Peucedanum japonicum Thunb. ameliorates age-related muscle loss by improving mitochondrial function in aged mice

Abstract

Peucedanum japonicum Thunb. (PJ), also known as costal hog fennel, is an edible medicinal plant recognized for its potential health benefits. We previously demonstrated the protective effects of PJ against muscle atrophy in young mice. In the present study, we investigated whether PJ also inhibits age-related muscle loss in aged mice. PJ treatment of primary myoblasts derived from aged mice attenuated myotube diameter reduction, suppressed the expression of atrogenes, and prevented the fast-to-slow myosin heavy chain (MHC) fiber type transition. Additionally, PJ improved mitochondrial respiratory capacity, accompanied by increased PGC1α expression and phosphorylation. Aged mice (20 months old) were fed diets supplemented with 0.1% or 0.2% PJ for eight weeks. PJ supplementation enhanced muscle strength and physicalrformance parameters, including treadmill endurance and stride length. Increases in lean body mass, muscle weight, cross-sectional area (CSA), and the ratio of MHCII to MHCI fibers were observed. These improvements were associated with reduced ubiquitin-dependent protein degradation and the downregulation of atrogenes. Furthermore, PJ supplementation promoted mitochondrial activity via PGC1α and stimulated mitochondrial biogenesis through the Nrf1–Tfam pathway. 4-Caffeoylquinic acid (4-CQA), a major bioactive compound in PJ, attenuated myotube atrophy and enhanced mitochondrial respiration in aged myoblasts. These findings suggested that PJ, along with its active compound 4-CQA, has therapeutic potential for combating age-related muscle loss.

Highlights

• Peucedanum japonicum Thunb. (PJ) improves myogenic capacity and mitochondrial function of primary myoblasts from aged mice.

• PJ supplementation ameliorates muscle mass and muscle function by suppressing protein degradation in C57BL/6 aged mice.

• PJ supplementation mitigates mitochondrial dysfunction and increases mitochondrial biogenesis in skeletal muscle of C57BL/6 aged mice.

• 4-caffeoylquinic acid, a bioactive compound in PJ, enhances myogenic capacity and mitochondrial function in primary myoblasts from aged mice.

Graphical Abstract

Introduction

Sarcopenia is the progressive reduction of muscle mass and function with aging, resulting in frailty and an increased risk of mortality. It is characterized by the progressive loss of muscle mass, strength, and function with age. Muscle loss is associated with disability, impairment, diseases, and mortality, consequently reducing quality of life [1]. Currently, there is no drug for sarcopenia, so preventive approaches such as protein supplementation are important. While the daily recommended dose for healthy adults is 0.8 g/kg, aging adults at risk of sarcopenia are advised to consume up to 1.2 g/kg per day [2]. However, excessive protein intake increases the risk of kidney dysfunction, digestive disorders, and osteoporosis [3]. Therefore, alternative strategies for preventing muscle atrophy are emerging.

The number and size of skeletal muscle fibers decrease with age; as a result, skeletal muscle function and mass are reduced [4]. Muscle fibers are classified as Type I (slow-twitch) or Type II (fast-twitch) based on their contractile speed in response to nerve stimulation [5]. Type I and IIA fibers rely on mitochondrial oxidative phosphorylation, whereas Type IIB and IIX fibers rely on glycolysis for energy production [6]. During muscle aging, the size and number of Type II fibers gradually decrease, which is accompanied by a fast-to-slow twitch transition [7]. Additionally, the muscle-type shift associated with muscle aging results in changes in skeletal muscle reliance from cytosolic glycolytic metabolism to oxidative metabolism, leading to an increase in the demand for functional mitochondria [8].

Mitochondria play key roles in cellular homeostasis, inflammation, and senescence. Mitochondrial dysfunction is a key contributor to muscle degeneration in aging. During aging, mitochondrial features change, affecting protein homeostasis, and ultimately leading to muscle loss [9, 10]. In addition, damaged mitochondria result in the impairment of oxidative phosphorylation (OXPHOS), ROS generation, and decreased ATP production, leading to a vicious cycle of muscle atrophy [9, 11]. Mitochondrial dysfunction has been reported to cause the early emergence of several age-related phenotypes, including sarcopenia, suggesting that inhibition of mitochondrial dysfunction may be effective in preventing sarcopenia [9, 12].

