Morin improves dexamethasone-induced muscle atrophy by modulating atrophy-related genes and oxidative stress in female mice

ABSTRACT

This study investigated the effect of morin, a flavonoid, on dexamethasone-induced muscle atrophy in C57BL/6J female mice. Dexamethasone (10 mg/kg body weight) for 10 days significantly reduced body weight, gastrocnemius and tibialis anterior muscle mass, and muscle protein in mice. Dexamethasone significantly upregulated muscle atrophy-associated ubiquitin ligases, including atrogin-1 and MuRF-1, and the upstream transcription factors FoxO3a and Klf15. Additionally, dexamethasone significantly induced the expression of oxidative stress-sensitive ubiquitin ligase Cbl-b and the accumulation of the oxidative stress markers malondialdehyde and advanced protein oxidation products in both the plasma and skeletal muscle samples. Intriguingly, morin treatment (20 mg/kg body weight) for 17 days effectively attenuated the loss of muscle mass and muscle protein and suppressed the expression of ubiquitin ligases while reducing the expression of upstream transcriptional factors. Therefore, morin might act as a potential therapeutic agent to attenuate muscle atrophy by modulating atrophy-inducing genes and preventing oxidative stress.

Graphical Abstract

Diagram showing anti-atrophic effect of morin. Morin attenuated dexamethasone-induced Klf15 expression and oxidative stress that upregulates Cbl-b followed by atrogin-1 and MuRF-1 via FoxO3a dephosphorylation.

C57BL/6J mouse, glucocorticoid, oxidative stress, ubiquitin ligase, muscle atrophy

Glucocorticoids are commonly used to alleviate inflammatory diseases. Prolonged or increased administration of glucocorticoids is associated with various side effects including osteoporosis, adrenal gland dysfunction, hyperglycemia, and muscle atrophy (Sato et al.2018). Dexamethasone is a potent synthetic glucocorticoid that stimulates muscle atrophy by rendering muscle protein degradation and exhausting protein synthesis (Schakman et al.2013). The proteolysis induced by dexamethasone is primarily mediated through the activation of the ubiquitin-proteasome system (UPS). Generally, following binding to the glucocorticoid receptor, glucocorticoids induce the expression of Krüppel-like factor 15 (Klf15), a member of the zinc finger transcription factor family of proteins. The upregulation of Klf15 has been found to induce the transcription of genes responsible for muscle atrophy, such as muscle atrophy F-box protein 1 (MAFbx1/atrogin-1) and muscle ring finger protein-1 (MuRF-1), via forkhead box O3a (FoxO3a) (Shimizu et al.2011).

Meanwhile, dexamethasone generates oxidative stress in mouse C2C12 and rat L6 skeletal muscle cells (Ohtsuka et al.1998; Oh et al.2021; Ulla et al.2021; Kim et al.2022) and it can initiate protein degradation and depress protein synthesis leading to muscle atrophy (Gomes-Marcondes and Tisdale 2002; Ábrigo et al.2018). We and others, previously reported that oxidative stress remarkably upregulates the ubiquitin ligase casitas B-lineage lymphoma proto-oncogene-b (Cbl-b), which induces muscle atrophy by stimulating the degradation of insulin receptor substrate (Uchida et al.2018; Oh et al.2021). This event leads to disturbed insulin-like growth factor-1 (IGF-1) signaling followed by the increased expression of atrogin-1 and MuRF-1 through FoxO3a dephosphorylation (Nakao et al.2009). Thus, both Klf15 and oxidative stress might be suitable targets to prevent dexamethasone-mediated muscle atrophy.

Morin (3,5,7,2′,4′ pentahydroxyflavone) is a plant-based, yellow-colored bioflavonoid widely distributed in different species of the Moraceae family of plants (Caselli et al.2016). Morin is a structural isomer of quercetin and exerts a wide range of pharmacological effects, including neuroprotective, cardioprotective, hepatoprotective, renoprotective, antidiabetic, anticancer, and anti-ulcer effects (Thakur et al.2020) mediated by its antioxidant and anti-inflammatory activity. We previously reported the beneficial effects of morin in cachexia-induced muscle atrophy in mice (Yoshimura et al.2018). Moreover, morin can regulate insulin/IGF-1 signaling to counteract various pathologies (Paoli et al.2013; Abuohashish et al.2021) indicating that morin could modulate muscle mass and function. We recently found the effect of morin in C2C12 cells treated with dexamethasone (Ulla et al.2021). However, the in vivo effect of morin on glucocorticoid-induced muscle atrophy is unknown. Therefore, we examined the effect of morin administration on dexamethasone-induced skeletal muscle atrophy in an in vivo model using C57BL/6J mice. Morin, the possible therapeutic agent in this study, was obtained from Sigma-Aldrich and used as a supplement to test its effectiveness against muscle atrophy in C57BL/6J mice.

