Duchenne muscular dystrophy is a hereditary muscular disease involving degeneration (i.e. atrophy and loss of muscle fibres) of skeletal muscles, including the diaphragm, and progressively severe functional decline. A previous study shows Polycan, a type of β-glucan derived from the black yeast Aureobasidium pullulans (SM-2001), promotes osteogenicity and bone loss, and possesses anti-inflammatory activity to induce inflammatory cytokines in human immune and cancer cells.
In this study, we evaluated changes in exercise load behaviour measurements and changes in muscle-related physiological indicators following oral administration of Polycan in mdx mice, an experimental animal model of Duchenne muscular dystrophy.
In mdx mice, Polycan prevented weight loss and thickness of skeletal muscle. In addition, by monitoring increases in running time of mice on treadmills and performing a grip strength test, we confirmed reduced muscle function was recovered to some extent after administering Polycan to mdx mice. In addition, we confirmed that Polycan significantly altered mRNA expression in a concentration-dependent manner, whereby myogenic transcription factors (MyoD, Myf5 and Myogenin) increased and FoxO3α, MuRF1 and Atrogin-1 decreased. We aimed to investigate the mechanism of action in Polycan on energy metabolism of p-AMPK, SIRT1 and PGC1α with apoptosis expression levels as factors related to signalling pathways. Expression ratios of cleaved-caspase-3/caspase-3 and Bax/Bcl-2 in the Polycan extract-administered group increased compared with the control group.
These results demonstrate that Polycan can improve and protect muscle atrophy by preventing apoptosis via pathway regulation related to myogenic transcription factors and energy metabolism in mdx mice.
Duchenne muscular dystrophy is the most common sex chromosome recessive genetic disorder in paediatric muscle, caused by a defect in the dystrophin gene (dys) located on chromosome Xp21 (Emery 2002). Duchenne muscular dystrophy (DMD) is a hereditary muscle disease that occurs in the absence of dystrophin, a protein important for maintaining stable cell membrane structure. In this disease, skeletal muscle, including the diaphragm, is degenerated (muscle fibre atrophy) and disappears, potentially showing gradual and serious decline in function (Alter et al. 2006). It affects approximately 1 in 3500 male newborns and typically begins between ages 3 and 5 and renders the child unable to walk by the age of 12. Furthermore, most patients struggle with breathing due to weakened diaphragmatic muscles and before the age of 20 will die from heart failure (Clerk et al. 1993). Since development of genetics revealed the existence of dystrophin, the physiological role of dystrophin and its interaction with other glycoproteins have occurred in the muscular membrane. Unfortunately, more than 100 years have passed since Duchenne and Meryon reported clinical and muscular histological findings in patients with DMD, but no cure has since been found. Typically, ultra-low intensity exercised is prescribed, primarily focusing on symptom relief and to prevent further complications (Barton et al. 2002; Blake et al. 2002).