Peucedanum japonicum Thunb. (PJ) belongs to the Umbelliferae family and is cultivated in East Asia. The leaves are consumed as vegetables, and the roots are used for herbal medicine [13]. PJ has been reported to exhibit anti-cancer, anti-allergic, antioxidant, and anti-obesity effects and it contains various phytochemicals, including rutin, coumarins, and caffeic acid derivatives [14–16]. Among caffeic derivatives, 4-caffeoylquinic acid (4-CQA) has been reported to exert antioxidant activity [17]. We previously reported that PJ prevents dexamethasone-induced muscle atrophy, with 4-CQA as the key functional compound responsible for these effects [18]. Since oxidative stress is recognized as one of the major underlying mechanisms of sarcopenia, the abundant presence of 4-CQA in PJ could contribute to its protective effects against muscle atrophy. However, the effects of PJ and 4-CQA, its bioactive compound, on age-related muscle loss remain unclear.

In this study, we revealed that 4-CQA-enriched PJ alleviates age-related muscle loss and mitochondrial dysfunction. We then evaluated the effects of PJ and 4-CQA on myogenesis and mitochondrial function in aged primary myoblast cells. In addition, the effects of PJ supplementation on age-related muscle loss were demonstrated in aged mice.

Materials and methods

Material preparation

PJ leaves were collected in Goheung, Korea, dried at 60℃, and ground into a fine powder using a grinder prior to extraction. The powdered material was subsequently extracted with 50% ethanol at 80℃ for 3 h. After filtration, the extract was concentrated using a rotary vacuum evaporator (Büchi Labortechnik AG, Flawil, Switzerland) and then lyophilized. The extraction yield was 20.8%.

Cell culture

Mouse myoblast cells were isolated as previously described [19]. Briefly, 7-month-old (young) and 22-month-old (old) C57BL/6 mice were euthanized and dissected to isolate skeletal muscles. The muscles were mechanically dissociated and enzymatically digested using a skeletal muscle dissociation kit (130-098-305, Miltenyi Biotec, Gergisch Gladbach, Germany). The isolated myoblasts were maintained in high-glucose DMEM (HyCloneTM, Logan, UT, USA), supplemented with 20% fetal bovine serum (FBS, HyCloneTM), 10% horse serum (HS, HyClonTM), 0.5% chicken embryo extract, 1% antibiotic-antimycotic (AA, Sigma-Aldrich, Ronkonkoma, NY, USA), 2.5 ng/mL bFGF, and 10 ug/mL gentamicin. For differentiation, cultured myoblasts were exposed to high-glucose DMEM (HyClonTM) containing 5% HS (HyCloneTM) and 1% AA (Sigma-Aldrich) for four days. The differentiated myotubes were then treated with PJ or 4-CQA for 24 h.

Measurement of tube diameter

To fix the differentiated myoblasts, 4% formaldehyde was applied for 30 min, followed by permeabilization with 0.1% saponin in PBS for 30 min and blocking with 1% bovine serum albumin (BSA) in PBS for 30 min. Then, myotubes were stained with 1:200 total myosin heavy chain (MHC) primary antibody (MF20, DSHB, University of Iowa, Iowa City, IA, USA) overnight at 4℃, followed by incubation with a 1:100 dilution of Alexa Fluor-488-conjugated secondary antibody (Cell Signaling Biotechnology, Beverly, MA, USA) at 25℃ for 30 min. The nucleus was stained with 1:10000 4’,6-diamidino-2-phenylindole (DAPI) at 25℃ for 1 min. Images were obtained using an Olympus IX71 microscope and Olympus DPO controller 3.1.1 software. Images were analyzed with ImageJ software (NIH, Bethesda, MD, USA), and myotube diameters were assessed by averaging five individual measurements.

Total RNA extraction and qRT-PCR

Total RNA was extracted from myoblasts and skeletal muscles using NucleoSpin RNA II (Macherey-Nagel, Düren, Germany) and the RNeasy mini kit (Qiagen, Hilden, Germany), respectively. cDNA synthesis was performed using ReverTra Ace qPCR RT Master Mix (Toyobo, Osaka, Japan) in a Thermal Cycler (Bio-Rad, Hercules, CA, USA). qRT-PCR was performed using SYBR Green Master Mix (Toyobo) on a ViiA7 system (Applied Biosystems, Foster City, CA, USA). The relative mRNA expression levels were normalized to 18S using 2^-ΔΔCt method. The primer sequences are listed in Table 1.

Table 1.