Materials and methods

Reagents and chemicals

Morin, dexamethasone, and dimethyl sulfoxide (DMSO) were purchased from Sigma-Aldrich (St. Louis, MO, USA), thiobarbituric acid and tetramethoxypropane were obtained from Tokyo Chemical Industry (Tokyo, Japan), potassium iodide and chloramine-T were procured from Nacalai Tesque (Kyoto, Japan). All the other chemicals and reagents used were analytical grade.

Animals and experimental designs

A total of 21 female C57BL/6J mice (age, 12-13 weeks; weight, 19-22 g) were purchased from Japan SLC (Shizuoka, Japan) and housed in individual cages in a temperature-maintained room with 12-h dark/light cycles. Female mice were used as they are more responsive to dexamethasone (Baehr et al.2011) and also male mice are resistant to dexamethasone-induced skeletal muscle atrophy possibly because of the anabolic effect of testosterone (Zhao et al.2008). All mice had free access to standard laboratory food and water throughout the experimental period. Following acclimatization for 1 week before experiments, mice were randomly divided into three experimental groups with seven mice per group: control (group I), dexamethasone (group II), and dexamethasone + morin (group III). Treatment with morin was initiated 7 days before the initiation of dexamethasone treatment, which was for 10 days. Morin was dissolved in 5% DMSO in saline and prepared as a fresh solution every day. The mice in group I were orally administered 5% DMSO in saline (vehicle for morin) in equal volume used for morin and intraperitoneally injected with saline (vehicle for dexamethasone). The mice in group II were orally administered 5% DMSO in saline and intraperitoneally injected with dexamethasone (10 mg/kg body weight) once a day for 10 days. The mice in group III were administered morin (20 mg/kg body weight) by oral gavage every day throughout the experimental period of 17 days, with the coadministration of dexamethasone injection for the last 10 days of the experimental period. The body weight was measured every other day throughout the experiments in all mice.

This study was conducted with approval from the Committee on Animal Experiments of Tokushima University (permit no: T30-126) and performed following the guidelines for the care and handling of laboratory animals approved by Tokushima University.

Sacrifice of animals and sample collection

After the last dexamethasone and/morin treatments, the mice were starved for 6 h before sacrifice and tissue harvesting. Blood was collected in ethylenediaminetetraacetic acid-coated tubes by cardiac puncture and centrifuged at 5870 g for 15 min at 4°C to collect the plasma. The separated plasma was stored at −80°C for further investigation. Gastrocnemius (GA), tibialis anterior (TA) and soleus muscle were harvested, weighed, and stored at −80°C until further analysis.

Western blotting

The GA muscle tissue samples were homogenized in 10 volumes of lysis buffer containing 50 m m Tris HCl (pH 7.5), 150 m m NaCl, 5 m m ethylenediaminetetraacetic acid, 10 m m NaF, 2 m m Na₃VO₄, 1% Triton X-100, protease inhibitor cocktail (Roche Diagnostics, Rotkreuz, Switzerland), and 10 µm MG-132, followed by centrifugation at 12 000 g for 15 min at 4°C. The supernatants were collected, and protein concentrations were measured using Pierce^TM BCA protein assay kit (Thermo Fisher Scientific, Waltham, MA). A total of 20 µg of protein was loaded in each lane of 8% sodium dodecyl sulfate-polyacrylamide gels, electrophoresed at 300 V, and transferred to polyvinylidene difluoride membranes. The membranes were blocked for 1 h using 4% block ACE^TM (DS Pharma Biomedical, Osaka, Japan) dissolved in milli-Q water for 1 h. The membranes were incubated with primary antibodies overnight at 4°C and with secondary antibodies for 1 h at room temperature. A C-DiGit scanner (LI-COR Biosciences, Lincoln, NE, USA) was used for densitometric analysis of the blots. In the study, antibodies against fast-type myosin heavy chain (MyHC) (1:10 000) (Sigma-Aldrich), alpha-tubulin (1:2000) (Sigma-Aldrich), slow-type MyHC (1: 2000) (Novus Biologicals, Littleton, USA), total FoxO3a (1: 1000) (Cell Signaling Technology, Danvers, MA, USA), rabbit IgG (1: 5000) (Cell Signaling Technology), and phosphorylated FoxO3a (1: 1000) (Invitrogen, Waltham, MA) were used.

