Porous Se@SiO₂ nanoparticles improve oxidative injury to promote muscle regeneration via modulating mitochondria

Acute skeletal muscle injuries are common among physical or sports traumas and are one of the most significant causes of disability [1]. Besides being prone to recurrence, this type of injury can also include pain, swelling and possibly osteofascial compartment syndrome [2]. Apart from mechanical damage to muscle cells, skeletal muscle injury also involves capillary rupture, hemorrhage, muscle tissue edema and inflammation, resulting in cytokines and inflammatory cells allowing direct access at the site of injury, induced mitochondrial dysfunction manifested as reactive oxygen species (ROS) overload, inhibition of mitochondrial respiration and disturbances in calcium homeostasis [3].

Despite advances in sports medicine, effective treatments for skeletal muscle injury are still lacking. Surgical intervention, ice and rehabilitation exercises are the most common treatments for skeletal muscle injury, but the outcomes are unsatisfactory, and they prolong the functional recovery of the muscle, sometimes even predisposing it to repeated injury [4]. Though the main aim of the pharmacological intervention is to suppress the activation of inflammatory pathways, related reports have shown that excessive inhibition of inflammatory signaling pathways would affect the immune system and ultimately impair muscle regeneration [5]. Inefficient muscle regeneration often leads to poor contractility and progressive loss of muscle strength, resulting in loss of function [6]. Therefore, there is a great need to develop a new therapeutic strategy for musculoskeletal injuries.

Satellite cells (SCs) are involved in repairing skeletal muscles after an injury [7]. They normally remain in a quiescent state and are characterized by the expression of PAX7 [8,9]. SCs initiate proliferation in response to specific stimuli such as direct trauma or extensive physical activity, and then they differentiate into new myofibers under the influence of specific factors, including MYOD and myogenin. They regulate the transcription of essential muscle-specific proteins such as MYHC [10]. However, in some cases, the injury is more severe than the compensatory ability of the skeletal muscle, and the number or function of SCs is restricted as a result of oxidative damage, which can lead to incomplete muscle regeneration and excessive fibrosis. Therefore, modulation of SCs at the site of injury has considerable therapeutic potential for promoting skeletal muscle regeneration.

Mitochondria are essential organelles found in the cytoplasm of almost all eukaryotic cells. They are double-membrane-structure organelles with independent, self-replicating genomes and regulated cell metabolism. The oxidation/reduction O₂-consuming metabolic pathways in mitochondria ensure survival at the cellular and organismal levels. In addition to their primary function, mitochondria also perform other essential functions, including biomolecule synthesis, ROS production, regulation of apoptosis and calcium homeostasis [11,12]. Mitochondrial dysfunction and oxidative stress play an important role in various metabolic disorders, including diabetes, obesity and hypertension [13]. As the largest metabolically active and most highly structured organelle, mitochondrial functions play vital roles in skeletal muscles. In addition to regulating the energy requirements of the cell, mitochondria are involved in many pathological processes [14].

Overproduction of ROS is known to be caused in part by mitochondrial dysfunction. Although mitochondria are the major site of ROS production, the antioxidant enzyme system in mitochondria can scavenge excessive ROS [15]. When mitochondrial function is impaired, severe oxidative stress conditions limit the proliferation and differentiation of SCs, and the process of muscle regeneration is hindered. Because the application of antioxidants protects cells from death, it is evident that excessive oxidative stress results in cell death. The levels of antioxidants in SCs are a major factor in the outcome of cell-based therapies [16]. In addition, SCs regulate the regeneration of skeletal muscles by regulating the structure, metabolism and activity of mitochondria [12,17,18]. RNA sequencing of SCs activated after injury has revealed significant upregulation of mitochondrial genes, suggesting active mitochondrial metabolism [19]. However, few studies have examined the effects of antioxidant treatment on the mitochondria of SCs that are under oxidative stress, or even on the mechanisms of ROS in SCs [20]. Therefore, reducing oxidative damage to SCs by modulating mitochondrial activity may be a promising strategy for further promoting muscle regeneration after injury.

Selenium (Se) is an essential micronutrient with a wide range of biological functions that can inhibit oxidative stress and scavenge excessive ROS [21]. However, the application of Se as a complementary therapy is narrow, and toxic effects due to excessive aggregation during Se administration in conventional or bulk form limit its application [22,23]. The development of nanotechnology appears to be a feasible solution. The authors designed porous Se@SiO₂ nanoparticles (NPs), since they would slowly release Se from porous Se@SiO₂ NPs and act as an ideal delivery system [24,25]. Porous Se@SiO₂ NPs also have been shown to alleviate many diseases, including improving mitochondrial dysfunction in acute lung injury and alleviating oxidative damage. They are also effective in treating various diseases associated with oxidative damage or mitochondrial dysfunction [26].

