Lycium Radicis Cortex and Its Kukoamine Constituents Attenuate Sarcopenia by Modulating Anabolic and Catabolic Pathways

Abstract

Lycium Radicis Cortex (LRC), derived from the root bark of Lycium chinense Mill., has traditionally been used in East Asian medicine to mitigate heat in the blood and consumptive fever. This study investigates LRC’s effects on skeletal muscle in aged mice subjected to forced exercise and examines the protective properties of its primary constituents, kukoamines A (KA) and B (KB), against dexamethasone (DEX)-induced muscle atrophy. Sixteen-month-old male C57BL/6 mice underwent regular swimming and received oral LRC supplementation for 8 weeks. The effects of KA and KB on muscle atrophy were further explored using C2C12 myotubes treated with DEX. LRC administration significantly enhanced muscle mass, strength, and endurance, while reducing plasma lactate and creatinine levels compared to the control group. LRC also upregulated mRNA expression of MyoD, myogenin, MHC, Akt, and mTOR, and downregulated myostatin, FoxO3a, MuRF1, and atrogin-1 in gastrocnemius and soleus muscles. Furthermore, KA and KB alleviated DEX-induced muscle atrophy in C2C12 myotubes by reducing proteolysis and ROS production, enhancing SOD activity, and improving mitochondrial function. Taken together, LRC may be a useful supplement in exercise-based muscle strengthening and amelioration of muscle disorders, and KA and KB have shown potential as preventive and therapeutic agents for muscle atrophy, indirectly suggesting that the efficacy of LRC is attributed to KA and KB.

INTRODUCTION

Sarcopenia, the age-associated loss of skeletal muscle mass and strength, is a growing public health concern due to its contribution to frailty, decreased mobility, and increased risk of falls in the elderly population (Larsson et al., 2019). This progressive condition is driven by multifactorial mechanisms, including chronic inflammation, oxidative stress, mitochondrial dysfunction, and impaired protein synthesis (Bilski et al., 2022). While physical activity remains a cornerstone of prevention and treatment of sarcopenia, aged individuals often respond less effectively to exercise interventions due to reduced regenerative capacity and elevated catabolic signaling (Chinvattanachot et al., 2024; Moretti et al., 2025). Therefore, the development of safe, effective therapeutic agents that can complement exercise and attenuate muscle atrophy in the elderly is of considerable interest.

Natural products have emerged as promising candidates for modulating muscle metabolism and improving muscle health (Jia et al., 2022; Kim et al., 2018; Lee et al., 2021a, 2022a). Among them, Lycium genus, traditionally used in East Asian medicine, has gained attention for its antioxidant, neuroprotective, and anti-aging properties (Jiang et al., 2024). Lycii radicis cortex (LRC), the root bark of Lycium chinense Mill., has been widely utilized as a traditional Korean herbal medicine for centuries, and is known to effectively benefit the kidney, lungs, and liver, and has been traditionally used for blood cooling, fever reduction, and lung fire removal (Wang and Luo, 2006). In addition, numerous in vitro and in vivo studies have demonstrated various pharmacological effects of LRC extracts, including reducing serum glucose and lipid levels, improving insulin resistance, protecting pancreatic β-cells, improving bone formation, anti-depressant, and anti-tumor (Cho et al., 2011; Jeong et al., 2012; Kim et al., 2013; Park et al., 2014; Wang and Ye, 2016; Ye et al., 2008).

LRC consists of phenolic acids, coumarins, alkaloids, flavonoids, and various other compounds (Qian et al., 2017). Among the various constituents, kukoamines A (KA) and B (KB), which have similar structures, are recognized as the primary ingredients of LRC (Kim et al., 2022). Kukoamines A and B, phenolic amides predominantly found in LRC, have demonstrated biological activities such as anti-inflammatory, antioxidant, anti-amyloid β (Aβ) aggregation, and anti-diabetic properties in various models (Jiang et al., 2020; Lee et al., 2022b; Li et al., 2018, 2019). Although the biological roles of LRC, KA, and KB in humans have been elucidated, their potential anti-sarcopenia effects remain largely unexplored. Herein, we investigated the efficacy of LRC extract in improving sarcopenia in aged mice subjected to a forced exercise regimen and also identified KA and KB as key active markers and evaluated their contributions to the observed effects.

MATERIALS AND METHODS

Sample preparation

The Lycii Radicis Cortex (LRC) was harvested from Henan Province, China, and purchased through Songlim Pharm. Co., Ltd. (Seoul, Korea). Prof. CY Kim, the corresponding author, authenticated the LRC specimens and stored them at ambient temperature until employed for the research. A voucher specimen (No. HYUP-LR-002) was archived at the Pharmacognosy Laboratory, College of Pharmacy, Hanyang University. Kukoamines A and B (KA and KB), the main compounds of LRC, were purchased from ChemFace (Hubei, China).