Western blotting to investigate the protein expression of slow-type MyHC was also performed using ProteinSimple^™ WES, a fully automated western blotting system, following the manufacturer's protocol. Briefly, 0.5 µg protein was mixed with Simple Western sample buffer and 5X Fluorescent Master Mix and denatured at 95^οC for 5 min. Protein samples, primary and secondary antibodies, and chemiluminescent substrate were dispensed in the designated wells of the Simple WES microplate according to the manufacturer's protocol. Finally, the WES microplate was inserted along with the suitable capillary cartridge to perform western blotting. After ∼3 h, software-generated results were analyzed to determine protein expression levels.

Reverse transcription-quantitative polymerase chain reaction

Total RNA was isolated from GA muscle homogenates using ISOGEN^TM (Nippon Gene, Tokyo, Japan), and RNA concentrations were measured using a Nanodrop 1000 spectrophotometer (Thermo Fisher Scientific). Next, 1 µg RNA from each sample was reverse transcribed to cDNA, and real-time polymerase chain reaction (PCR) was performed using the SYBR Green dye with the StepOnePlus real-time PCR system (Applied Biosystems, CA, USA). The internal standard was 18S ribosomal RNA. PCR oligonucleotide primer sequences used in the study are listed in Table 1.

Table 1.

PCR oligonucleotide primer sequences

Target gene		Sequence	Length(bp)
MAFbx1/atrogin-1	S	GGCGGACGGCTGGAA	101
	AS	CAGATTCTCCTTACTGTATACCTCCTTGT
MuRF-1	S	TGTCTGGAGGTCGTTTCCG	183
	AS	CTCGTCTTCGTGTTCCTTGC
Cbl-b	S	GAGCCTCGCAGGACTATGAC	222
	AS	CTGGCCACTTCCACGTTATT
Klf15	S	CCAGGCTGCAGCAAGATGTACAC	125
	AS	TGCCTTGACAACTCATCTGAGCGG
Nrf2	S	AGGACATGGAGCAAGTTTGG	482
	AS	TCTGTCAGTGTGGCTTCTGG
SOD1	S	ACCAGTGCAGGACCTCATTTTAA	78
	AS	TCTCCAACATGCCTCTCTTCATC
18sr	S	CATTCGAACGTCTGCCCTA	119
	AS	CCTGCTGCCTTCCTTGGA

18Sr, 18S ribosomal RNA; Cbl-b, casitas B-lineage lymphoma proto-oncogene-b; Klf15, Krüppel-like factor 15; MuRF-1, muscle ring finger protein-1; MAFbx1, muscle atrophy F-box protein-1; Nrf2, nuclear factor erythroid 2-related factor 2; SOD1, superoxide dismutase.

Analysis of oxidative stress markers

GA muscle tissue was homogenized in lysis buffer as described in western blotting section, followed by centrifugation at 13200 g for 15 min at 4^οC. The supernatants were collected to determine oxidative stress markers.

Evaluation of lipid peroxidation

The level of malondialdehyde (MDA), an oxidative stress marker, in plasma and GA muscle lysates was measured using the thiobarbituric acid reactive substance colorimetric assay, following the previously described method (Ohkawa et al.1979; Ulla et al.2017).

Assessment of advanced protein oxidation products

The levels of advanced protein oxidation products (APOP) in plasma and tissue samples were measured using a colorimetric method described by Alam et al. (Alam et al.2015). The supernatants from GA muscle homogenates were prepared as described above, and APOP concentration was expressed as nmol, mL⁻¹ or per mg chloramine-T equivalent using a chloramine-T standard curve that showed a linear regression between concentration and absorbance (at 340 nm) within a range from 0 to 100 nmol/mL.