The authors of this study hypothesized that porous Se@SiO₂ NPs could promote muscle regeneration and protect the myogenic differentiation of SCs. They created an oxidative stress environment using H₂O₂ in an in vitro cell culture. To induce myofiber necrosis and regeneration, they injected cardiotoxin (CTX) into the tibialis anterior (TA) muscle of rats to construct a model of acute injury. The potential mechanisms underlying the effect of porous Se@SiO₂ NPs were investigated. This study provides preliminary evidence that porous Se@SiO₂ NPs promote the proliferation and myogenic differentiation of SCs by scavenging mitochondrial ROS and improving mitochondrial activity under oxidative stress, ultimately promoting skeletal muscle regeneration in rats.

Material & methods

Preparation & characterization of porous Se@SiO₂ NPs

The porous Se@SiO₂ NPs were synthesized following the procedure used in the authors' previous research [27]. A mixture of 2 ml of Cu_2–x Se nanocrystals synthesized in nitrogen was mixed thoroughly with 20 ml of n-hexane, 20 ml of n-hexanol, 2 ml of Triton X-100, 0.6 ml of deionized water and 0.08 ml of ethyl orthosilicate. Subsequently, 0.1 ml of ammonia was added and orthosilicate was hydrolyzed to encapsulate the silica in the Se quantum dots prepared by oxidation to form solid Se@SiO₂ nanospheres. Finally, they were dispersed in 10 mg/ml of polyvinylpyrrolidone solution and etched in hot water to form porous structures. The porous Se@SiO₂ NPs were characterized by transmission electron microscopy (HT7700, HITACHI, Tokyo, Japan) and D/max2550 x-ray diffractometer (Cu K-α radiation; Rigaku, Tokyo, Japan).

Isolation & culture of SCs

All experiments were approved by the Animal Ethics Committee of Yangzhou University and conducted in accordance with the regulation and guidance. SCs were prepared as described previously [28]. Newborn male Sprague-Dawley suckling rats were euthanized, and the TA and gastrocnemius muscles were dissected. The muscle tissue was homogenized and enzymatically digested by adding type II collagenase. The enzymatically cleaved tissue homogenate was then centrifuged at 400–500 r.p.m. for 3 min. The supernatant was removed, and the pellet was resuspended in Collagenase/Dispase (Roche, Basel, Switzerland) and incubated in a water bath on a rocker. The homogenate was passed through a series of mesh cell strainers. The filtrate was centrifuged in a tabletop centrifuge at 1200 r.p.m. for 10 min. The pellet was resuspended gently in a growth medium containing DMEM (HyClone, UT, USA) supplemented with 20% fetal bovine serum (HyClone), 3% chicken embryo extract, 4 mM L-glutamine, 10 mM sodium pyruvate and 1% penicillin–streptomycin (Sangon Biotech, Shanghai, China). The solution was poured into Petri dishes and preplated in an incubator (5% CO₂; 37°C) to remove fibroblasts. At about 3–5 days, the SC culture was shifted from growth medium to differentiation medium (DMEM supplemented with 5% horse serum, 1% penicillin/streptomycin, 4 mM L-glutamine and 10 mM sodium pyruvate). SCs from passages 3–4 were used for later experiments. The cells proliferated faster in about the first 3 days, then gradually started to differentiate, and differentiation was more obvious around day 7. Subsequent assays related to cell proliferation, death and senescence were selected on day 3, and assays related to myogenic differentiation and mitochondria were performed on day 7.

Establishment of H₂O₂-induced damage models of SCs

Based on the presence or absence of 20 μg/ml porous Se@SiO₂ NPs and 100 μM H₂O₂, cells were divided into four groups: control group, in which cells were cultured in differentiation medium only; Se@SiO₂ NPs group, in which cells were treated with differentiation medium and porous Se@SiO₂ NPs; H₂O₂ group, in which cells were treated with differentiation medium and then with 100 μM H₂O₂ for 24 h; Se@SiO₂ + H₂O₂ group, in which cells were treated with differentiation medium and porous Se@SiO₂ NPs and then with 100 μM H₂O₂ for 24 h. To reduce the impact of H₂O₂ evaporation on the experiment, the authors increased the number of liquid changes in the medium within 24 h of adding H₂O₂. A day later, H₂O₂ solution was removed, and the cells were washed with phosphate-buffered saline (PBS) solution. All groups were cultured for 7 days.