Recently, the proportion of elderly people aged 65 and over is increasing worldwide and clinical follow-up studies show decrease in muscle mass and bone density with the progress of ageing is a typical physiological change. Conversely, economic and social problems continue to emerge (Jagoe and Goldberg 2001). The silence of muscle loss refers to a disease in which muscle mass, strength and muscle function all decrease with age. Muscle atrophy is caused by several reasons, including ageing, undernourishment and chronic disease, and involves many intracellular changes, such as regulation of protein synthesis, protease activation, ubiquitin conjugation and autophagy (Ammar et al. 2016; Kandarian and Jackman 2006). Since such muscular atrophy can cause poor quality of life and disease worsening, resulting in increased mortality, studies continue to investigate its mechanisms with no current clear mechanism known. Muscle injury can be caused by strenuous exercise, muscle and skeletal disorders and can occur with limited movement, such as with patients in intensive care (Kim et al. 2013). Here, muscle mass is reduced, muscle fibre diameter (Cross Sectional Areas) may decrease and satellite cell activity, which are muscle stem cells, also decreases (Kasper et al. 2002). In addition, muscle strength is reduced and musculoskeletal ultrastructure is disrupted. In particular, patients with muscular dystrophy in the absence of cytoskeletal protein dystrophin are vulnerable to physical muscle damage due to contraction and muscle damage from chemical factors, such as oxidative stress (Matsumura and Campbell 1994). Therefore, patients with muscular dystrophy experience muscle atrophy, disappearance and regression from muscle injury (Haller et al. 1983). Such symptoms can be due to increased proteolysis in the human body. Of the numerous proteolytic pathways in the human body, two main pathways exist. First, the autophagy-mediated proteolytic pathway performs 10% of proteolysis (Rayavarapu et al. 2013). Self-predation is a mechanism manifested in response to various internal and external stress developments and degrades damaged proteins and organelles by fusion with lysosomes (Yang and Klionsky 2009). The remaining 80–90% of proteolysis occurs via ubiquitin and the ubiquitin–proteasome system (UPS) (Rayavarapu et al. 2013). Damaged protein is labelled with ubiquitin, whereby ubiquitinated protein is transferred to macromolecular proteasome and degraded. Following completed degradation, ubiquitin separates again and binds to other damaged proteins, resulting in most proteolysis in the human body (Ehlers 2003). These UPSs play an important role in degrading damaged proteins and maintaining homeostasis in the human body (Rayavarapu et al. 2013). However, overactivation in response to muscle damage and associated increased muscle contraction protein damage triggers excessive degradation of muscle contraction protein and promotes proximal axis and musculoskeletal disorder (Hilt and Wolf 2004).
In recent years, due to increased awareness of natural products, there is a demand for functional foods using natural products, ingredients separated from natural products, bioactive substances and food materials and functional foods that use bioactive substances as alternative medicine (Kim et al. 2021). As the number of commercial medicinal plants increases, the food industry's use of natural products, including medicinal plants, tends to increase. Research on natural products that impact human activity is an active field. Among the natural extracts attracting attention in immune potentiation, β-glucan is non-resistant and shows anti-inflammatory and anti-cancer effects in addition to immune modulators applicable in various bioactive fields (Bacha et al. 2017; Novak and Vetvicka 2008; Smiderle et al. 2008). In particular, Polycan, which is a type of β-glucan derived from the black yeast Aureobasidium pullulans SM-2001, promotes bone formation and anti-inflammatory processes (Shin et al. 2007; Wang et al. 2016). Furthermore, it can reduce IL-1β, IL-6 and TNF-α, which are inflammatory cytokines (Chan et al. 2009) and increase anti-inflammatory activity when combined with non-steroidal anti-inflammatory agents (Choi et al. 2015). Therefore, in this study, we explore how Polycan extract isolated from the black yeast Aureobasidium pullulans can facilitate changes in exercise load and improve physiological indicators of muscles in mdx mice.
Materials and methods
Preparation of Polycan extract
The Polycan used in this experiment was β-glucan fermented with the black yeast Aureobasidium pullans SM-2001. According to previous studies, β-1,3/1,6-glucan (β-1,3 linkage 68: β-1,6 linkage 32) and an average molecular weight of 2.6 × 105 Da. The β-glucan used in the experiment was provided by Glucan (Jinju, Korea) and used in the experiment. Hereinafter, the extract is referred to as “Polycan”.
In this study, 4-week-old mdx mice (C57BL/10ScSn-Dmdmdx/J, N = 32) and control mice (C57BL/10SnJ, N = 8; Jackson Laboratory, Bar Harbor, ME) were used in the same environment (12: 12 h light/dark cycle). It was bred with a standard diet and a free water intake at 22 °C. Eight mice (CTL) and eight mdx mice were divided into two groups, eight each, a control group (mdx), Polycan -administered (Polycan 250, 500 mg/kg) experimental group, and a positive control (Sc 500 mg/kg) group. During the experiment, mice were bred in a 12-h light–dark cycle environment, and the same diet was fed unlimitedly to the experimental and control groups. All the animal procedures were accepted and approved by the Korea Institute of Oriental Medicine-Daegu-Institutional Animal Care and Use Committee (KIOM-IACUC, reference number 21-053) and conducted following the US guidelines (NIH publication #83-23, revised in 1985).