Primer sequences for qRT-PCR

Gene	Primer Sequences
Atrogin-1	Forward Reverse	5'- AAGGCTGTTGGAGCTGATAGCA −3' 5'- CACCCACATGTTAATGTTGCCC −3'
Murf1	Forward Reverse	5'- TGTCTCACGTGTGAGGTGCCTA −3' 5'- CACCAGCATGGAGATGCAGTTAC −3'
Myostatin	Forward Reverse	5'- ACG CTA CCA CGG AAA CAA TC −3' 5'- GGA GTC TTG ACG GGT CTG AG −3'
Nrf1	Forward Reverse	5'- AATGTCCGCAGTGATGTCC −3' 5'- GCCTGAGTTTGTGTTTGCTG −3'
Nrf2	Forward Reverse	5'- TGAAGTTCGCATTTTGATGGC −3' 5'- CTTTGGTCCTGGCATCTCTAC −3'
Tfam	Forward Reverse	5'- CACCCAGATGCAAAACTTTCAG −3' 5'- CTGCTCTTTATACTTGCTCACAG −3'
Ndufv1	Forward Reverse	5'- ATC GCT CGA CAG ACA TTG TG −3' 5'- GTG GCC TTC TAT CTG CTT GC −3'
Ndufs3	Forward Reverse	5'- CTG ACT TGA CGG CAG TGG AT −3' 5'- CAT ACC AAT TGG CCG CGA TG −3'
Ndufb3	Forward Reverse	5'- AAG GGA CGC CAT TAG AAA CG −3' 5'- TAC CAC AAA CGC AGC AAA CC −3'
Ndufs8	Forward Reverse	5'- GTTCATAGGGTCAGAGGTCAAG −3' 5'- TCCATTAAGATGTCCTGTGCG −3'
Sdhb	Forward Reverse	5'- ACCCCTTCTCTGTCTACCG −3' 5'- AATGCTCGCTTCTCCTTGTAG −3'
Sdhd	Forward Reverse	5'- GAT CCC TGC TGG GTA CTT GA −3' 5'- AAG TAG CAA AGC CCA GCA AA −3'
Uqcrc1	Forward Reverse	5'- ATCAAGGCACTGTCCAAGG −3' 5'- TCATTTTCCTGCATCTCCCG −3'
Cox5b	Forward Reverse	5'- ACCCTAATCTAGTCCCGTCC −3' 5'- CAGCCAAAACCAGATGACAG −3'
Atp5a	Forward Reverse	5'- CATTGGTGATGGTATTGCGC −3' 5'- TCCCAAACACGACAACTCC −3'
18s	Forward Reverse	5'- CTCAACACGGGA AACCTCAC −3' 5'- CGCTCCACCAACTAAGAACG −3'

PGC-1α, peroxisome proliferator-activated receptor gamma coactivator 1-alpha; NRF1 nuclear respiratory factor 1; NRF2, nuclear factor erythroid-derived 2-related factor 2; TFAM, mitochondrial transcription factor A; NDUFV1, NADH dehydrogenase [ubiquinone] flavoprotine 1; NDUFB3, NADH dehydrogenase [ubiquinone] 1 beta subcomplex 3; NDUFS8, NADH dehydrogenase [ubiquinone] iron-sulfur protein 8; SDHB, succinate dehydrogenase [ubiquinone] iron-sulfur subunit; SDHD, succinate dehydrogenase complex subunit D; UQCRC1, ubiquinol-cytochrome c reductase core protein 1; COX5b, cytochrome c oxidase subunit 5β; ATP5a, ATP synthase subunit alpha; 18S, 18S ribosomal

Protein extraction and western blot

Protein extraction from myoblasts and quadriceps muscle tissues was performed with RIPA (Thermo Fisher Scientific, Rockford, IL, USA), which contains protease and phosphatase inhibitors. A total of 10 μg of protein separated using SDS-PAGE were transferred to polyvinylidene fluoride membranes (Bio-Rad). The membranes were blocked with 5% BSA in TBST for 1 h and then incubated with primary antibodies in TBST (Tris buffered saline with 0.1% Tween 20) overnight at 4℃. Following washes with TBST, the membranes were incubated with a horseradish peroxidase (HRP)-conjugated secondary antibody. Protein bands were developed using chemiluminescent HRP substrates (Thermo Fisher Scientific) and subsequently quantified using ImageJ software (NIH). The primary antibodies used are listed in Table 2.

Table 2.