In vitro safety of porous Se@SiO₂ NPs & cell proliferation assay

First, the SCs were cultured with different concentration gradients of porous Se@SiO₂ NPs (0–160 μg/ml), and cell samples were harvested after 1 day. In addition, the four groups of cell samples were harvested on days 1, 2 and 3. The effects of porous Se@SiO₂ NPs and H₂O₂ on cell proliferation were assessed by MTT assay. Briefly, culture medium containing dimethyl sulfoxide (Invitrogen, MA, USA) served as control, and MTT solution (Sigma-Aldrich, MO, USA) was added to each well. Then the mixed liquid was sucked out, 150 μl/well of dimethyl sulfoxide was added and it was shaken for 10 min. The absorbance was measured by a microplate reader (MK3 type, Multiskan GO; Thermo Fisher Scientific, MA, USA) at 492 nm.

Staining of live–dead cells & senescence-associated β-galactosidase (SA-β-gal) staining

Cell samples were obtained after 3 days of induction. The diluted calcein AM/PI kit (Biovision, CA, USA) was added according to the instructions and incubated for 30 min, followed by washing. It was observed using a fluorescence microscope (Olympus IX71, Olympus, Tokyo, Japan). Red and green denoted dead cells and live cells. The ratio of living cells to dead cells was measured by ImageJ software, version 1.39.

The presence of senescent cells was also evaluated through the activity of senescence-associated β-galactosidase (SA-β-gal). Cells collected at the same time were washed with PBS. SA-β-gal staining fixation solution (Beyotime, Shanghai, China) was added, followed by fixing at room temperature. The solution was then removed and the cells were washed with PBS. PBS was removed and dye solution was added per well according to the instructions, followed by overnight incubation at 37°C. The observations were recorded using an optical microscope.

Detection of mitochondrial ROS

On days 3 and 7, cell samples were taken. Dimethyl sulfoxide was added to MitoSoX Red Mitochondrial Superoxide Indicator (Yeasen, Shanghai, China) and mixed well to prepare storage solution and was diluted with Hanks' balanced salt solution; 1–2 ml probe working solution was added and the cells were fully covered, followed by incubation at 37°C in the dark for 10 min. Cells were then counterstained with Hoechst (Beyotime) for 5 min, and after sealing it was observed by confocal laser scanning microscopy (TCS SP5, Leica, Heidelberg, Germany).

Measurement of mitochondrial morphology

The mitochondrial morphology and actin filaments of SCs were observed under a confocal microscope on day 7. First, SCs in each group, as mentioned earlier, were plated on confocal Petri dishes. Then they were incubated with the MitoTracker Red CMXRos (Beyotime) in the ratio of 1:1000–1:10,000 and F-actin dye solution (Abcam, MA, USA) at 37°C for 30 min. The intracellular fluorescence was observed under a confocal fluorescence microscope (Leica). MitoTracker Red fluorescence staining of mitochondria in live cells was determined by ImageJ software.

Evaluation of mitochondrial respiratory chain & catalase activities

On day 7, the activities of the mitochondrial respiratory chain (MRC) were measured by the MRC Complex Activity Assay Kit (Solarbio, Peking, China) and the ATP Synthase Activity Assay Kit (YBio; Shanghai, China) according to the instructions. MRC activities were determined using a colorimetric assay at 340 nm (MRC complex I), 550 nm (MRC complex III) and 450 nm (ATP synthase). The unit enzyme activity was exhibited as nmol/min/mg protein. At the same time, the CAT Activity Detection Kit (Solarbio) was used to measure the activity of catalase (CAT). According to the instructions, the enzyme labeling instrument (ThermoMK3; Thermo Fisher Scientific) section wavelength was set to 405 nm for reading and calculating the absorbance value. The unit enzyme activity was exhibited as μmol/min/mg protein.