Statistical analysis

Results were presented as means ± standard error of the mean (SEM). One-way analysis of variance followed by Tukey's post hoc test was used to compare data among the groups. Two-way repeated measure analysis of variance followed by Tukey's post hoc test was used to compare body weight changes of mice over the experimental period. All analyses were performed using GraphPad Prism version 9.3.1 (GraphPad Software, San Diego, CA, USA). Statistical significance was considered at a P-value of <.05.

Results

Effect of morin treatment on body weight

First, we measured the body weight of mice throughout the experimental period. The body weight in dexamethasone-treated mice started to decrease after 4 days of dexamethasone injection and it showed a significant difference compared with that of control mice after Day 6 and later (Figure 1a). Morin treatment partially restored body weight although the value did not reach that in the control group (Figure 1a).

Figure 1.

Morin prevents dexamethasone-induced loss of body and muscle weight in mice. (a) Mice were treated with dexamethasone (Dex; 10 mg/kg body weight) and/or morin (20 mg/kg body weight) for indicated durations, and body weights were measured at indicated time points. ^*P < .05, ^**P < .01. ^*Indicates significance difference between control and Dex treated group. Results are presented as means ± standard error of the mean (SEM), (n = 7/group). Total weight (b and d) and normalized weight (c and e) of GA and TA muscles. Results are presented as means ± SEM, n = 7/group. ^*P < .05, ^**P < 0.01, ^***P < .001. BW, body weight; Dex, dexamethasone; Dex + M, dexamethasone with morin; GA, gastrocnemius; TA, tibialis anterior; Normalized weight = total muscle weight/body weight.

Effect of morin treatment on muscle weight

The measurement of muscle weight of all mice at the end of the study revealed that both the total and normalized weights (muscle weight/body weight) of the GA and TA muscles were significantly lower in the dexamethasone-treated mice compared to the vehicle-treated control mice (Figures 1b-e), whereas the weight of the soleus muscle was not affected by dexamethasone treatment (data not shown). Interestingly, morin treatment significantly attenuated the loss of GA and TA muscle induced by dexamethasone. Both the total and normalized weights of the GA and TA muscle were increased in mice treated with dexamethasone and morin than in those treated with dexamethasone alone (Figures 1b-e). These results suggested that morin might be an effective phytochemical agent to prevent muscle atrophy.

Effect of morin treatment on muscle protein degradation

Glucocorticoids majorly affect fast-type MyHC by increasing MuRF-1 expression while exerting less impact on slow-type MyHC (Schakman et al.2013). We investigated the protein expression levels of fast- and slow-type MyHC in the GA muscle following dexamethasone and morin treatment. As shown in Figure 2a, treatment with dexamethasone significantly reduced the fast-type MyHC protein levels compared with the vehicle treated mice (Figure 2a). Notably, morin treatment effectively attenuated the dexamethasone-mediated degradation of fast-type MyHC. In contrast, no comparable changes in slow-type MyHC protein levels were noted among the three experimental groups (Figure 2b). These results indicated that dexamethasone reduced fast-type but not slow-type MyHC protein and that morin was effective in attenuating dexamethasone-induced breakdown of MyHC.

Figure 2.

Morin prevents dexamethasone-induced muscle protein degradation in mice. Fast- (a) and slow-type (b) myosin heavy chain (MyHC) protein levels in the GA muscle. Results are presented as means ± SEM, n = 3-4/group. ^*P < .05, ^**P < .01. Dex, dexamethasone; Dex + M, dexamethasone with morin; PC, positive control with sedentary C2C12 myotubes; MMSTD, molecular mass standard; GA, gastrocnemius.

Effect of morin administration on mRNA expression of E3 ubiquitin ligases and upstream regulators

We analyzed the mRNA expression levels of muscle atrophy-associated ubiquitin ligases, namely, atrogin-1 and MuRF-1, in the GA muscle using reverse transcription-quantitative PCR. The administration of dexamethasone significantly increased the expression of atrogin-1 and MuRF-1 compared with the vehicle treatment in mice (Figures 3a and b). Interestingly, morin treatment effectively suppressed dexamethasone-induced upregulation of the mRNA levels of these two ubiquitin ligases. Atrogin-1 expression in dexamethasone with morin treated group was found significantly higher than the control group.

