muscle fibers with western blotting, respectively. Our results showed that the expression of Troponin I-SS was decreased and the expression of Troponin I-FS was increased in denervated soleus muscles and PQQ could reverse the phenotype induced by denervation, which was consistent with the results from MyHC II staining analysis (Figure 5C,D,E). PGC-1α and Sirt1 are all involved in maintaining the oxidative fiber type. In current study, PGC-1α and Sirt1 all displayed a significant decrease in denervated soleus muscles and this decrease of PGC-1α and Sirt1 was rescued by PQQ administration (Figure 5F,G). These findings demonstrated that PQQ reversed slow-to-fast myofiber-type transition through, at least in part, enhancing the expression of PGC-1α and Sirt1 in skeletal muscles after denervation.
Skeletal muscle atrophy, characterized by loss of muscle mass, shrinkage of muscle fibers, mitochondrial dysfunction, alteration in fiber-type composition, and reduced physical function, occurs when protein degradation exceeds protein synthesis under the conditions of denervation, cancer cachexia and aging (48). More recently, understanding about the molecular mechanisms regulating skeletal muscle atrophy has taken a quantum leap, though most of the current researches were focused on the role of various effector kinases and some downstream transcription factors (49,50). The initial events that trigger the activation of atrophying signaling pathways remain poorly defined. Our study demonstrated that inflammation might be a major upstream regulator of skeletal muscle atrophy in response to denervation, as evidenced by the increased proinflammatory cytokines and the activated inflammatory signaling pathways in denervated skeletal muscles, which was in agreement with recent reports that inflammation was an important contributor and driver to skeletal muscle atrophy in other conditions, such as diabetes and cancer (6-8,51). Although it remains unknown what elicits the inflammatory response in atrophying muscle in response to denervation, it was of interest to note that inflammation might be an important mediator and driver to skeletal muscle atrophy, and blocking inflammation in skeletal muscle might be a novel and exciting therapeutic approach for muscle wasting in condition of denervation.
PQQ is a natural antioxidant found in soil (39). Recent studies have found that PQQ also has anti-inflammatory function. Liu et al. found that PQQ could reduce the production of TNF-α and IL-6, and inhibit the signal activation of p65, p38 and JNK in IL-1β-stimulated human synovial sarcoma SW982 cells (40). PQQ also could inhibit the activation of TGF-β1/Smad signaling (41). Although it remains unknown how PQQ suppresses inflammation, it was of interest to note that PQQ might provide a potent therapeutic benefit for the treatment of muscle atrophy.
Intriguingly, the results of the present study suggested that PQQ could alleviate skeletal muscle atrophy, mitigate mitophagy and inhibit slow-to-fast muscle fiber type transition in denervated skeletal muscles. The anti-muscular atrophy function of PQQ has been confirmed by Kuo’s research that demonstrated PQQ inhibiting skeletal muscle atrophy in response to denervation by activating PGC-1α (52). Our previous studies have also confirmed that PQQ can alleviate muscle atrophy by inhibiting reactive oxygen species (ROS) (42,53). The inflammatory cytokine could decrease PGC-1α expression, and promotes ROS generation (54,55). Whether PQQ improves skeletal muscle after denervation through suppressing inflammation is an area of research for future investigation.
It is not clear whether PQQ could protect against denervated skeletal muscle atrophy through alleviating inflammation. We further investigated the effects of PQQ on inflammation in skeletal muscles upon denervation. Previous studies have emphasized that inflammatory cytokines were important triggers and mediators of muscle atrophy in both animal models and patients with advanced cancer (56-59). Interestingly, our findings demonstrated that inflammation was significantly induced, and PQQ could not only reduce the production of proinflammatory cytokines IL-6, IL-1β and TNF-α, but also block the signal activation of Jak2/STAT3, TGF-β1/Smad3, JNK/p38 MAPK, and NF-κB in skeletal muscles after denervation. These findings revealed that PQQ could inhibit proinflammatory cytokines and their downstream signaling pathways during denervated skeletal muscles atrophy.
Jak/STAT3 pathway is now recognized as a trigger of muscle protein loss, and the inhibition of p-STAT3 in muscle decreased the expression of MAFbx and myostatin, alleviated muscle atrophy, and increased body weight and grip strength (18-21). STAT3/NF-κB inhibition increased protein synthesis in skeletal muscles in cachectic mice, alleviated muscle atrophy (60). The increased activation of STAT3 could upregulate the markers of the mitophagy/autophagy (LC3B, BNIP3) in left ventricular tissue of metastatic melanoma mice (61), which indicated that inactivation of STAT3 might inhibit mitophagy. Sirt1 suppresses the activation of STAT3, and improves the activation of PGC-1α (62,63). Moreover, sirt1 overexpression could trigger the transition of myofiber fast-to-slow, increase the expression of PGC-1α, and decrease the expression of atrophic genes (64). In this study, PQQ suppressed the activation of Jak/STAT3, and improved the expression of Sirt1 and PGC-1α, accompanied with decreased the expression of UPS and ALS related proteins, reduced Troponin I-FS (fast twitch protein), and increased Troponin I-SS (slow twitch protein), which was likely responsible for the mechanism by which PQQ alleviated skeletal muscle atrophy, mitigated mitophagy and inhibited slow-to-fast muscle fiber type transition in denervated TA muscles.