HPLC analysis of LRC, KA, and KB

The purity and substance determination of LRC, KA, and KB were confirmed by HPLC analysis according to previous literature (Son et al., 2024). In brief, an Agilent 1260 system (Agilent, Santa Clara, CA, USA) equipped with a binary pump, autosampler, column oven, and diode array detector was used to perform the HPLC analysis. A Zorbax C18 SB-AQ column was used to achieve separation. The mobile phase consisted of trifluoroacetic acid (0.1%) in acetonitrile (A) and trifluoroacetic acid (0.1%) in water (B), with a gradient elution as follows: 0-5 min, 10% A; 5-25 min, 15% A; 25-35 min, 100% A; 35-40 min, 100% A. The flow rate was set at 1.0 mL/min, and detection was performed at 280 nm.

Mouse maintenance

Mouse experimentation was approved by the Animal Ethics and Welfare Committee of Chaon (Approval No. CE22492) and was conducted in strict accordance with ethical guidelines and regulations under the NIH guide for the care and use of Laboratory animals. A total of 30 male C57BL/6 mice, aged 16 months, were acquired from Orient Bio (Seongnam, Korea) and kept in an animal facility specifically designated for that purpose. The facility ran on a reversed 12:12 h light/dark cycle, with a temperature range of 23 ± 3 °C, relative humidity between 50 ± 20%, and illumination intensities between 150-200 lux. Mice were given unlimited access to water and a standard chow diet during the trial.

Mouse experimental design

The mice were randomly divided into five experimental groups (n=6 per group) after a one-week acclimatization period. These groups included: (1) control group, aged with no exercise; (2) swim group, aged mice with swimming; (3) swim+LRC 50 group, aged mice with swimming plus LRC 50 mg/kg supplementation; (4) swim+LRC 100 group, aged mice with swimming plus LRC 100 mg/kg supplementation; (5) swim+LRC 200 group, aged mice with swimming plus LRC 200 mg/kg supplementation. The LRC (50, 100, and 200 mg/kg) was orally administered for 8 weeks at a volume of 10 ml/kg using gastric gavage. The forced swimming was performed in the chamber (L427 x W287 x H198 mm) 30 min after feeding and for 2 min, 3 times a week throughout the experimental period. Weekly monitoring of body weight was conducted, and upon completion of the experiment, whole blood and muscles (soleus and gastrocnemius) were collected following euthanasia. The soleus and gastrocnemius muscles were swiftly separated and weighed before being preserved in liquid nitrogen for subsequent analysis.

Measurements of physical performance

Following 8 weeks of LRC administration, grip strength was assessed using a BIO-GS3 meter (BIOSEB, Vitrolles, France). The procedure involved positioning the mouse on a grid with its body aligned to its height and all four paws placed on the grid (Ueno et al., 2020). Subsequently, the tail was gently pulled back, repeating this process five times per mouse, and calculating the average value. The treadmill test was conducted over 3 days. On the first day, mice underwent treadmill training at 10 m/min with a 10% incline for 10 min. On the second day, the mice were acclimated by starting at 10 m/min with a 10% incline, increasing the speed by 1 m/min every minute until reaching 25 m/min. Additionally, an electronic grid with a 1 mA current was used on the first and second days. On the third day, an endurance test was carried out. The test began at 10 m/min with a 10% incline, increasing by 1 m/min every minute until 25 m/min was reached. This speed was maintained for 15 min. Exhaustion was determined when a mouse could not maintain the treadmill speed for more than 10 s.

Plasma analysis

Blood was collected from the abdominal vena cava of the euthanized mice. The plasma was separated from the collected whole blood by centrifuging it at 3000 rpm for 10 min at room temperature. Creatinine levels were assessed with an automated analyzer (BECKMAN Coulter, Brea, CA, USA) and lactate levels were evaluated employing an ELISA kit.

Real-time polymerase chain reaction (RT-PCR)

Gastrocnemius and soleus muscles RNA were extracted and purified using RNeasy kits and QIAzol lysis reagent (Qiagen Inc, Germantown, MD, USA). Subsequently, cDNA synthesis was carried out utilizing total RNA (5 μg) with the High-Capacity Kit (Applied Biosystems, Waltham, MA, USA). Specific primers and the Thunderbird^TM qPCR mix (Toyobo, Osaka, Japan) were used for quantitative real-time PCR (qRT-PCR). The qRT-PCR reactions were conducted on a CFX 384 Touch™ System (Bio-Rad, Hercules, CA, USA). Quantification of the amplified products was performed utilizing the comparative cycle threshold (Ct) method, with normalization of each gene’s expression level to GAPDH. CFX Manager^TM software (Bio-Rad, Hercules, CA, USA) was used to evaluate the data. Table 1 provides detailed information regarding the primer sequences for the target genes.