Figure 3.

Morin suppresses dexamethasone-induced expression of ubiquitin ligases and upstream mediators associated with muscle atrophy. The mRNA expression levels of (a) atrogin-1, (b) MuRF-1 and (d) Klf15, n = 7. (c) Protein levels of total and phosphorylated FoxO3a n = 4. PC, positive control with Dex-treated C2C12 myotubes. All results are expressed as means ± SEM. ^*P < .05, ^**P < .01, ^***P < .001. Dex, dexamethasone; Dex + M, dexamethasone with morin. Klf15, Krüppel-like factor 15; MuRF-1, muscle ring finger protein-1; MMSTD, molecular mass standard.

We next examined the mRNA expression levels of Klf15 and the expression levels of total and phosphorylated FoxO3a, which are upstream of atrogin-1 and MuRF-1. FoxO3a, a transcription factor and downstream of Klf15, regulates the expression of atrogin-1 and MuRF-1, leading to muscle atrophy (Sandri et al.2004). As shown in Figure 3c, compared with the vehicle-treated control mice, dexamethasone treatment significantly increased the expression of total FoxO3a, whereas the expression of phosphorylated FoxO3a was reduced. The treatment with morin reversed the dexamethasone-mediated increase in the expression of FoxO3a and increased the expression of phosphorylated FoxO3a (Figure 3c).

Furthermore, the dexamethasone-treated mice exhibited significantly elevated mRNA levels of Klf15 compared to the vehicle-treated control mice (Figure 3d). Intriguingly, morin significantly reduced the dexamethasone-mediated increase in Klf15. These results indicated that morin could regulate the expression of atrogin-1 and MuRF-1 by modulating the expression of Klf15 and FoxO3a.

Effect of morin treatment on Cbl-b expression and oxidative stress markers

The dexamethasone treatment significantly increased the mRNA expression of Cbl-b compared with the vehicle treatment in mice (Figure 4a). Similarly, dexamethasone treatment significantly increased the accumulation of MDA, a lipid peroxidation product, as well as APOP in plasma and tissue samples compared with the vehicle treatment in control mice (Figures 4b and c). The treatment with morin effectively suppressed the increased mRNA expression of Cbl-b and the elevated levels of MDA and APOP induced by dexamethasone in both the plasma and tissue samples. The mRNA expression of nuclear factor erythroid 2-related factor 2 (Nrf2), an oxidative stress response gene, was upregulated in response to the dexamethasone treatment (Figure 4d) compared with the vehicle treatment and was followed by an increase in the superoxide-neutralizing enzyme superoxide dismutase (SOD1) (Figure 4e). The treatment with morin showed decreased expression of Nrf2 and SOD1 compared to the dexamethasone treatment (Figures 4d and e). The increased expression of MDA, APOP, and Nrf2 indicated that the dexamethasone-induced oxidative stress in the skeletal muscle, and it was suppressed by morin via its reactive oxygen species-scavenging effects.

Figure 4.

Morin attenuates dexamethasone-induced Cbl-b expression and oxidative stress in mice. (a) The mRNA expression levels of the ubiquitin ligase Cbl-b. (b and c) Levels of malondialdehyde (MDA) and advanced protein oxidation products (APOP) in plasma and tissue samples of mice respectively. Data are expressed as means ± SEM, n = 6. (d and e) The mRNA expression levels of Nrf2 and SOD1. Data are expressed as means ± SEM, n = 7. ^*P < .05, ^**P < .01, ^***P < .001. Dex, dexamethasone; Dex + M, dexamethasone with morin; Cbl-b, casitas B-lineage lymphoma proto-oncogene-b; Nrf2, nuclear factor erythroid 2-related factor 2; SOD1, superoxide dismutase.

Discussion

This study investigated the potential beneficial effects of morin, a natural antioxidant polyphenol, against dexamethasone-induced skeletal muscle atrophy in mice. Supplementation with morin for 17 days significantly attenuated dexamethasone-induced loss in GA and TA muscles. Moreover, morin effectively suppressed the expression of the ubiquitin ligases atrogin-1, MuRF-1, and Cbl-b, which are involved in muscle atrophy, consistent with our previous in vitro study demonstrating the effect of morin against dexamethasone-induced muscle atrophy in C2C12 myotubes (Ulla et al.2021).