Primary antibody for western blot

Antibody	Source	Dilution
Total MHC	DSHB	1:1000
MHCⅠ	DSHB	1:1000
MHC2A	DSHB	1:1000
MHC2B	DSHB	1:200
Atrogin-1(Fbx32)	Abcam	1:1000
MuRF1	Abcam	1:1000
Ubiquitin	Cell signaling	1:1000
PGC1α	Abcam	1:1000
p-PGC1α	R&D systems	1:1000
NCoR1	Cell signaling	1:1000
α-tubulin	Santa Cruz	1:1000
β-actin	Santa Cruz	1:1000

Determination of oxygen consumption rate (OCR) in primary mouse myoblasts

OCR was measured using an XF Cell Mito Stress Test Kit (Seahorse Bioscience, Billerica, MA, USA) and an XF24 extracellular Flux Analyzer (Seahorse Bioscience), according to the manufacturer’s protocol. Briefly, the primary myoblast cells were seeded in XF24 cell culture microplate coated with matrigel and then treated with PJ or 4-CQA for 24 h. After 24 h, the culture medium was changed to assay medium with 25 mM glucose, 1 mM pyruvate, and 1 mM glutamine and incubated at 37℃ for 1 h. OCR measurements were obtained by sequentially injecting 1.5 μM oligomycin, 4 μM FCCP, and 1 μM antimycin A.

Animal experiments

Eight-week-old male C57BL/6 mice were obtained from Orient Bio, Inc. (Seongnam, Korea). The mice were housed in an environment with a controlled temperature of 21−23℃ and a 12-hour light/12-hour dark cycle. Following one week of acclimatization, 28 mice were randomly divided into four groups (n = 7/group): Ⅰ) Young control group (Young), Ⅱ) Aged control group (Aged), Ⅲ) 0.1% PJ-supplemented group (PJL), and IV) 0.2% PJ-supplemented group (PJH). The young and aged control groups were fed an AIN-93M diet (D10012M; Research Diet, Inc., New Brunswick, NJ, USA). The PJL and PJH groups were fed the AIN-93M diet (Research Diet) containing 0.1% and 0.2% PJ supplementation, respectively. All experimental groups were fed the diet for eight weeks. Five-month-old (young) and 20-month-old (aged) male C57BL/6 mice were used in the study. The animals were humanely sacrificed using 5% isoflurane, and muscle tissues to be used in the experiment were collected. All animal experiments were performed in accordance with the guidelines of the Institutional Animal Care and Use Committee of the Korea Food Research Institute (KFRI-IACUC, KFRI-M-20018).

Measurements of muscle mass, function, and performance

Body composition was measured using the InAlyzer (Medikors, Korea). Mouse grip strength was measured five times using BIO-GS3 (Bioseb, Chaville, France) to evaluate skeletal muscle function, and results were standardized by body weight. To determine skeletal muscle performance, the total running distance (m) and running time (min) were measured on a rodent treadmill (Ugo Basile, Varese, Italy). Mice were acclimatized for 20 min at a speed of 10 m/min, 10% incline, and 0.1 mA shock for two days. On the third day, the mice were examined at speeds of 10 m/min, 10% incline, and 0.1 mA shock, with the speed increasing 2 m/min every 2 min. The maximal running time did not exceed 20 min. The mice stopped running when they remained on the shock grid for 10 s.

Step length

Step length was measured using a method modified from a previous study [20]. Mouse paws were painted with commercial ink and allowed to walk on a strip of paper (6 cm wide, 100 cm long). After the ink dried, stride lengths were measured, and the average was calculated.

Histological analysis

The gastrocnemius muscles were embedded in Tissue-Tek optimal cutting temperature (OCT) compound (Sakura Finetek, Tokyo, Japan) using a cryomold and frozen in liquid nitrogen. The frozen muscles were stored at −80℃ until use. For immunohistochemical analysis, the gastrocnemius muscles were sectioned and stained with laminin to measure cross-sectional area (CSA) and measured muscle fiber type transition with MHC isoforms [21]. The stained tissues were imaged using a confocal microscope (Digital Eclipse C1 plus, Nikon, Tokyo, Japan). For nicotinamide adenine dinucleotide (NADH) staining, sectioned gastrocnemius muscles were incubated with nitro blue tetrazolium (NBT) solution (2 mg/mL NBT in 0.05 M Tris buffer, pH 7.6) and NADH solution (1.6 mg/mL NADH in Tris buffer, pH 7.6) for 30 min at 37℃. Then, the muscles were washed three times with tap water and unbound NBT was removed using sequential washes with 30, 60, and 90% acetone solutions. After rinsing with tap water three more times, the stained muscles were mounted with a cover slip. The stained tissues were imaged using a confocal microscope (Digital Eclipse C1 Plus, Nikon).