Table 1.

Primer information

Samples	Species	Gene	Sequence (F)	Sequence (R)
Soleus and gastrocnemius muscle	Mouse	MyoD	5’-AGCGACACAGAACAGGGAAC-3’	5’-TGCTGCAGTCGATCTCTCAA-3’
	Mouse	Myogenin	5’-AGCAGGGGGCTTTTAAGTGG-3’	5’-TAAATTCCCTCGCTGGGCTG-3’
	Mouse	MHC	5’-CGACACAGCCTACAGAAGCA-3’	5’-TTTCCAGCTCCCCCGTGAT-3’
	Mouse	Akt	5’-ATGAACGACGTAGCCATTGTG-3’	5’-TTGTAGCCAATAAAGGTGCCAT-3’
	Mouse	mTOR	5’-TATCCGCTACTGTGTCTTGGC-3’	5’-GCTCGCGGATCTCAAAGACC-3’
	Mouse	Myostatin	5’-TACGACGTCCAGAGGGATGA-3’	5’-TTGCCATCCGCTTGCATTAG-3’
	Mouse	FoxO3a	5’-AAGGGAAGGAGCCGAGGTAG-3’	5’-GCTTGGGCTCTTGCTCTCTC-3’
	Mouse	MuRF1	5’-TGCCAAGCAGCTCATCAAGA-3’	5’-TCCCAAAGTCAATGGCCCTC-3’
	Mouse	Atrogin-1	5’-GCCCTCCACACTAGTTGACC-3’	5’-GACGGATTGACAGCCAGGAA-3
	Mouse	GAPDH	5’-CATGGCCTCCAAGGAAGA-3’	5’-GAGGGAGATGCTCAGTGTTGG-3’

Histological analysis

For histopathological evaluation, gastrocnemius tissue underwent preservation in 10% formalin (Biosesang, Seongnam, Korea), followed by embedding in paraffin utilizing an automated tissue processor (Thermo Fisher Scientific, Waltham, MA, USA). Each block was sectioned at 4 µm thickness and subjected to hematoxylin and eosin (H&E) staining. Imaging of the stained specimens was carried out utilizing an optical microscope. Estimation of the cross-sectional area of muscle fibers in each mouse involved calculating the average of 50 fibers, facilitated by CellSens image analysis software (Olympus, Tokyo, Japan).

C2C12 cell maintenance and treatment with dexamethasone, kukoamines A and B

C2C12 myoblast cells were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA), which were then grown in high-glucose (25 mM) Dulbecco’s Modified Eagle Medium (DMEM; Gibco, Waltham, MA, USA) containing 10% fetal bovine serum (FBS; Gibco, Waltham, MA, USA), 1% penicillin/streptomycin (P/S) under conditions of 5% CO₂ at 37°C. Upon reaching over 90% confluence, the growth medium was changed every two days to DMEM containing 2% horse serum (HS; Gibco, Waltham, MA, USA) to stimulate differentiation into myotubes for 6 days. Following complete differentiation, myotubes were exposed to 10 μM dexamethasone (DEX; Sigma-Aldrich, St. Louis, MO, USA), either alone or combined with KA and KB at concentrations of 50 and 100 μM, for 24 h. Control cells were incubated in DMEM supplemented with 2% HS and 0.1% DMSO (Sigma-Aldrich, St. Louis, MO, USA), serving as the vehicle for DEX.

Cell viability assay

Cell counting kit-8 (CCK-8; Dojindo, Kumamoto, Japan) was used to assess the viability of C2C12 myotubes. For 24 h, fully differentiated C2C12 myotubes were subjected to 10 μM DEX with or without KA or KB at 50 and 100 μM doses. After adding 20 μL of CCK-8 reagent, the plates were incubated in a CO₂ incubator for 4 h. After that, absorbance was measured with a plate reader at 450 nm (PerkinElmer, Waltham, MA, USA).

Immunofluorescence staining

After treatment with 10 μM DEX with or without KA or KB (50 and 100 μM) for 24h, the myotubes were fixed for 10 min using 4% paraformaldehyde (Biosesang, Seongnam, Korea). After washing 3 times with PBS, permeabilization was performed with 0.1% Triton X-100 for 20 min. The cells were then blocked with 3% BSA for 1 h and incubated overnight at 4°C with a myosin heavy chain (MHC) antibody (1:300, Santa Cruz, TX, USA). The next day, myotubes were treated with an Alexa Fluor 488 goat-mouse IgG conjugated secondary antibody (1:500, Invitrogen, Carlsbad, CA, USA) for 1 h at 37°C. Subsequently, Hoechst 33342 was used to counterstain the nuclei. The diameter, MHC-positive area, and fusion index of C2C12 myotubes were assessed following the literature (Kim et al., 2023) after the stained myotubes were viewed using a JuLI^TM stage (Nano Entek, Seoul, Korea).