Measurement of body weight
After confirming overall health and behavioral changes in all mice prior to physiologic testing, body weight was measured and recorded weekly every Monday. The body weight of 4 weeks after birth and the body weight of 9 weeks after birth, which is the end of the experiment, were measured, and the weight gain ratio was compared among each group.
A treadmill test was used to compare the endurance of the experimental and control groups. A Columbus Instrument Rodent Treadmill (Columbus Instruments, Ohio, USA) installed on a horizontal surface was run at a speed of 10 mpm for 30 min. A 1.0 mA electrical wire was placed at the back of the belt to keep the mouse running so that a light electrical stimulus was applied when the mouse was pushed to the back of the belt. When the mouse became tired, it stopped running and showed sitting on the electrical stimulation line, it stayed on the electrical stimulation line for 10 s or more, it was regarded as a resting period. The number of 30-min rest periods was recorded for each mouse. All exercises were performed between 11:00 am and 2:00 pm to reduce the difference due to changes in biological rhythms in both the control group of C57 mice control group and experimental group of mdx mice.
Grip strength meter
A grip strength meter (Ugo Basile, Comerio, Italy) was used to examine the muscle strength of the distal part of the mouse. This device applies the tendency to instinctively capture the object in front of the rat when the rodent is pulled backwards, after the rat captures the rod connected to the force transducer and then the rat. A device designed so that if the tail is carefully pulled back, the rat resistance cannot be overcome and the maximum vicious force at the moment of missing the rod is recorded on the computer. The average of the values obtained by five discontinuous tests was used as a bad force, and the test was performed before the Treadmill exercise to reduce errors due to muscle fatigue, and the mice were stabilized for 10 min or more before the experiment was started.
Extraction of muscle tissue
After euthanasia on the day the experiment was completed, each group immediately removed the foreground bone muscle and measured skeletal muscle weight and thickness on an electronic balance. Immediately after that, it was quickly frozen in liquefied nitrogen and stored at − 80 °C. The right foreground bone muscle was used to measure muscle fibre cross-sectional area for immunohistochemical and the left foreground bone muscle was used to quantify muscle regulators through Western blot.
After removing the muscle tissue by dividing it into the thigh muscle and TA (tibialis anterior) each tissue was immersed and fixed in 4% paraformaldehyde for 24 h, and after undergoing a paraffin embedding process. It was sliced into a thickness of 3 μm using a microtome to prepare a tissue slide. For hematoxylin and eosin (H&E) staining of each tissue, tissue slides were stained with hematoxylin for 5 min and then with eosin for 2–3 min. In addition, each tissue slide was oxidized with 2% periodic acid solution for 10 min and stained with Schiff solution at 37 °C for 30 min. Stained tissue slides were observed for histological changes using a microscope.
Changes of serums
Serum hematological markers were analyzed to examine the changes in various serum factors related to drug and muscle loss. That is, after the experiment was completed, whole blood from the abdominal vena cava was collected from the mouse blood. Plasma was separated from the blood, and used in the experiment. Glutamic oxaloacetic transaminase (GOT), glutamic oxaloacetic transaminase (GPT), glucose, total protein, cholesterol, triglyceride, HDL, LDL, UREA, CREA, creatine kinase (CK) are blood biochemical automatic analyzers (XL-200; Germany) was analyzed using.