Rat model of skeletal muscle injury

All experimental procedures with rats were approved by Animal Ethics Committee of Yangzhou University. Common methods were adopted, for which 24 male, 8-week-old rats were placed in a supine position; the left limb was rinsed with 95% ethanol. Approximately 30 μl of saline solution dissolved with CTX (20 μM; WeiGa, Guangzhou, China) was injected into the middle belly of the left TA muscle. After the injection, analgesia and intramuscular injection of penicillin were given to prevent infection. Then the rats were randomly divided into four groups. In the CTX + Se@SiO₂ group, porous Se@SiO₂ NPs (1 mg/kg) were injected intramuscularly in the same place 2 h after CTX injection and porous Se@SiO₂ NPs (1 mg/kg) were injected intraperitoneally per day; meanwhile, the right leg of the CTX + Se@SiO₂ group of rats was labeled as the Se@SiO₂ group. In the CTX group, normal saline was used in intramuscular injections 2 h after CTX injection and daily intraperitoneal injection of normal saline; at the same time, the right leg of the CTX group of rats was labeled as the control group. The concentration of porous Se@SiO₂ NPs was chosen to be 1 mg/kg, according to the authors' previous studies; there was no significant toxic effect on the vital organs of the animals, and no significant aggregation was found in vivo [29]. According to previous studies, morphological changes occurred immediately after CTX injection, and over time the localization went from an inflammatory response to muscle regeneration, with maturation of regenerated muscle fibers in about 1 month [30,31]. The animals were sacrificed on days 1, 7, 14 and 28 after CTX injection. After euthanasia, TA muscles were dissected and snap-frozen in liquid nitrogen or preserved in formalin. Follow-up experiments were conducted by three different experimenters blinded to the study.

Histological evaluation of muscle regeneration & fibrosis development

The TA muscle samples preserved in formalin were then dehydrated, embedded in paraffin and cut into 4 μm sections. Subsequently, the sections were deparaffinized, rehydrated and subjected to histological staining. Hematoxylin and eosin staining was performed to evaluate injury and regeneration of myofibers; the Masson Modified Trichrome Staining Kit (Solarbio) was used to evaluate the deposition of collagen according to standard protocols.

Western blot analysis

For cell samples, whole-cell extracts in all four groups were prepared on day 7; for animal samples, rat TA muscle was collected on days 14 and 28. The samples were washed with PBS, followed by lysis in precooled RIPA lysate (P0013B, Beyotime), and then the protein was quantified by a BCA protein assay kit (KeyGEN, Shanghai, China). The protein samples were separated on a sodium dodecyl sulfate-polyacrylamide gel and then transferred to a polyvinylidene difluoride membrane. After blocking, the membrane was incubated with primary antibodies: SIRT1 (Proteintech, Wuhan, China), SOD1 (Proteintech), MYHC (Thermo Fisher Scientific), MYOD (Proteintech) and GAPDH (Proteintech). The polyvinylidene difluoride membrane was then treated with rabbit secondary antibody or rat secondary antibody (ZSGB-BIO, Peking, China). By employing enhanced chemiluminescence solution (Dingguo, Peking, China), the proteins were visualized, and the immunoreaction bands were quantified by ImageJ software. Protein expression was represented as the densitometric ratio of protein normalized to GAPDH.

Immunofluorescence staining

With immunofluorescence staining, the expression of markers was detected using antibodies against desmin (Proteintech), PAX7 (Proteintech), MYHC (Thermo Fisher Scientific), MYOD (Invitrogen), F-actin (Abcam) and dystrophin (Santa Cruz Biotechnology, TX, USA). Briefly, cell samples were washed thrice with PBS and then fixed with 4% paraformaldehyde for 15 min. After washing with PBS, the cells were permeabilized with 0.25% Triton X-100 (Rich Joint, Shanghai, China) for 10 min and incubated in 5% fetal bovine serum at 37°C for 1 h. Afterward, cells were incubated with primary antibodies overnight. After washing, secondary antibody was used to incubate the cells for 1 h at room temperature. The cells were then washed and treated with Hoechst (Beyotime) to stain the cell nuclei. In the end, samples were imaged under a fluorescence microscope (Olympus). Using ImageJ software according to the method described by Menconi et al. [32,33], quantitative image analysis was performed by analyzing five randomly selected injury fields per sample; fusion index was calculated as the number of nuclei in myotubes divided by the total number of nuclei counted. The average number of nuclei per myotube was determined by dividing the number of nuclei in myotubes by the total number of myotubes.

Statistical analysis

Qualitative data were representative of at least three independent experiments. Quantitative or semiquantitative data are expressed as the mean ± standard deviation. The data were analyzed using GraphPad Prism 6. Data were managed through one-way analysis of variance and Student's unpaired t-test. A difference was defined as statistically significant only when p < 0.05.