Western blotting analysis

C2C12 myotubes were lysed using RIPA lysis buffer (Invitrogen, Carlsbad, CA, USA) supplemented with 1% protease/phosphatase inhibitor cocktails. The lysates were then quantified and separated on 8% or 12% SDS-PAGE. Following electrophoresis, proteins were transferred to PVDF membranes (Merck, Darmstadt, Germany). Subsequently, PVDF membranes were blocked at room temperature with 3% BSA for 2 h. Following blocking, the membranes were incubated overnight with various primary antibodies targeting MHC, FoxO3a, MuRF1, atrogin-1, Akt, and p-Akt (diluted at 1:1000, Santacruz, Dallas, TX, USA), as well as GAPDH (diluted at 1:5000, Santacruz, Dallas, TX, USA) at 4 °C. The subsequent day involved 3 washes with 0.2% PBST followed by treatment with corresponding HRP-linked anti-mouse or anti-rabbit secondary antibodies (diluted at 1:5000, Santacruz, Dallas, TX, USA) for 2 h at room temperature. The expression of the protein on each membrane was then visualized utilizing an ECL solution (Thermo Fisher Scientific, Waltham, MA, USA) and Chemidoc (Biorad, Hercules, CA, USA). The band densitometry for each protein was calculated as the expression ratio compared to GAPDH expression using ImageJ software.

Measurement of intracellular ROS

Intracellular ROS levels in C2C12 myotubes were assessed using the DCF-DA method. Fully differentiated C2C12 myotubes were exposed to 10 µM DEX, with or without 50 and 100 µM KA or KB, for 24 h. Following treatment, the myotubes were washed with PBS. Then, 10 µM CM-H2DCFDA (Invitrogen, Carlsbad, CA, USA) was added to each well, and the cells were incubated for 15 min in the dark state at room temperature. After incubation, the wells were washed with fresh medium. Fluorescent imaging was conducted using a JuLI^TM stage fluorescence microscope and Image J software was used to assess the fluorescence intensity.

Measurement of SOD activity

SOD levels were assessed utilizing a SOD assay kit (Dojindo, Kumamoto, Japan) according to the manufacturer’s guidelines. After 24 h co-treatment with DEX (10 μM) and either KA or KB (50 and 100 μM, respectively), myotubes were lysed by scraping to prepare the samples. Each sample (20 µL) was added to a 96-well plate, followed by the addition of 200 µL of WST and 20 µL of enzyme working solution. Following the incubation of the plate at 37°C for 20 min, SOD activity was determined at 450 nm using a plate reader.

Mitochondria staining

C2C12 myotubes were subjected to 10 µM DEX in the presence or absence of KA or KB (at concentrations of 50 and 100 µM) for 24 h. Following this, the myotubes were treated with 500 nM Mito Tracker Deep Red (Invitrogen, Carlsbad, CA, USA) and incubated at 37°C for 30 min. Following their incubation, the myotubes underwent a PBS wash and a 4% paraformaldehyde fixation. The myotubes were PBS-washed three times following fixation. The stained myotubes were captured and visualized using a JuLI^TM stage and Image J software was used to quantify red fluorescence intensity.

Determination of the ATP level

To quantify the total amount of ATP levels in C2C12 myotubes, an ATP colorimetric assay kit was utilized (Thermo Fisher Scientific, Waltham, MA, USA). The myotubes were exposed to KA or KB (50 and 100 µM, respectively) and/or DEX (10 µM) for 24 h. Subsequently, myotubes were washed with PBS and lysed using a sample buffer. The lysate was centrifuged at 13,000× g for 10 min to collect the supernatant, and then 10 µL was carefully transferred to a plate. After transportation, a 20 min room temperature incubation was conducted with the addition of 100 µL of ATP reaction mix solution. The absorbance was then measured at a wavelength of 560 nm following incubation.

Statistical analyses

All data were presented as mean ± standard deviation (SD) for at least three independent experiments. Statistical significance was assessed and determined by one-way analysis of variance (ANOVA) using GraphPad Prism 5.0 (GraphPad Software Inc., San Diego, CA, USA), followed by Tukey’s post-hoc test. Differences were considered statistically significant at p<0.05.

RESULTS

LRC supplementation increases muscle mass in aged mice without affecting overall body weight

Body weight did not vary among the groups, however, there were notable alterations in the weights of the gastrocnemius and soleus muscles (Fig. 1). Specifically, the swimming group (swim group) displayed higher weights for both the gastrocnemius and soleus muscles compared to the control group (Fig. 1C, 1D). The swim+LRC 100 and 200 mg/kg groups exhibited a significant increase in gastrocnemius and soleus muscle masses, respectively, in comparison to the swim group (Fig. 1C, 1D). However, no dose-dependent discrepancies were detected.