Statistical analysis

Data were analyzed using GraphPad Prism software (version 8.0; GraphPad Software, San Diego, CA, USA). Statistical results are expressed as mean ± standard deviation (SD) for in vitro experiments or mean ± standard error of the mean (SEM) for in vivo experiments. One-way analysis of variance (ANOVA) was used to compare quantitative data among groups, followed by Dunnett’s multiple comparison test. A p-value of < 0.05 was considered statistically significant.

Results

PJ improves myogenesis and mitochondrial function in primary myotubes from aged mice

Muscle regeneration and myogenic capacity depend on satellite cells and decrease with age [22]. To investigate the effect of PJ on myogenesis, we induced myogenic differentiation of primary myoblasts using PJ. PJ treatment effectively improved the myogenic capacity of aged primary myoblasts (Fig. 1a and b). The mRNA expression of atrophy-related genes, including Atrogin-1 and Murf1, was upregulated in aged primary myoblasts. Moreover, PJ treatment markedly decreased the levels of atrophy-related genes (Fig. 1c). Western blotting was performed to assess the protein expression of MHC isotypes (Fig. 1d and e). In aged myotubes, the expression of MHC1 increased and the expression of MHC2A and MHC2B decreased. However, PJ tended to increase MHC2A expression, and PJ markedly upregulated MHC2B expression in a dose-dependent manner (Fig. 1e). Consequently, the elevated ratio of MHC1/MHC2B in aged primary mouse myotubes was dose-dependently reduced following PJ treatment (Fig. 1f). These results suggested that PJ inhibited aging-related fast-to-slow MHC type transitions. Next, we measured OCR and the protein expressions of p- PGC1α, PGC1α and NCoR1 in primary myotubes from aged mice (Fig. 1g-j). Basal and maximal respiration were reduced and effectively increased with PJ treatment. PGC1α and NCoR1 have a key role in mitochondria function and mitochondria homeostasis and PGC1α activity is regulated by phosphyrylation. PGC1α is a coactivator and NCoR1 is a corepressor of mitochondria, acting in opposition to each other [23]. We measured the protein expression of p- PGC1α, PGC1α and NCoR1 using western blot. p- PGC1α/PGC1α ratio was increased by PJ treatment in primary myotube from aged mice (Fig. 1i and j). These results showed that PJ treatment improved myogenic capacity and mitochondrial function in primary myotubes from aged mice.

Fig. 1.

Effect of PJ on myogenic capacity and mitochondrial function of primary myoblasts from aged mice. (a) Immunofluorescence staining of total MHC (green) and DAPI (blue) in differentiated primary mouse myoblasts. (b) Mean myotube diameter (μm). (c) The mRNA expressions of Atrogin-1 and Murf1 in myotubes. (d) The expressions of MHC isoforms detected by western blot analysis in myotubes. (e) The quantification of (d). (f) MHC1/MHC2B ratio in myotubes. (g) Extracellular flux analysis to measure the oxygen consumption rate (OCR) in primary mouse myoblasts. (h) Mitochondrial parameters were calculated from the mitochondrial stress profiles. (i) The protein expressions of p- PGC1α, PGC1α and NCoR1 in myotubes. (j) The quantification of (i). Values are mean ± SD. *P < 0.05, ** P < 0.01, and *** P < 0.001; one-Way ANOVA compared with Aged (without any treatment)

PJ supplementation ameliorates aging-related muscle loss in aged C57BL/6 mice

To examine whether PJ ameliorates aging-related muscle loss, we fed experimental diets containing 100 mg/kg (PJL) or 200 mg/kg (PJH) PJ to 20-month-old aged mice for eight weeks. PJ supplementation increased the body weight of the mice (Fig. 2a). Moreover, Dual-Energy X-ray Absorptiometry (DEXA) analysis showed an increase in lean body mass and a decrease in fat mass in PJ-supplemented aged mice (Fig. 2b and c). We performed grip strength and treadmill tests to measure muscle function and physical performance, respectively (Fig. 2d-f). PJH markedly enhanced muscle strength and physical performance in aged mice. In addition, PJ supplementation effectively prevented the reduction in stride length (Fig. 2g). Consistent with the observed decrease in lean body mass and muscle function, skeletal muscle weight was reduced in aged mice. Unlike the results for muscle performance and function, an increase in muscle weight was only observed in the quadriceps muscle following PJH supplementation (Fig. 2h). Based on these findings, subsequent in vivo analyses were mainly focused on the PJH supplemented group.

Fig. 2.