Results

Characterization of porous Se@SiO₂ NPs & identification of SCs

Figure 1A illustrates the fabrication process of porous Se@SiO₂ NPs. Figure 1B shows the phase structure of Se@SiO₂ NPs analyzed by x-ray diffractometer, and the characteristic peaks of Se@SiO2 NPs show the standard Se hexagonal phase except for the amorphous silica leading to an increase in the low-angle region.

Transmission electron microscopy was used to measure the size and morphology of the porous Se@SiO₂ NPs. Porous Se@SiO₂ NPs had an average diameter of 55 nm; from the center to the surface, there were numerous irregular quantum dots (<5 nm; Figure 1C & D). The porous Se@SiO₂ NPs exhibited a porous structure being treated with hot water (Figure 1E). The SCs were identified by immunofluorescence staining for PAX7 and desmin, which are specific markers, and cytoskeletal alterations were detected by staining for F-actin (Figure 1F). This result confirmed that the isolated cells were SCs because PAX7 was distributed in the nucleus and desmin was present in the cytoplasm. The safety of porous Se@SiO₂ NPs was ascertained by determining the proliferation of cells in the presence of porous Se@SiO₂ NPs. Porous Se@SiO₂ NPs were added to the cells at concentrations ranging from 0 to 160 μg/ml. Compared with the control group, SCs showed different cell proliferation patterns, wherein the concentrations of 10 and 20 μg/ml significantly promoted cell proliferation, and concentrations above 40 μg/ml inhibited cell proliferation (Figure 1G). Therefore, for the porous Se@SiO₂ NPs used in the subsequent experiments, the authors chose to conduct them at a concentration of 20 μg/ml, which has a positive effect on cell proliferation.

Porous Se@SiO₂ NPs protect SCs against H₂O₂-induced injury

The MTT assay showed that porous Se@SiO₂ NPs promoted cell proliferation (Figure 2A). Compared with the control group, cell proliferation was significantly enhanced in the Se@SiO₂ group on days 1, 2 and 3. The cell proliferation was also higher in the Se@SiO₂ + H₂O₂ group than in the H₂O₂ group. These results demonstrated that porous Se@SiO₂ NPs remarkably enhanced the proliferation of SCs, and the inhibitory effect of H₂O₂ on cell growth was reduced.

In the staining of live–dead cells (Figure 2C & D), the green dye can permeate the cell membrane of live eukaryotic cells and stain the cell green, whereas the red dye stain can only label dead cells, since it is membrane impermeable. On day 3 of cell culture, H₂O₂ significantly increased cell death compared with the control group; however, the addition of porous Se@SiO2 NPs reduced cell death compared with that in the H₂O₂ group.

On day 3, senescent cells were detected based on the expression of SA-β-gal, a prominent biomarker of aging. The addition of porous Se@SiO₂ NPs did not affect cellular senescence (Figure 2B). Blue-stained SA-β-gal positivity was significantly elevated in the SCs after treatment with H₂O₂ compared with the control. Blue-stained SA-β-gal positivity was reduced significantly in the SCs treated with porous Se@SiO₂ NPs. These results show that porous Se@SiO₂ NPs significantly reduce the increase in cell senescence caused by oxidative stress.

Porous Se@SiO₂ NPs promote mitochondrial function under oxidative stress

The authors examined the ability of porous Se@SiO₂ NPs to promote the clearance of ROS in mitochondria on days 3 and 7. Porous Se@SiO₂ NPs did not significantly affect the control group but elevated mitochondrial ROS levels after exposure to H₂O₂ (Figure 3A & B).

To illustrate the underlying antioxidant mechanisms of porous Se@SiO₂ NPs, the authors studied the protein levels of SIRT1. This regulates mitochondrial biogenesis by interacting with the transcription coactivator PGC-1α and the removal of damaged mitochondria. It is also associated with the self-renewal of SCs [16,34] and SOD1, which is an important antioxidant enzyme. Compared with the control group, H₂O₂ treatment considerably decreased SIRT1 and SOD1 expression. Compared with the H₂O₂ group, the addition of porous Se@SiO₂ NPs enhanced the expression of SIRT1 and SOD1 in an oxidative stress environment (Figure 3C–E).