Fig. 1.

Influence of LRC on body and muscle weight in mice (n=6 per group). (A) Body weight changes of mice during 8 weeks. (B) Representative images of the soleus and gastrocnemius muscles. (C) Gastrocnemius muscle weight. (D) Soleus muscles weight. These data are presented as means ± SD. ^##p<0.01, ^###p<0.001 vs. Control; **p<0.01 vs. Swim.

LRC supplementation enhances muscle strength and exercise capacity while alleviating muscle fatigue

Swimming resulted in a significant elevation in grip strength, exhaustion running duration time, and distance when compared to the control group (Fig. 2A-2C). Grip strength increased significantly in the swim+LRC 100 and 200 mg/kg groups as compared to the swim group (Fig. 2A). During the exhaustion running experiments, significant enhancements were observed in the swim+LRC 50, 100, and 200 mg/kg groups compared to the swim group (Fig. 2B, 2C). Additionally, compared to the control group, the swim group had higher plasma lactate levels (Fig. 2D), which is a sign of post-exercise muscular fatigue. However, in comparison to the swim group, this rise was significantly reduced in the swim+LRC 100 and 200 mg/kg groups (Fig. 2D). Conversely, creatinine levels, reflecting muscle metabolism byproducts, notably decreased in the swim group relative to the control group, with a more pronounced decrease observed in the swim+LRC 50, 100, and 200 mg/kg groups compared to the swim group (Fig. 2E).

Fig. 2.

Changes in exercise capacity and plasma biochemistry (n=6 per group). (A) Grip strength. (B) Duration time of the treadmill. (C) Distance of the treadmill. (D) Plasma lactate level. (E) Plasma creatinine level. These data are presented as means ± SD. ^#p<0.05, ^###p<0.001 vs. Control; *p<0.05, **p<0.01, ***p<0.001 vs. Swim.

LRC regulates the expression of anti-sarcopenic-related factors in soleus and gastrocnemius

In both the soleus and gastrocnemius tissues, the mRNA levels associated with muscle differentiation (MyoD, myogenin, and MHC), synthesis (Akt and mTOR), and degradation (myostatin, FoxO3a, MuRF1, and atrogin-1) were assessed. As shown in Fig. 3, compared to the control group, the swim group promoted mRNA levels associated with muscle differentiation (MyoD, myogenin, and MHC) and synthesis (Akt and mTOR), while decreasing degradation (myostatin, FoxO3a, MuRF1, and atrogin-1). The swim+LRC groups showed considerably higher mRNA expression of MyoD, myogenin, MHC, Akt, and mTOR in the soleus and gastrocnemius when compared to the swim group, while the degradation markers myostatin, FoxO3a, MuRF1, and atrogin-1 showed significantly lower mRNA expression (Fig. 3).

Fig. 3.

The mRNA expression levels involved in muscle differentiation, synthesis, and degradation in the soleus and gastrocnemius muscles (n=6 per group). (A-C) Genes related to muscle differentiation mRNA (MyoD, Myogenin, and MyHC). (D, E) Genes related to muscle protein synthesis mRNA (Akt and mTOR). (F-I) Genes related to muscle degradation mRNA (Myostatin, FoxO3a, MuRF1, and Atrogin-1). These data are presented as means ± SD. ^#p<0.05, ^##p<0.01, ^###p<0.001 vs. Control; *p<0.05, **p<0.01, ***p<0.001 vs. Swim.

LRC improves muscle fiber cross-sectional area in gastrocnemius

H&E staining was performed to confirm histological changes in the gastrocnemius. As a result, swimming significantly increased the mean muscle fiber cross-sectional area of the gastrocnemius muscles when compared to the control group (Fig. 4). In addition, LRC supplementation enhanced swimming-induced gastrocnemius muscle hypertrophic changes dose-dependently (Fig. 4).

Fig. 4.

The histology of gastrocnemius muscle (n=6 per group). (A) Representative Hematoxylin & Eosin (H&E) staining (scale bar=25 μm). (B) Quantitative analysis of the cross-sectional area. These data presented as means ± SD. ^#p<0.05, *p<0.05 vs. Control; **p<0.01, ***p<0.001 vs. Swim.