Effect of PJ supplementation on age-related muscle loss in aged C57BL/6 mice. (a) The final body weights (g). (b and c) Lean body mass (g) and fat mass (g) in aged mice were measured by Dual-Energy X-ray Absorptiometry (DEXA). (d-e) Muscle performance was measured by treadmill. Running distance (m) and running time (min) were shown. (f) Muscle strength was measured by grip strength (g/body weights). (g) The stride length (cm) was measured. (h) The skeletal muscle weights normalized by body weights (mg/g). (i) The representative images of quadriceps muscle by immunofluorescent staining for laminin. (j) The mean cross sectional area (CSA) of quadriceps muscles myofibers. (k) The distribution of CSA of myofibers (l) Immunohistochemical staining of quadriceps muscles for the triple-labeling with MHC1 (blue), MHC2A (green), and MHC2B (red). (m) The ratio of MHC isoforms. Values are mean ± SEM. *P < 0.05, ** P < 0.01, and *** P < 0.001; One-Way ANOVA compared with aged

To demonstrate the inhibitory effect of PJH on aging-induced CSA reduction and fast-to-slow transition, we performed immunohistochemical staining. CSA was measured through laminin staining of the muscle tissue (Fig. 2i). Similar to the changes in muscle mass, CSA was decreased in aged mice compared to young mice (Fig. 2j). However, the PJH group showed a significant increase in CSA. In young mice, the most common fiber was distributed between 2,000 and 2,500 μm². However, in aged mice, the most common fiber was distributed between 500 and 1000 μm² (Fig. 2k). The left shift in the distribution of muscle fiber size was ameliorated in the PJH group.

To investigate the effects of PJ on aging-induced Type II fiber damage, we performed immunohistochemical staining of the muscle fiber type. PJH effectively prevented aging-related MHC2 loss (Fig. 2l and m). These results indicated that PJ inhibited age-related skeletal muscle loss and fast-to-slow muscle type transition, and enhanced muscle function in aged mice, suggesting the suppression of age-related muscle loss.

PJ reduced ubiquitin-dependent protein degradation in aged muscles

We investigated the effect of PJ on muscle protein degradation by measuring the ubiquitination of muscle proteins and mRNA levels of genes related to muscle atrophy in aged mice. The increase in polyubiquitinated proteins in aged muscles was suppressed by PJH supplementation (Fig. 3a). In aging muscles, atrophy is associated with proteolysis via the ubiquitin–proteasome pathway, and key genes, including Atrogin-1 and MuRF1, are involved [24]. Consistent with the increase in ubiquitinated proteins, the mRNA levels of muscle atrophy-related genes such as Atrogin-1, Murf1, and Mstn were upregulated in the quadriceps muscle of aged mice, which were effectively downregulated by PJH supplementation (Fig. 3b). These results were consistent with those of the in vitro experiments using primary myotubes from aged mice. The increased protein expression of Atrogin-1 and MuRF1 was notably decreased by PJH supplementation (Fig. 3c and d). These results demonstrated that PJ mitigated aging-associated proteolysis and protein degradation.

Fig. 3.

Effect of PJ on ubiquitin-dependent protein degradation in aged muscle. (a) The poly-ubiquitinated protein was detected by western blotting in quadriceps muscles. (b) The mRNA expressions of Atrogin-1, Murf1, and Mstn (Myostatin) in quadriceps muscles. (c) The protein expressions of Atrogin-1 and MuRF1 in quadriceps muscles. (d) The quantification of (c). Values are mean ± SEM. *P < 0.05, ** P < 0.01, and *** P < 0.001; One-Way ANOVA compared with Aged

PJ supplementation improved mitochondrial dysfunction and increased mitochondrial biogenesis in skeletal muscles of aged C57BL/6 mice

To explore the impact of PJ on mitochondrial function in aged mice, we measured total ATP production and citrate synthase activity (Fig. 4a and b). Aged muscles showed decreased ATP content and citrate synthase activity. However, PJH supplementation effectively increased ATP content and modestly increased citrate synthase activity.

Fig. 4.

Effect of PJ supplementation on mitochondria function and mitochondrial biogenesis in skeletal muscle of aged C57BL/6 mice. (a) Total ATP content (pmol) was measured in quadriceps muscles. (b) Citrate synthase activity (μmole/min/mg) was measured. (c) The representative images of NADH staining (d) Percentages of NADH positive fiber (%). (e) The mtDNA contents (f) The mRNA expressions of mitochondrial complex genes (g) The mRNA expressions of mitochondrial biogenesis related genes (h) The protein expressions of PGC1α and NCoR1 (i) The quantification of (h). (j) PGC1α/NCoR1 ratio. Values are mean ± SEM. *P < 0.05, ** P < 0.01, and *** P < 0.001; One-Way ANOVA compared with aged