Reverse transcription-quantitative PCR (RT-qPCR)
Total RNA of mouse tissues prepared under muscle atrophy conditions was isolated and quantified using TRIzol Reagent (Invitrogen, Carlsbad, CA, USA), followed by the same amount of total RNA in the first-strand cDNA synthesis kit (first-strand cDNA synthesis kit). Converted to cDNA using Applied Biosystems Inc., Framingham, MA, USA). Total RNA extraction and cDNA synthesis were conducted using the TRIzol Reagent and AccuPower® CycleScript RT PreMix (Bioneer, Daejeon, Korea), respectively. A total of 1 μg RNA was reverse-transcribed into cDNA, qPCR oligonucleotide primers for macrophage cell cDNA are indicated in Table 1. The qPCR reactions were performed in triplicate 20 μL reactions with 1 μL of 0.3 μM of the forward and reverse primer each, 10 μL of the AccuPower® 2 × Greenstar qPCR master mix, 5 μL of template DNA, and 3 μL of RNase-free water. The PCR cycle was as follows: 95 °C for 10 min, 40 cycles. At the each cycle end, a melting curve analysis was conducted to confirm that a single product per primer pair was amplified. Amplification and analysis were performed using the QuantStudio 6 Flex Real-time PCR System (Thermo Fisher, Waltham, Massachusetts, USA), and each sample was compared using the relative CT method. Fold changes in gene expression were determined relative to the blank control after normalization to GAPDH expression using the 2 − ΔΔCt method.
Western blot analysis
The proteins in mouse tissues prepared under muscle atrophy conditions were extracted with RIPA lysis buffer containing a protease and phosphatase-inhibitor cocktail (Roche, Basel, Switzerland). The protein was separated by centrifuging this at 12,000 rpm for 20 min. The concentration of each isolated protein was quantified by a protein assay solution, then 30 μg of protein was mixed with 5 × sample buffer and separated by 8–15% SDS-PAGE. The proteins on the isolated gels were transferred to NC membranes and each membrane was blocked with 5% BSA at room temperature for 1 h. The primary antibody related to differentiation, cell death, and muscle loss was added to the membrane and reacted at 4 °C. overnight, and then washed three times with TBS containing 0.05% Tween. After re-inserting the anti-IgG conjugated HRP antibody into the Membrane, the reaction was carried out at room temperature for 1 h, washed three times with TBS (1 × TTBS) containing 0.05% Tween, and the ChemiDocTM touch imaging system (BioRad, California, USA) was used with the ECL solution.
All the results are presented as mean ± SD (or SEM). One-way analysis of variance using GraphPad Prism version 5.03 for Windows (GraphPad Software Inc., San Diego, CA, USA) was performed for multi-group comparisons. T tests were applied to analyze the significance of the variations among the groups.
Effect of Polycan on body weight and muscle dystrophy in mdx mice
Body weight changes in 4-week-old control mice (CTL), mdx mice (mdx), Polycan-treated administered mice (mdx + Polycan 250 mg/kg, mdx + Polycan 500 mg/kg) and positive control-treated mice (mdx + Sc 500 mg/kg) are shown in Fig. 1A. There was no significant difference in body weight between the five groups at the end of the study in Polycan-treated animals. In contrast, we found body weight of mice in the CTL and mdx groups significantly decreased from the 4th to 8th week after feeding concluded. Results of body weight measurements in the 1st and 8th week of drug administration showed the weight gain ratio could be calculated and compared among groups. Weight gain in the control group (CTL) was 12.0 ± 4.1 and showed an increase of 4.2 g (Fig. 1B). In contrast, weight gain in the mdx group was 6.9 ± 0.7 g. In the mdx + Polycan 250 mg/kg, mdx + Polycan 500 mg/kg and mdx + Sc 500 mg/kg groups, body weight was 12.5 ± 2.1 g, 17.9 ± 1.5 g and 9.3 ± 2.0 g, respectively, and body weight increase in the drug-administered groups was similar to the CTL group; here, increases in the mdx group was significant. Time to exhaustion from treadmill exercises for the CTL and mdx groups were 1962.0 ± 61.4 and 1430.4 ± 196 s, respectively, which represented a significant decrease. Time recorded for the mdx + Polycan 250 mg/kg and mdx + Polycan 500 mg/kg groups was 1567.9 ± 164.8 and 1661.3 ± 62.4 s, respectively, while time recorded for the Sc 500 mg/kg group was 1600.2 ± 46 s (Fig. 1C). For grip strength tests performed on the same mice, the CTL group showed a significant decrease in adverse force of 86.9 ± 6.5 g compared with 139.8 ± 15.4 g. In contrast, the mdx + Polycan 250 mg/kg, mdx + Polycan 500 mg/kg and Sc groups showed vicious forces of 109.9 ± 12.7 g, 125.2 ± 9.5 and 91.6 ± 11.0 g, respectively, which were statistically significant compared with the mdx group. Therefore, Polycan-feeding is likely to alleviate muscle dysfunction in mdx mice (Fig. 1D).