Kukoamines A and B protect C2C12 myotubes from DEX-induced atrophy

The viability of the myotubes exposed to DEX exhibited a significant reduction when compared to the control group (Fig. 5A). However, co-treatment with 100 µM KA or KB and DEX resulted in a significant restoration of the DEX-induced decrease in viability of myotubes (Fig. 5A). Morphological changes were assessed by MHC immunostaining (Fig. 5B). DEX treatment induced a significant decrease in myotube diameter, MHC staining intensity, and fusion index in myotubes compared to the control group (Fig. 5B-5E). In contrast, KA- or KB-treated myotubes significantly reversed the DEX-induced reduction in myotube diameter, MHC staining intensity, and fusion index, respectively (Fig. 5B-5E). Furthermore, the MHC protein expression was remarkably lower in DEX-treated myotubes than in the control (Fig. 5F, 5G). However, this decrease in MHC protein expression was significantly restored by 50 or 100 µM of KA and KB, respectively (Fig. 5F, 5G).

Fig. 5.

Effects of KA and KB on cell viability, morphological changes, and MHC protein expression in DEX-treated C2C12 myotubes. (A) C2C12 myotubes’ viability. (B) Immunofluorescence staining with MHC (scale bar=250 μm). (C) Relative changes of MHC-stained area, (D) diameters, and (E) fusion index. (F) MHC expression in C2C12 myotubes treated with DEX and KA and (G) MHC expression in C2C12 myotubes treated with DEX and KB. GAPDH was used as the loading control. These data are presented as means ± SD. ^###p<0.001 vs. Control; *p<0.05, **p<0.01, ***p<0.001 vs. Swim.

Kukoamines A and B ameliorate atrophic signaling by DEX in the C2C12 myotubes

The effects of KA and KB on the expression of muscle protein degradation-related factors induced by DEX in C2C12 myotubes using western blot analysis were analyzed. Our results showed that DEX markedly reduced Akt phosphorylation and up-regulated FoxO3a, MuRF1, and atrogin-1 (Fig. 6). KA and KB treatment restored the changes in DEX-induced FoxO3a, atrogin-1, and MuRF1 expression, and notably, despite DEX treatment, 50 or 100 µM KA or KB-treated myotubes increased Akt phosphorylation compared to the control group (Fig. 6A, 6B, 6F, 6G).

Fig. 6.

Effects of KA and KB on the expression of markers associated with muscle atrophy. (A) and (F) The expression of p-Akt, Akt, FoxO3a, MuRF1, and Atrogin-1 were measured using western blot analysis. GAPDH was used as the loading control. (B) and (G) Quantitative analysis of p-Akt/Akt, (C) and (H) FoxO3a, (D) and (I) MuRF1, and (E) and (J) Atrogin-1. These data are presented as means ± SD of three independent experiments. ^#p<0.05, ^##p<0.01, ^###p<0.001 vs. Control; *p<0.05, **p<0.01 vs. Swim.

Kukoamines A and B improve antioxidant activity in DEX-treated C2C12 myotubes

DEX treatment significantly increased the positive intensity of DCFH-DA compared to the control group (Fig. 7A, 7B) accompanied by a decrease in SOD activity (Fig. 7C). The myotubes co-treated with KA or KB with DEX inhibited ROS generation (Fig. 8A, 8B) and increased SOD activity compared to DEX-treated myotubes (Fig. 7C). Despite the DEX treatment, both the KA or KB treatments suppressed ROS formation to levels similar to or lower than the control group and restored SOD activity similar to the control group (Fig. 7).

Fig. 7.

Effects of KA and KB on ROS production and SOD activity in DEX-treated C2C12 myotubes. (A) The production of cellular ROS analysis using a 2,7-dichlorodihydrofluorescein diacetate (DCFH-DA) dye (scale bar=250 μm). (B) The graph shows the green fluorescence intensity of DCF-positive C2C12 myotubes. (C) The antioxidant enzyme (SOD) activity in C2C12 myotubes. These data are presented as means ± SD of three independent experiments. ^##p<0.01 vs. Control; **p<0.01, ***p<0.001 vs. Swim.

Fig. 8.

Effects of KA and KB on mitochondrial content and ATP levels in DEX-treated C2C12 myotubes. (A) Under a fluorescence microscope, the contents of the mitochondria were measured using MitoTracker Deep Red dye (scale bar=250 μm). (B) The graph is representative of the quantification of the MitoTracker Deep Red stained area. (C) ATP production levels in the C2C12 myotubes. These data are presented as means ± SD of three independent experiments. ^##p<0.01, *p<0.05 vs. Control; **p<0.01, ***p<0.001 vs. Swim.

Kukoamines A and B ameliorate DEX-induced reduction in mitochondrial content and ATP levels in C2C12 myotubes

When compared to the control group, treatment with DEX dramatically reduced the mitochondrial content and ATP levels in C2C12 myotubes (Fig. 8). However, treatment with KA or KB in combination with DEX significantly reversed the decrease in mitochondrial content in comparison to myotubes treated with DEX alone, respectively (Fig. 8A, 8B). In addition, 100 µM of KA and KB also significantly restored the ATP levels reduced by DEX (Fig. 8).