Moreover, PJH supplementation significantly (P < 0.001) ameliorated aging-related NADH reduction (Fig. 4c and d). The reduction of mtDNA content in aged mice was reversed by PJH supplementation (Fig. 4e). These data showed that PJ improved both the mitochondrial function and quantity in aged mice. The mRNA expression of OXPHOS genes was downregulated in aged mice muscles, and PJ effectively upregulated the levels of complexes I, II, III, and IV (Fig. 4f). We also observed that the mRNA expression levels of Nrf1, Nrf2, and Tfam, genes related to mitochondrial biogenesis, were downregulated with aging, and were effectively restored by PJH supplementation (Fig. 4g). Moreover, we measured the protein expression levels of PGC1α and NCoR1 using western blot analysis (Fig. 4h-j). PJ supplementation led to a trend toward increased PGC1α expression and a significant decrease in NCoR1 levels. The reduced PGC1α/NCoR1 ratio observed in aged muscle was elevated by PJH supplementation (Fig. 4j). These findings indicated that PJ improved mitochondrial function and stimulated mitochondrial biogenesis by regulating PGC1α and NCoR1 expression in aged mice.

4-CQA promotes myogenic capacity and mitochondrial function in primary myoblasts from aged mice

Next, we investigated whether 4-CQA is the functional component of PJ responsible for the mitigation of muscle aging. We treated aged myoblasts with 4-CQA and induced their differentiation. 4-CQA significantly (P < 0.05) increased myogenesis in a dose-dependent manner (Fig. 5a and b). Conversely, the mRNA expression levels of Atrogin-1 and Murf1 decreased in a dose-dependent manner after 4-CQA treatment (Fig. 5c). 4-CQA suppressed the elevation of MHC1 expression, which increased with aging, leading to a reduction in the MHC1/MHC2B ratio (Fig. 5d-f). These results demonstrated that 4-CQA improved myogenesis in primary myoblasts from aged mice. In addition, 4-CQA markedly increased maximal respiration, basal respiration, and spare respiratory capacity, and elevated the protein expression ratio of p- PGC1α/PGC1α (Fig. 5g-j). These data showed that 4-CQA improved mitochondrial function in primary myotubes derived from aged mice. Taken together, our results suggested that 4-CQA contributes to the protective effects of PJ against muscle aging.

Fig. 5.

4-CQA improves myogenesis and mitochondrial function in aged mouse primary myotubes. (a) Immunofluorescence staining of total MHC (green) and DAPI (blue) in differentiated myotubes. (b) Mean myotube diameter (μm). (c) The mRNA expressions of Atrogin-1 and Murf1. (d) The expressions of MHC isoforms detected by western blot analysis. (e) The quantification of (d). (f) MHC1/MHC2B ratio in aged-mice myotubes. (g) Extracellular flux analysis to measure the oxygen consumption rate (OCR) in primary mouse myoblasts. (h) Mitochondrial parameters were calculated from the mitochondrial stress profiles. (i) The protein expressions of p- PGC1α, PGC1α and NCoR1 in myotubes. (j) The quantification of (i). Values are mean ± SD. *P < 0.05, ** P < 0.01, and *** P < 0.001; One-Way ANOVA compared with aged (without any treatment)

Discussion

In this study, we presented both in vitro and in vivo evidence that PJ effectively ameliorated muscle aging in mice. PJ improved myogenic differentiation capacity and mitochondrial respiration in primary aged myoblasts via PGC1α activation. Dietary supplementation with PJ mitigated age-related muscle loss, particularly in fast-twitch MHC2 myofibers. Moreover, it improved muscle strength and physical performance, as evidenced by grip strength and stride length, respectively. PJ supplementation reduced ubiquitin-dependent protein degradation and increased mitochondrial function in the skeletal muscles of aged mice. 4-CQA is the bioactive compound in PJ responsible for preventing muscle aging.

Sarcopenia includes muscle fiber atrophy and the transition from fast-type MHC (MHC2X and MHC2B) to slow-type MHC (MHC1). Depending on the cause of muscle atrophy, muscle fiber type transitions show different patterns. The loss of MHC II (fast-twitch fibers) is considered more susceptible to age-related muscle atrophy [25–27]. In aged human muscles, Type 2X fibers almost disappear, while type 2A and type 2A-2X hybrid fibers show no significant change, and the proportion ofType 1 fibers is notably increased [28]. In the present study, we observed muscle loss and a fast-to-slow muscle type transition in aged mice. However, PJ supplementation led to increased muscle mass and inhibited the fast-to-slow type transition in aged mice.