Effect of Polycan on changes in histology, thickness and weight
We observed a significant difference in muscle thickness following Polycan- and Polycan-feeding treatments (P < 0.01) in the mdx group compared with the CTL group. However, a significant increase in thickness and weight of mouse thigh muscle was observed in Polycan- and Sc-treated groups compared with the mdx group. Polycan (250 and 500 mg/kg) showed a dose-dependent inhibitory effect on weight loss of thigh muscles, which was greater than the effects seen in the positive control group (Fig. 2A & B). In general, an important feature of skeletal muscle atrophy is reduced size and number of muscle fibres. Therefore, we performed H & E staining of skeletal muscle tissue to compare histological characteristics from morphological changes in muscle fibre. Compared with the CTL group, muscle fibres in the mdx, Polycan-feeding mdx and SC-feeding mdx were atrophied and muscle fibre thickness decreased. In comparison, we observed increased muscle fibre thickness in the Polycan and Sc-treated groups (Fig. 3).
Results from blood tests are shown in Table 2. We found statistically significant results between mdx and CTL groups, as indicated by *p > 0.05, 0.01 and 0.001, and between Polycan- or Sc-treated groups and the mdx group, as indicated by #p > 0.05, 0.01 and 0.001. No haematologic markers differed among groups. Creatine kinase (CK) significantly increased in the mdx group compared with the CTL group and significantly decreased in Polycan-treated and Sc-treated groups compared with the mdx group.
Changes in gene expression associated with muscle production and muscle cell proteolysis
In mdx mice, mRNA expression in muscle tissue confirmed the effect of Polycan on muscle cell production and proteolytic factors. Relative mRNA expression levels of Myf5, MyoD, myogenin and MEF2, which are transcription factors involved in muscle cell production, are shown in Fig. 3. mRNA expression level decreased overall in the mdx group compared with the CTL group. Furthermore, gene transcript level increased significantly in Polycan-treated groups (250 and 500 mg/kg) in a concentration-dependent manner compared with the Sc-treated group (Fig. 3A). Furthermore, we observed changes in mRNA expression levels of FoxO3α, MuRF1 and atrogin-1, whereby expression in the mdx group was approximately twice as high compared with the control group (CTL). In contrast, mRNA expression level was significantly lower in a concentration-dependent manner following 250 and 500 mg/kg administration of Polycan extract.