DISCUSSION

Regular exercise is a well-established strategy for preventing and managing sarcopenia, as it stimulates muscle protein synthesis and improves muscle strength and function (Landi et al., 2014). Swimming is a regular aerobic exercise that, as a non-drug intervention, is physiologically and structurally useful in preventing diseases and injuries associated with skeletal muscle disorders in old age (Zargani et al., 2023). In this study, we implemented a moderate swimming protocol (2 min per session, 3 times per week) to mimic a low-intensity, consistent exercise routine suitable for aging or frail populations. As shown in Fig. 1A, forced swimming did not influence the body weight changes of 16-month-old mice, and likewise, oral administration of LRC had no effect on body weight changes. However, swimming significantly increased the weight of both the gastrocnemius and soleus muscles compared to the control group, and this increase was mediated by LRC administration relative to the swim group. (Fig. 1D, 1E). These findings indicate that the intensity of swimming and LRC intake do not influence body weight change, and that increases in muscle weight are not associated with body weight change. Furthermore, it also indicates that LRC may alleviate sarcopenia by promoting swimming-induced muscle weight gains.

In the present study, administration of LRC significantly increased grip strength, treadmill running distance, and prolonged the time to exhaustion compared with swim group (Fig. 2A-2C). Grip strength assessment and treadmill running are reliable methods for assessing musculoskeletal health and endurance in animals (Reynolds and Lee, 2020). Our current findings are consistent with previous reports showing enhanced grip strength by Ishige okamurae (IO) extract administration or improved endurance capacity by apple pomade (AP) extract supplementation (Hyun et al., 2022; Jeong et al., 2015). Meanwhile, LRC reduced serum lactate and creatinine levels compared with swim group, respectively (Fig. 2D, 2E). Serum lactate and creatinine are indicators of muscle fatigue and age-related muscle damage (Lee et al., 2021b). There was no significant difference in serum lactate levels between the swim group and control group, but creatinine levels were significantly lower in the swim group compared to the control group (Fig. 2D, 2E). These differences are likely due to the frequency and intensity of swimming performed in this study, and are consistent with meta-analyses showing that plasma creatinine levels were reduced in patients with chronic kidney disease (CKD) when combined with aerobic exercise, such as walking or running, compared to the control group (Ma et al., 2022). These findings support the beneficial role of even low-intensity physical activity in mitigating age-related declines in muscle function and LRC may enhance the physiological adaptations induced by exercise. Collectively, our findings highlight the potential of LRC as a complementary intervention to exercise in promoting muscle performance and reducing fatigue and damage in aging.

MyoD, Myogenin, and MHC are genes involved in muscle differentiation, with MyoD initiating the myogenic program and determining muscle cell lineage (Guo et al., 2023; Liu et al., 2014). Myogenin is crucial for the maturation and terminal differentiation of muscle cells, while MHC is a major contractile protein in skeletal muscle (Mastroyiannopoulos et al., 2012; Short et al., 2005). According to previous studies, extracts of Schisandra or Turmeric have been reported to increase the expression of muscle differentiation-promoting factors in aged mice, thereby demonstrating effectiveness in improving sarcopenia (Kim et al., 2015; Lee et al., 2021a). Our present results found that swimming increased mRNA levels of MyoD and myogenin in soleus muscles (Fig. 3A, 3B), and MHC levels in both soleus and gastrocnemius muscles (Fig. 3C). Furthermore, mRNA levels of MyoD, myogenin, and MHC were significantly increased due to LRC supplementation in both soleus and gastrocnemius muscles compared to the swim group (Fig. 3A-3C). These results suggest that LRC can promote swimming-induced muscle differentiation in aged mice.

Exercise training stimulates protein synthesis through activation of the Akt/mTOR pathway (Bacurau et al., 2016). Akt is a major regulator that controls various cellular functions, including survival, growth, differentiation, metabolism, and migration, and it promotes mTOR phosphorylation (Sugiyama et al., 2019; Xiao et al., 2009). The mTOR is activated by Akt and is involved in various cellular responses, including cell growth, proliferation, and survival (Hahn-Windgassen et al., 2005). Recent studies have revealed that administration of soluble whey protein hydrolysate or fermented Tenebrio molitor larvae extract in immobilization- or DEX-induced muscle atrophy mouse models enhances muscle protein synthesis through activation of the PI3K-Akt-mTOR pathway (Han et al., 2024; Shin et al., 2020). In the present study, swimming significantly increased Akt and mTOR mRNA expression in aged mice, and LRC administration induced an even greater increase (Fig. 3D, 3E). These findings suggest that LRC can enhance swimming-induced muscle synthesis, potentially contributing to the prevention of sarcopenia in aged mice.