Muscle atrophy is defined as decreased skeletal muscle mass and function due to reduced protein synthesis and increased protein degradation. Aging, muscle inactivity (e.g. immobilization, disuse, denervation, bed rest, space flight), cachexia, and glucocorticoid treatment are among the major causes of muscle atrophy. Glucocorticoids favor atrophy by activating the UPS, where the target protein is ubiquitinated by ubiquitin ligases atrogin-1 and MuRF-1. Dexamethasone activates the UPS by upregulating Klf15 and subsequently activating the FoxOs family of proteins (Shimizu et al.2011). FoxO3a, a transcriptional regulator of atrogin-1 and MuRF-1, is inactivated by phosphorylation that prevents its translocation to the nucleus, whereas the dephosphorylation of FoxO3a activates the UPS (Sandri et al.2004). In this study, morin administration prevented the dexamethasone-induced upregulation in the mRNA levels of atrogin-1, MuRF-1, and Klf15. Moreover, morin reversed the dexamethasone-mediated induction of FoxO3a by leading to an increase in phosphorylation of FoxO3a, thereby inhibiting its nuclear translocation and downstream induction of ubiquitin ligases. Based on these findings, we suggest that morin can prevent muscle atrophy in vivo by inhibiting the expression of ubiquitin ligases through the modulation of Klf15 and FoxO3a.

Contrary to these in vivo results, morin had failed to suppress Klf15 expression in our previous in vitro study using C2C12 myotubes, which may be due to insufficient dose or suboptimal dosage regimen necessary for effective Klf15 suppression. Furthermore, the metabolites of morin produced after its ingestion by mice might account for the effective suppression of Klf15 expression. Future studies are warranted to investigate the mechanism underlying the differences in morin-induced Klf15 suppression between in vivo and in vitro settings.

Oxidative stress, an important mechanism underlying muscle atrophy (Ábrigo et al.2018; Oh et al.2021), can cause mitochondrial dysfunction, protein degradation, increased ubiquitin-proteasome activity, and reduced protein synthesis via the upregulation of atrogin-1 and MuRF-1 and the downregulation of the phosphoinositide 3-kinase/Akt pathway (Gomes-Marcondes and Tisdale 2002). In this study, we measured the levels of MDA and APOP to assess oxidative stress. MDA is produced by lipid peroxidation following the reaction of reactive oxygen species with polyunsaturated fatty acids. Both MDA and APOP levels were significantly increased in the plasma and muscle tissue of mice treated with dexamethasone compared to the control group, which are in agreement with previous studies (Kim et al.2015), whereas morin treatment led to a significant reduction in MDA and APOP levels. The hydroxyl group at 2′ and 4′ positions in the β-ring of morin plays a notable role in the reduction of lipid peroxidation (Wolfe and Liu 2008). Similarly, the 4′-hydroxyl group of morin is credited for its free radical scavenging effect (Farkas et al.2004). In the present in vivo study, we also found that the mRNA expression of Nrf2, which is responsible in mounting response to oxidative damage in tissues, was increased in parallel with the increased mRNA expression of SOD1 in dexamethasone-treated mice; these findings might reflect an early protective response against oxidative stress.

Consequent to oxidative stress, we also found that the mRNA expression of the stress-sensitive ubiquitin ligase Cbl-b was significantly upregulated by the dexamethasone treatment. Cbl-b impairs the IGF-1 signaling pathway by degrading insulin receptor substrate-1, which leads to the dephosphorylation of FoxO3a and increases in atrogin-1 and MuRF-1. The IGF-1 signaling is involved in various pathways responsible for muscle growth, differentiation, and regeneration (Scicchitano et al.2009). In this study, morin effectively reduced the dexamethasone-induced Cbl-b expression and prevented the upregulation in atrogin-1 and MuRF-1 expression. Previous studies reported that morin could enhance insulin/IGF-1 signaling and induce the expression of genes involved in insulin signaling such as insulin receptor substrate-1 and phosphoinositide 3-kinase in rat L6 myotubes (Abuohashish et al.2021; Issac et al.2021). Thus, we suggest that the beneficial effects of morin may also include the prevention of oxidative stress through the suppression of Cbl-b and the downstream Cbl-b-mediated ubiquitin ligases.