Effects on pathways related to energy metabolism regulation
When AMPK is activated in tissues, such as the liver and skeletal muscle, fatty acid oxidation and glucose absorption increase. SIRT1 is an essential factor for AMPK activation and contributes to PGC1a metabolism. To investigate the Polycan mechanism of action on energy metabolism, we confirmed expression levels of p-AMPK, SIRT1 and PGC1α, which are factors related to signal transduction pathways. As expected, mRNA expression levels of SIRT1, p-AMPK and PGC1a decreased significantly in the mdx group, but were restored in a concentration-dependent manner in the two Polycan-treated groups (Fig. 4A). In contrast, in muscle, mRNA expression levels of genes in p-AMPK, SIRT1 and PGC1α pathways significantly decreased in the mdx group. In the Polycan-treated group, mRNA expression level of p-AMPK, SIRT1 and PGC1α increased for all concentration ranges together with the SC-treated group (Fig. 4B). These results indicate that Polycan has a protective effect on the energy metabolism-related pathway control in the muscular dystrophy model.
Effects on muscle protein synthesis pathway and apoptosis
A decrease in Akt signal activity and cell death due to decreased mitochondrial function in muscle cells are typical phenomena that induce muscle hypotrophy. The Akt signal activity is a major regulator of skeletal muscle hypertrophy. To investigate the mechanism of inhibitory effects of Polycan on muscular atrophy, we determined expression of Akt and PI3k proteins, which are involved in protein synthesis, using western blotting. As expected, protein expression level of p-Akt and PI3k were significantly lower in the mdx group, but higher in the Polycan 250 mg/kg and Polycan 500 mg/kg treatment groups, compared with controls (CTL; Fig. 5A). Protein expression levels of bax, bcl-2, caspase-3 and cleaved-caspase-3, which are major regulators of apoptosis, were also examined and showed Bcl-2 expression increased and Bax expression decreased. In addition, when ratios of cleaved-caspase-3/caspase-3 and Bax/Bcl-2 expression following treatment with Polycan extract were compared with the CTL group, protein expression related to cell death changed in a concentration-dependent manner, showing significant recovery (Fig. 5B).
DMD is a typical hereditary neuromuscular disease with a prevalence of approximately 4 in 100,000 people and an incidence of 1 in 3,500 (Clerk et al. 1993). It occurs due to a genetic abnormality at the p21 site of the X chromosome preventing dystrophin expression, a cell membrane protein (Barton et al. 2002). The absence of dystrophin, which connects the cytoskeleton and extracellular matrix of muscle fibres and contributes to muscle fibre protection and intracellular/extracellular signal transduction, causes skeletal muscle atrophy, wasting and degeneration in patients with DMD. There is also evidence of muscle replacement for fat and connective tissue, pseudohypertrophy, fibrosis, inflammation and necrosis, resulting in muscle weakness. Here, mdx (C57BL/10ScSn-Dmdmdx/J) mice are a useful representative animal model for Duchenne-type muscular dystrophy and show an early stop codon (codon) at exon 23, the site that ultimately produces the protein in DNA and restricts expression of the cytoskeletal protein dystrophin (Coley et al. 2016). Therefore, mdx mice show pathological symptoms similar to patients with muscular dystrophy and are used as a representative animal model for muscular dystrophy (Dunckley et al. 1998). However, the pathological symptoms of mdx are less critical than pathological symptoms of patients with human muscular dystrophy because mdx mice, which show accelerated muscle regression from 3 to 4 weeks after birth, actively undergo muscle regeneration at 3–12 months. This difference is due to weight differences between humans and mice (Grounds and Torrisi 2004). However, differences in pathological symptoms exist and pathological progressions in skeletal and respiratory muscle are almost similar and genetically homologous (Hartel et al. 2001). In addition, mdx mice show muscle damage similar to patients with muscular dystrophy, which results in fibrotic hypertrophy and necrosis, as well as destruction of muscle structure. Owing to their study model advantages, mdx mice are widely used in preclinical studies of muscular dystrophy (Barton et al. 2002). Studies using these mdx mice have investigated potential therapeutic agents for cancer cachexia (Lee et al. 2022), skeletal muscle wasting (Marquez et al. 2020) and improving muscle mass and strength in tumour induction (Cheng et al. 2021), reducing oxidative stress (Resveratrol) and muscle fibre regeneration (Evans et al. 2010).