In aged muscles, a fast-to-slow transition is observed with an increase in oxidative-type fibers. However, glycolytic enzyme expression is increased and PGC1α, a key marker of mitochondrial biogenesis, is decreased [28]. These results indicated that despite an increase in oxidative-type fibers, glycolysis is increased, and mitochondrial function is damaged in aged muscles. Consistent with our results, primary skeletal muscle cells isolated from elderly individuals exhibit lower mitochondrial respiratory function compared with those derived from young individuals [29]. Activation of PGC1α has been shown to protect mitochondrial function and increase mitochondrial biogenesis. This leads to enhance the expression of genes involved in oxidative phosphorylation and elevating OCR [30, 31]. PGC1α binds to target nuclear transcription factors such as Nrf1 and Nrf2 to induce the expression of Tfam, which in turn promotes mtDNA replication and transcription [32]. Although PGC1α is regulated through phosphorylation, total PGC1α expression is widely used as an indicator of mitochondrial biogenic capacity [30, 33]. In our study, PJ treatment increased mitochondrial OXPHOS respiration and induced PGC1α in aged myoblasts. Moreover, PJ supplementation in aged mice increased PGC1α expression in skeletal muscle, leading to improved mitochondrial content and function.

During aging, the disruption of mitochondrial biogenesis, proteostasis, and dynamics leads to the accumulation of dysfunctional mitochondria, which is considered a key contributor to the multifaceted pathophysiology of sarcopenia [34]. As mitochondrial dysfunction occurs with aging, NADH levels in the mitochondrial electron transport chains and OXPHOS, which generates ATP through mitochondrial metabolism, decline [35, 36]. The age-associated decrease in the OXPHOS complex results in decreased ATP production and compromised mitochondrial dysfunction in skeletal muscles [37]. In the present study, PJ supplementation ameliorated age-related muscle loss, which was accompanied by increased OXPHOS and ATP contents. This showed that PJ supplementation enhanced overall mitochondrial function in aged mice.

The accumulation of damaged proteins, such as abnormal aggregates, ubiquitinated proteins, and misfolding forms, is observed in aged muscles and is related to age-associated diseases, including Alzheimer’s disease, Parkinson’s disease, and muscle atrophy [38]. This accumulation results from disrupted proteostasis, which is mediated by the ubiquitin–proteasome system (UPS) [39]. The UPS plays a pivotal role in maintaining proteostasis by degrading misfolded and aggregated proteins [40]. However, UPS dysfunction is observed in aged skeletal muscles during the early stages of aging. UPS dysfunction causes an imbalance between protein synthesis and degradation, eventually leading to loss of muscle mass [41]. Substrate proteins are ubiquitinated by covalent linking, a process that includes muscle-specific E3 ligases, such as Atrogin-1 and MuRF1 [42]. In aged muscles, the level of 26S proteasome is elevated two- to five-fold [43]. In this study, we found that PJ suppressed the aging-associated upregulation of atrophic genes and the increase of polyubiquitinated proteins. These results demonstrated that PJ delays muscle loss by maintaining protein homeostasis in aged mice.

Oxidative stress-induced mitochondrial dysfunction is widely recognized as a fundamental mechanism in the progression of sarcopenia [34]. The potent antioxidant activity of 4-CQA may help attenuate reducing ROS production and protecting mitochondrial function. 4-CQA has been reported to have antioxidant, anti-inflammatory, and anti-hypertrophy effects in the heart [44]. In a previous study, we reported that 4-CQA, an abundant component of PJ, could prevents dexamethasone-induced muscle atrophy [18]. We further demonstrated that 4-CQA is the functional component of PJ responsible for the amelioration of age-related muscle loss and improved the myogenic capacity and mitochondrial dysfunction in aged myoblasts.

In conclusion, our findings revealed that PJ attenuated age-related muscle atrophy by promoting myogenesis and preventing UPS and mitochondrial dysfunction. Moreover, we identified 4-CQA as a functional compound in PJ. Our results provided the first evidence that PJ effectively alleviates age-related muscle loss.

Author contributions

Young In Kim: Methodology, Resources, Investigation, Formal analysis, Visualization, Writing. Young-soo Kim: Conceptualization, Investigation. Chang Hwa Jung: Conceptualization, Investigation. Ahn Jiyun Ahn: Conceptualization, Project administration, Investigation, Writing.

Funding

This work was supported by the Korea Food Research Institute [grant number E0210101].

Data Availability

Data will be made available from the corresponding author upon reasonable request.

Declarations

Conflict of interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.