One of the hallmarks of dexamethasone-induced atrophy is the fast-to-slow muscle fiber type transition. The degradation of fast-type MyHC is more pronounced than that of slow-type MyHC in response to dexamethasone, which might be due to the higher expression of glucocorticoid receptors in the fast-type muscle. Moreover, MuRF-1 causes the breakdown of MyHC during dexamethasone-induced atrophy (Peris-Moreno et al.2020). In this study, dexamethasone-induced a significant decrease in fast-type MyHC, whereas no noticeable changes were observed in slow-type MyHC. Morin efficiently attenuated the dexamethasone-induced fast-type MyHC, which may be due to the suppression of MuRF-1 by morin.

The bioavailability of morin after its oral ingestion has rarely been studied. After the oral administration, the dietary morin goes through the stomach and reaches the small intestine as free or glycosylated, methylated, or sulphated derivatives (Heim et al.2002; Xie et al.2006). Pancreatic and intestinal enzymes convert these derivatives into their respective aglycone during the digestive action, which are essential for the absorption from the intestine. Furthermore, the unabsorbed fraction of morin from the small intestine moves to the large intestine and gets metabolized to the preferred morin aglycon by gut microbes (Rajput et al.2021). A study conducted by Li et al. found that the plasma concentration of morin was 3.38 µg/mL (11.2 µm) after 30 min of oral administration at 200 mg/kg in Wistar rats (Li et al.2019). Another study reported that the Cmax was 84.9 µm following gastric gavage of morin at 50 mg/kg in rats (Hou et al.2003). These results suggest the limited bioavailability of morin after single dose oral administration. In the present study, we administered morin (aglycon) which may have less absorption at 20 mg/kg, suggesting that the plasma concentration and the amount of morin that reached muscle might be less than the estimated value. However, Li et al. further showed that orally administered morin (Sigma-Aldrich) showed a longer plasma half-life than that were intravenously injected (Li et al.2019). In our present study, orally and daily administered morin might retain in plasma and be metabolized to metabolites that functioned to restore muscle abnormality. No previous reports, at least to our knowledge, described the precise amount of morin in plasma or muscle of mice after its oral administration. Hence, it is our future experimental plan to investigate the concentration of morin in plasma or muscle after oral administration.

Conclusion

In conclusion, we demonstrated that morin prevented dexamethasone-induced muscle atrophy in C57BL/6J mice. Morin downregulated the expression of genes involved in muscle atrophy and reduced the dexamethasone-induced protein degradation of MyHC. Oxidative stress and Cbl-b expression were effectively attenuated by morin in atrophied muscles. Therefore, we suggest that morin is a natural supplement with beneficial effects in muscle atrophy.

Acknowledgments

The authors would like to thank Enago (www.enago.jp) for the English language review and editing.

Data availability

The data underlying this article will be shared on reasonable request to the corresponding author.

Author contribution

A.U.: Conceptualization, methodology, investigation, data curation, formal analysis, writing—original draft. K.O.: Methodology, writing original—draft, formal analysis. M.M.R.: Investigation, data curation, formal analysis. R.N.: Methodology, visualization, validation, writing-review and editing. T.U.: Investigation, formal analysis. I.M.: Investigation, writing original—draft, formal analysis. K.M.: Methodology, visualization, formal analysis. T.F.: Conceptualization, methodology, visualization, writing—review and editing. H.O.K.: Methodology, visualization, writing—review and editing. I.S.: Visualization, writing—review and editing. K.H.: Methodology, visualization, writing—review and editing. T.N.: Conceptualization, methodology, visualization, funding acquisition, writing—review and editing, supervision.

Funding

This work was supported by Cabinet Office, Government of Japan, Cross-ministerial Moonshot Agriculture, Forestry and Fisheries Research and Development Program, “Technologies for Smart Bio-industry and Agriculture” (funding agency: Bio-oriented Technology Research Advancement Institution), Grant Number JPJ009237.

Disclosure statement

No potential conflict of interest was reported by the authors.