In this study, we evaluated muscle improvement and its efficacy in Polycan-treated mdx mice based on previous study outcomes (Hwang et al. 2021). First, when examining changes in body weight, there was no clear increase in the mdx group compared with the control group (CTL), whereas the Polycan- and Sc-administered groups showed a significant increase compared with the mdx group. Weight gain in mdx mice was also prominent as per previous studies (Yang et al. 2017). The treadmill test and grip test functionally evaluated muscle strength, but in an experiment comparing treadmills, values in the Polycan-treated group were significantly different to those in the mdx group. In addition, for the grip strength test conducted on the same mice, we found reduced force was recovered to some extent following Polycan extract administration to mdx mice compared with the CTL group (Fig. 1D).
Previous studies have consistently demonstrated elevated levels of CK in patients with muscular dystrophy and mdx mice, which are known to cause muscle loss (Bogdanovich et al. 2002). Concentrations of CK in mdx mice significantly increased compared with the CTL group and confirmed concentrations significantly decreased following Polycan extract administration to mdx mice (Table 2).
Muscle atrophy is caused by degraded muscle protein by activating the ubiquitin–proteasome pathway (Jagoe and Goldberg 2001; Marinovic et al. 2007). MyoD and Myf5, which are classified as primary myogenic factors among myogenic transcription factors, help induce muscle blast cells, while myogenin and Myf-6 are secondary factors. Myocyte enhancer factor 2 (MEF2) helps differentiate myogenic cells into root canal cells (Bentzinger et al. 2012; Cheng et al. 2021). In addition, myostatin, a substance in transforming growth factor-β (TGF-β) family that lowers Akt/mTOR/p70S6K signalling and suppresses skeletal muscle hypertrophy, is expressed in muscles. MAFbx/atrogin-1, an F-box type E3 ligase specifically expressed in skeletal muscle, and MuRF1, a ring finger type E3 ligase that increases under various conditions, induce the proximal axis, resulting in increased cell density. This promotes degradation of myogenin and MyoD involved in myogenic disorders through cycle suppression and activation of muscle-specific gene expression (Kandarian and Jackman 2006; Lawler et al. 2003). Therefore, in this study, we confirmed how the efficacy of Polycan affects expression of transcription factors involved in producing and degrading muscle during muscular atrophy. As shown in Fig. 3, muscle cell production in the mdx group confirms that mRNA expression of MyoD, myogenin, MEF2 and Myf5 decreased compared with the CTL group and that Polycan treatment facilitated increases in gene expression of these factors. Among the muscle-specific transcription factors involved in protein degradation of muscle cells, forkhead box O (FoxO) activity induced autophagic protein activation to promote the downstream signal transduction system and FoxO activity. Finally, inhibition of intracellular mTOR expression affects cell proliferation (Ardite et al. 2004).
AMPK is an important signal transduction enzyme that functions as a sensor to maintain homeostasis of intracellular energy and the human body regulates sugar, lipid and protein metabolism by sensing intracellular energy states based on the AMPK to ATP ratio. In addition, AMPK can increase expression of energy metabolism regulators, such as PGC1α, Sirt1, NRF1 and TFAM, to promote glycolysis and depletion and stimulate fatty acid oxidation to promote intracellular ATP synthesis. PGC1α is an important factor that regulates energy metabolism in muscle, regulates oxidation of fatty acids and increases energy levels by promoting mitochondrial biosynthesis, which in turn activates transcription factors that affect regulation and respiratory activity (Kovacheva et al. 2010; Park et al. 2020). In this study, we confirmed changes in gene expression and protein expression of PGC1α, Sirt1 and AMPK to confirm the energy-increasing effect of Polycan extract and found expression of these regulators increased following treatment with Polycan extract. Notably, PGC1α and SIRT1 expression increased significantly depending on Polycan extract treatment concentration compared with the control group (CTL). These results show Polycan extract can promote mitochondrial energy production by regulating expression of energy metabolism regulators in muscle cells.