Porous Se@SiO₂ NPs enhance mitochondrial activity & dynamics under H₂O₂ stimulation in SCs

Overaccumulation of H₂O₂ can cause oxidative damage, mitochondrial dysfunction and ultimately cell death. Therefore, the authors examined the most important H₂O₂ scavenging enzyme, CAT. Compared with the control group, the addition of porous Se@SiO₂ NPs promoted CAT activity, but the activity of CAT decreased after treatment with H₂O₂. Compared with the H₂O₂ group, the activity of intracellular CAT increased after the addition of porous Se@SiO₂ NPs (Figure 4A). To investigate the effects of porous Se@SiO₂ NPs on mitochondrial respiration, the authors tested the activities of MRC complex I and III and ATP synthase in the respiratory chain (Figure 4B–D). Compared with the control group, the activity of MRC complex I and III, but not ATP synthase, was increased by adding porous Se@SiO₂ NPs. The activity of MRC complex I and III and ATP synthase decreased after H₂O₂ treatment. Compared with the H₂O₂ group, the activity of MRC complex I and III and ATP synthase increased after the addition of porous Se@SiO₂ NPs.

The mitochondrial dynamics and morphology of SCs were studied by confocal laser microscopy. The general morphology of mitochondria was observed by labeling them with a red MitoTracker probe, and cytoskeletal alterations were detected by staining with F-actin to observe the overall shape of the cell and to make sure that the fluorescent probe correctly labeled the mitochondria in cells (Figure 4E & F). Both the control group and the Se@SiO₂ group had numerous mitochondria distributed in the perinuclear areas with high density. Compared with the control group, mitochondrial fragmentation increased, and the density was decreased in H₂O₂-stimulated cells. Compared with the H₂O₂ group, the addition of the porous Se@SiO₂ NPs increased the mitochondrial density, but the mitochondrial structure was still quite different from that in the control group.

Porous Se@SiO₂ NPs promote myogenic differentiation of SCs

The authors further investigated the influence of porous Se@SiO₂ NPs on the myogenic differentiation of SCs under oxidative stress on day 7. MYHC, which is responsible for muscle production in mammals, and MYOD, the early marker of myogenesis, were used to determine the extent of myogenic differentiation. Compared with the control group, the addition of porous Se@SiO₂ NPs increased the expression of MYOD; however, the expression of MYHC and MYOD decreased significantly after H₂O₂ treatment. Compared with the H₂O₂ group, the addition of porous Se@SiO₂ NPs increased the expression of both (Figure 5A–C). Immunofluorescence staining was used to observe myotubes by MYHC-specific antibody and the expression of MYOD (Figure 5D & F). The results were consistent with the results of the western blot analysis. Compared with the control group, the addition of porous Se@SiO₂ NPs can increase the mean fluorescence intensity of MYOD, and exposure to H₂O₂ decreased the fusion index. It also reduced the mean fluorescence intensity of MYOD. Compared with the H₂O₂ group, the addition of porous Se@SiO₂ NPs increased the fusion index and mean fluorescence intensity of MYOD (Figure 5E & G).

Porous Se@SiO₂ NPs accelerate muscle regeneration & reduce collagen deposition in vivo

The authors further verified the therapeutic effect of porous Se@SiO₂ NPs on muscle regeneration after oxidative stress injury in rats. By intramuscularly injecting CTX into rats, the authors established a classical rat model of acute muscle injury (Figure 6A & B). Compared with the control group, porous Se@SiO₂ NPs alone did not affect the normal skeletal muscles (Figure 6C). On day 1 after CTX injection, severe degeneration and necrosis of muscle fibers and exudation of numerous nuclei were seen in the damaged area, indicating successful model construction (Figure 6C). Compared with the control group, more newly formed muscle fibers with nuclei located in the center appeared 7 days after injury, accompanied by neovascularization in the CTX + Se@SiO₂ group (Figure 6D). On day 14, many newly formed muscle cells fused into muscle tubes compared with the CTX group. The regenerated skeletal muscle fibers displayed a more orderly arrangement and had a larger diameter after porous Se@SiO₂ NP treatment. On day 28, the muscle cells differentiated and matured. The transverse diameter of the muscle fibers was close to that of normal muscle, and the nucleus was located under the membrane. In the CTX group, the muscle cells were completely repaired, but they were still different from normal muscle cells because they had a disordered arrangement of muscle fibers. In the CTX + Se@SiO₂ group, the muscle tissue was completely regenerated and showed almost no difference from normal muscle tissue.

At 14 days after injury, images of Masson trichrome staining showed the presence of collagenous tissue, which indicated active collagen deposition in the CTX group compared with the CTX + Se@SiO₂ group (Figure 6E). At 28 days after injury, less positive staining was observed in the gaps around regenerated myofibers in each group. Compared with the CTX group, porous Se@SiO₂ NP treatment reduced collagen deposition in the later stages of muscle regeneration (Figure 6F).