NF-κB activation in skeletal muscle is important for atrophy upon denervation or cancer (24). Inflammatory signaling pathways, especially NF-κB, potentially contribute to diabetes-mediated muscle atrophy (8). Overexpression of PGC-1α could increase mitochondrial content (regulating myofiber-type transition), suppress the activation of NF-κB, and inhibit the expression of atrophic genes such as MAFbx and MuRF1, alleviating muscle atrophy (65). In current study, PQQ could improve the expression of PGC-1α, and suppress the activation of NF-κB, which might be responsible for PQQ attenuating muscle atrophy and the transition of slow-to-fast in condition of denervation.
The activation of p38 MAPK and JNK were markedly induced by cigarette smoke extract in myotubes, and the p38 MAPK inhibitor significantly alleviated muscle atrophy (27). JNK and p38 MAPK signaling pathways were also activated, and augmented the expression of MAFbx in skeletal muscles suffered from limb immobilization and denervation (30,31). JNK/c-Jun pathway could inhibit the expression of PGC-1α in cerebellar granule neurons (66). The inactivation of JNK and p38 MAPK resulted in a reduction of palmitate-induced autophagy (67). These reports revealed that inactivation of p38 MAPK and JNK might be beneficial for the treatment of muscle atrophy. Intriguingly, our findings suggested that PQQ could inhibit the activation of p38 MAPK and JNK, as evidenced by decreased levels of phosphorylated p38 MAPK and JNK in denervated TA muscles. It was likely responsible, at least part, for the mechanism by which PQQ alleviated skeletal muscle atrophy, mitigated mitophagy and inhibited slow-to-fast muscle fiber type transition induced by denervation.
TGF-β1 could induce skeletal muscle atrophy through activating Smad3, and inactivation of Smad3 could reverse the atrophic effect induced by TGF-β (33,34). The activation of TGF-β signaling pathway is also crucial for autophagy, and it could activate autophagy of human hepatic carcinoma HepG2 cells (68). TGF-β/Smad3 signaling pathway could suppress the expression of PGC-1α (69), and the activation of TGF-β/Smad3 pathway could be inhibited by sirt1 (70). These data indicated that regulating the activity of TGF-β/Smad3 pathway might be associated the skeletal muscle atrophy, autophagy and the expression of PGC-1α and sirt1, which were responsible for oxidative metabolism in muscles. Thus, it can be seen that inhibition of TGF-β/Smad3 pathway might be a novel therapeutic intervention for prevention of skeletal muscle atrophy. Interestingly, PQQ could reduce the activity of TGF-β/Smad3 pathway, as illustrated by decreased TGF-β1 and phosphorylated smad3 in denervated muscles, which revealed that PQQ could alleviate skeletal muscle atrophy, mitigate mitophagy and inhibit slow-to-fast muscle fiber type transition through, at least part, inhibiting the activation of TGF-β/Smad3 pathway.
Collectively, these findings highlighted the importance of the inflammatory signaling mechanisms in denervation-induced muscle atrophy and suggested that inflammation might be an important therapeutic target to prevent skeletal muscle wasting induced by denervation. Interestingly, PQQ could attenuate skeletal muscle atrophy, mitophagy and fiber type transition due to denervation through suppressing the Jak2/STAT3, TGF-β1/Smad3, JNK/p38 MAPK, and NF-κB signaling pathways (Figure 6). This study not only enriches the molecular regulatory mechanism of denervated muscle atrophy, but also provides a novel and exciting therapeutic approach for muscle wasting in condition of denervation.
Funding: This work was supported by National Natural Science Foundation of China (Grant No. 31730031, 81871554 and 81671230), the 973 Program (Grant No. 2014CB542202 and 2014CB542203), a project funded by Jiangsu Provincial Key Medical Center and the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD), and 2019 Jiangsu College Students Innovation and Entrepreneurship Training Program.
Conflicts of Interest: The authors have no conflict of interests to declare.
Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All animal procedures in this study were carried out in accordance with the recommendations of the Institutional Animal Care and Use Committee of Nantong University, and were approved by the Institutional Animal Care and Use Committee of Nantong University (No. 20170305-003).
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