Suppressed self-predatory action in aged skeletal muscle reduces protein accumulation modified by impaired protein conversion, ultimately diminishing function of various biological pathways required for homeostasis and survival of muscle fibres. It regulates self-predation by activating PI3K and Akt signals via the insulin-like growth factor 1/PI3K/Akt signalling pathway and regulating mTOR via its involvement in ageing that cause degeneration and weakening (Kovacheva et al. 2010; Pavlicek et al. 2013). By analysing protein expression of PI3K/Akt and mTOR in this study, we confirmed Polycan extract improved reduced protein expression of p-Akt, p-PI3k and p-mTOR in the mdx group. This revealed that Polycan suppresses the proximal axis of the PI3K/Akt/mTOR signalling pathway.
Apoptosis is an important physiological process for tissue regeneration, homeostasis maintenance and growth, but the physiological changes that appear with age are characterised by rapid acceleration of muscle cell apoptosis. This is strongly correlated with muscle loss that appears in the elderly population, but the mechanism through which apoptosis is accelerated is not yet fully known. However, chronic inflammation, such as the abovementioned oxidative stress, is indeed involved as is myonuclear apoptosis, which is characterised by a reduced number of multinuclear muscle cells (Cea et al. 2016; Spencer et al. 1997). Caspases are the most important factors in apoptosis and play essential roles. Specifically, caspase-3 is known as the primary effector caspase that destroys various substrates. When cell death occurs in the cell membrane, activity of pro-apoptotic factor Bax (bcl-2-associated X protein) increases and expression of the relatively anti-apoptotic factor Bcl-2 is suppressed, thereby suppressing expression of the relatively anti-apoptotic factor Bcl-2 and facilitating cytochrome c release. The released cytochrome c forms an apoptosome with pro-caspase-9 and apoptotic protease activator (APAF)-1, which triggers PARP digestion by increasing activity of caspase-3 and DNA. Following this, apoptosis occurs due to increased DNA fragmentation. In this study, we investigated expression levels of Bax/Bcl-2 and cleaved-caspase-3/caspase-3 after administering Polycan extract to determine its involvement in muscle cell apoptosis in mdx mice. Our results showed that cell death was suppressed.
Furthermore, our results indicate that Polycan could affect activation of muscle cell-producing transcription factors and muscle cell proteolytic factor suppression, which may be linked to ameliorated muscular atrophy and protection of muscular status in mdx mice, a model of muscular dystrophy.
This study demonstrated that Polycan resulted in activated myogenic transcription and inhibited muscle proteolytic factors, resulting in improvement and protection of muscle atrophy in the mdx mouse, a model of muscular dystrophy.
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Conflict of interest
Young-Suk Kim declares that he/she has no conflict of interest. Jong-Min Lim declares that he/she has no conflict of interest. Jae Suk Shin declares that he/she has no conflict of interest. Hyun Jun Kim declares that he/she has no conflict of interest. Kwang-Il Park declares that he/she has no conflict of interest. Tae Woo Oh declares that he/she has no conflict of interest.
All the animal procedures were accepted and approved by the Korea Institute of Oriental Medicine-Daegu-Institutional Animal Care and Use Committee (KIOM-IACUC, reference number 21-053) and conducted following the US guidelines (NIH publication #83-23, revised in 1985).
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Kim, YS., Lim, JM., Shin, J.S. et al. Extracellular polysaccharides purified (Polycan) from Aureobasidium pullulans SM‑2001 improves pathophysiology of dystrophin-deficient mdx mice. Mol. Cell. Toxicol. 18, 285–297 (2022). https://doi.org/10.1007/s13273-022-00245-x
- Duchenne muscular dystrophy (DMD)
- Aureobasidium pullulans (SM-2001)
- Mdx mice
- Anti-atrophic effect