Porous Se@SiO₂ NPs promote the formation of myotubes & increase the number of PAX7-positive cells

The authors assessed skeletal muscle regeneration at protein levels on days 14 and 28. The changes were the same for both MYHC and MYOD. Thus, porous Se@SiO₂ NP treatment can be considered to significantly increase the expression of MYHC and MYOD on days 14 and 28 after muscle injury (Figure 7A–C). This validated the previous results from in vitro experiments. The authors also used immunofluorescence analysis to observe muscle regeneration more intuitively (Figure 7D & E). Double immunostaining for MYHC and dystrophin with nuclear staining of sections indicated that the fluorescence intensity of MYHC increased with time. Local magnification of the image allowed clearer visualization of the double staining of MYHC and dystrophin, and the nucleus was located under the membrane. Treatment with porous Se@SiO₂ NPs could accelerate skeletal muscle regeneration on days 14 and 28 after injury.

PAX7 was specifically expressed in SCs. Double immunostaining for PAX7 and dystrophin with nuclear staining of sections was performed, and the number of PAX7-positive nuclei was determined in the muscle samples. The number of PAX7-positive cells was significantly higher in skeletal muscle after 14 days of action of porous Se@SiO₂ NPs than in the CTX group. (Figure 7F & G).

Discussion

The proliferation and differentiation of SCs are vital for skeletal muscle regeneration after injury and cannot be replaced by unorthodox myogenic precursor cells [35]. To enhance the regenerative ability of a skeletal muscle, it is necessary to recognize the control mechanisms that determine the fate of SCs. Advancements in SCs have revealed that the dynamic metabolism, mitochondrial structure and activity are associated with SCs differentiation, and consequently drives muscle regeneration [12,17,18]. This study was based on exploring the effect of porous Se@SiO₂ NPs on the proliferation and differentiation of SCs by modulating mitochondria under oxidative stress, and it examined the effect of these NPs on muscle regeneration after injury.

Se is an essential micronutrient for the body and is incorporated into selenoproteins in the form of selenocysteine. Selenoproteins have oxidoreductase activity, thereby maintaining the physiological redox balance [36]. Se supplementation attenuates ischemia–reperfusion injury in the skeletal muscle and heart due to its antioxidant and anti-inflammatory properties [37–39]. However, excessive aggregation of Se may have harmful effects on health. There is disagreement on the safe upper limit of Se. According to research, Se at the nanoscale level can reduce the risk of Se toxicity [27]; therefore, the authors of the present study developed porous Se@SiO₂ NPs. The diameter of NPs made them suitable for absorption by cells. In a previous study, porous Se@SiO₂ NPs labeled with fluorescein isothiocyanate were observed by confocal microscopy to enter the cytoplasm and elicit biological effects [26]. The release curve of porous Se@SiO₂ NPs in vitro simulates the levels in the human body, indicating that NPs can achieve sustained release and get effectively distributed in the body [24]. Major organs are not affected by tolerable concentrations of porous Se@SiO₂ NPs [29,40]. With the extension of time due to porous Se@SiO₂ NPs, the problem of local excessive Se accumulation in the tissue is decreased dramatically, avoiding toxicity concerns [41]. The safety of porous Se@SiO₂ NPs was confirmed in this study as well. The MTT assay showed that appropriate concentrations of porous Se@SiO₂ NPs did not inhibit cell proliferation, indicating that porous Se@SiO₂ NPs may be an ideal and safe material for further studies.

When serious muscle trauma occurs, pathological stages, including the destruction phase, repair phase and remodeling phase, occur [29,30]. A large amount of ROS is produced after injury. ROS are a class of molecules that are continuously generated, converted and consumed in all living organisms, and they include H₂O₂, superoxide and hydroxyl radicals, which are generated mainly by electron leakage from the electron transport chain in mitochondria [6,42,43]. For SCs, the role of ROS is extremely complex. The production of ROS is related to the activation and subsequent differentiation of SCs. Moderate levels of ROS are necessary for myogenic differentiation as a signal molecule, while high levels of ROS lead to apoptosis or necrosis. High ROS levels also upregulate NF-κβ, which downregulates MYOD levels, thereby impairing myogenic differentiation [44,45]. In this study, a high concentration of H₂O₂ was used to induce oxidative stress injury in SCs. The porous Se@SiO₂ NPs significantly decreased mitochondrial ROS levels under H₂O₂ exposure. Thus, porous Se@SiO₂ NPs protected SCs from H₂O₂-induced cell damage and senescence. The fate of SCs in myogenic differentiation is regulated by MYOD at an early stage, and eventually cells exit the cell cycle and merge into the regenerated myotubes [46]. The level of myogenic differentiation was based on the expression of MYOD and MYHC; the expression of both these markers was reduced after H₂O₂ treatment. However, porous Se@SiO₂ NPs suppressed the inhibitory effect of H₂O₂ on MYHC and MYOD. Thus, porous Se@SiO₂ NPs could counteract the H₂O₂-induced inhibition of myogenic differentiation in SCs.