Skeletal muscle atrophy induced by aging or DEX inhibits the PI3K-Akt signaling pathway, leading to the activation of forkhead box transcription factors (FoxO) (Wang et al., 2022). FoxO activates muscle-specific E3 ubiquitin ligases, muscle-specific ring finger protein 1 (MuRF-1), and atrogin-1, resulting in increased protein degradation and muscle mass loss (Lee et al., 2020). Additionally, aging and DEX also increase myostatin, an inhibitor of muscle synthesis, causing muscle loss and reduced grip strength (Han et al., 2011; Wang et al., 2016; Yarasheski et al., 2002). Swimming can help reduce these levels in both the soleus and gastrocnemius muscles, and supplementation with LRC significantly enhances the effects of swimming (Fig. 3F-3I). Taken together, our findings suggest that swimming inhibits protein degradation, and LRC enhances the effects of swimming, indicating its potential effectiveness in alleviating sarcopenia.

An increase in muscle cross-sectional area, also known as muscle hypertrophy, means that the muscle fibers have become larger and thicker, resulting in a larger overall muscle mass and size (Haun et al., 2019). As shown in Fig. 4, swim+LRC groups exhibited even more substantial enhancements in muscle fiber cross-sectional area within the gastrocnemius muscle in comparison to the swim group. These results are consistent with the increase in gastrocnemius muscle weight (Fig. 1D) and suggest that LRC has a synergistic effect on muscle remodeling induced by forced swimming.

In a previous study, we demonstrated that Lycii Radicis Cortex (LRC) extract prevented 10 μM dexamethasone (DEX)-induced muscle atrophy in C2C12 myotubes (Son et al., 2024). However, the results regarding the possibility that these effects may be due to the main ingredients KA and KB are not yet known. Although KA and KB, the major components and isomers of LRC extract, have been reported to possess various biological activities, including antioxidant, anti-inflammatory, and anti-cytotoxic effects, their potential role in ameliorating muscle damage has not yet been investigated. We evaluated the preventive effects of KA and KB on muscle damage using C2C12 myotubes and dexamethasone (DEX), which are frequently used as an in vitro model of muscular atrophy. According to our current results, DEX-treated C2C12 myotubes’ survival, diameter, MHC-stained area, fusion index, and MHC protein expression were all dramatically restored by KA or KB treatment (Fig. 5A-5G). Furthermore, KA and KB treatment decreased DEX-induced upregulation of FoxO3a, MuRF1, and atrogin-1 and enhanced Akt phosphorylation decreased by DEX (Fig. 6). These results suggest that KA and KB inhibit DEX-induced cytotoxicity, C2C12 myotubes morphological changes, and protein degradation. These effects are consistent with recent studies showing the efficacy of LRC in ameliorating DEX-induced muscle atrophy (Son et al., 2024).

Treatments for muscle atrophy may involve reducing ROS generation or increasing SOD activity. In C2C12 myotubes, DEX treatment causes an increase in ROS generation and a decrease in SOD activity, which is closely associated with skeletal muscle atrophy (Kim et al., 2023). Our current results showed that KA or KB treatment significantly reduced ROS production in DEX-treated myotubes (Fig. 7A, 7B), and restored SOD activity to control levels (Fig. 7C). These results suggest that KA and KB can reduce muscle atrophy by preventing oxidative damage caused by DEX.

Mitochondrial dysfunction promoted muscle atrophy and DEX-treatment induced skeletal muscle atrophy in C2C12 myotubes by reducing mitochondrial content and ATP production capacity (Huang et al., 2018; Roshanravan et al., 2021). Improving the quantity and quality of mitochondria can be considered an important factor in preventing muscle atrophy. KA and KB restored mitochondrial contents to levels similar to those in the control group, and ATP levels also increased (Fig. 8). According to these findings, KA and KB may help inhibit muscle atrophy by preventing mitochondrial dysfunction in C2C12 myotubes treated with DEX.

In conclusion, this study suggests that LRC and its main active ingredients, KA and KB, have effective ameliorating effects on aging and dexamethasone-induced muscle damage (Fig. 9), which may play an important role in muscle maintenance and functional improvement. In particular, LRC has the potential to be developed as a functional food or natural product-based complementary medicine, and can be used as basic data for clinical trials. However, future studies should include an LRC-only treatment group and a young control group, and expand biomarker analyses to more clearly elucidate the role of LRC in the improvement of sarcopenia.

Fig. 9.

Schematic diagram of this study. LRC increased muscle synthesis and inhibited muscle degradation in aged mice, thereby improving muscle strength and endurance. Its main components, KA and KB, were found to have the effect of inhibiting muscle damage caused by DEX, suggesting that LRC administration is effective in preventing sarcopenia.