Excessive levels of ROS are detrimental to mitochondria in SCs because they could target mitochondrial DNA and mitochondrial function, causing swelling and disruption of mitochondria [47,48]. Therefore, it is necessary to promote cellular homeostasis for protecting mitochondria. Mitochondrial damage and dysfunction can further lead to damage to the entire cell. To protect SCs from the underlying damage of excess ROS, mitochondria have their antioxidant systems, particularly superoxide dismutase and glutathione peroxidase, which scavenge ROS to sustain intracellular stability [45]. Thus, antioxidants are essential for mitigating cellular oxidative damage through mitochondria. Appropriate doses of antioxidants can maintain the level of ROS at an optimal level, so that ROS can act as physiological signaling molecules [25]. In this study, the porous Se@SiO₂ NPs protected the morphological and functional stability of mitochondria by increasing the number of mitochondria and promoting mitochondrial respiration. To illustrate the potential antioxidant mechanism of porous Se@SiO₂ NPs, the authors evaluated the expression of SOD1, which acts as an antioxidant enzyme, and SIRT1, which regulates mitochondrial genesis and removes damaged mitochondria. The results demonstrated that porous Se@SiO₂ NPs upregulated the expression of SIRT1 and SOD1. This was consistent with previous studies that demonstrated that porous Se@SiO₂ NPs reduce ROS levels by promoting the production of major endogenous antioxidants, including superoxide dismutase and glutathione [29].

The effect of porous Se@SiO₂ NPs was next investigated in a CTX-induced TA injury model. The results were consistent with the results of the in vitro experiments. Porous Se@SiO₂ NPs upregulated the expression of MYHC and MYOD in the injured muscle on days 14 and 28. Morphological changes in muscle regeneration were observed in hematoxylin and eosin-stained sections, and treatment with porous Se@SiO₂ NPs promoted more rapid muscle fiber regeneration and more regular arrangement between muscle fibers. Interestingly, porous Se@SiO₂ NPs significantly increased the number of PAX7-positive cells in the injured region on day 14 compared with the CTX group, but there was no statistical significance on day 28. The authors speculate that this is because of the complete regeneration of the muscle on day 28 and because no additional SCs were required for skeletal muscle regeneration. Skeletal muscle regeneration is accompanied by fibrosis development, muscle fibrosis and oxidative stress, which are closely related to each other [49,50]. SCs may also contribute to fibrosis through the WNT signaling pathway [51]. In addition to a longer time for repair, fibrotic repair also disrupts the mechanical properties of the muscle and makes the affected muscle more vulnerable to reinjury [52]. The authors confirmed that porous Se@SiO₂ NPs attenuated collagen deposition after injury, but its detailed mechanism is not known.

Although this study confirmed that porous Se@SiO₂ NPs promote skeletal muscle regeneration by modulating mitochondria to attenuate oxidative damage, additional studies are necessary to elucidate the potential impact of porous Se@SiO₂ NPs on other mechanisms regulating mitochondrial homeostasis such as mitochondrial autophagy in SCs. Furthermore, the biological effects of NPs depend on their targeting efficiency, and further improvements in material properties will enable them to enter the mitochondria more effectively to exert their biological effects. In addition, acute degenerative necrosis of the skeletal muscle induced by CTX in this experiment is still somewhat different from skeletal muscle injuries seen in the clinic. Porous Se@SiO₂ NPs should be further explored to investigate their potential therapeutic effects.

Conclusion & future perspective

This study provides initial evidence that porous Se@SiO₂ NPs significantly promote the proliferation and myogenic differentiation of SCs under oxidative stress by scavenging mitochondrial ROS and improving mitochondrial activity (Figure 8). Treatment with porous Se@SiO₂ NPs accelerated skeletal muscle regeneration and reduced fibrosis deposition after injury. Our results reveal the application potential of porous Se@SiO₂ NPs in skeletal muscle diseases or other pathologies related to mitochondrial dysfunction. Further research is necessary in the future to improve the targeting efficiency of NPs; and safety studies in large animal models to facilitate better clinical translation.