Effects of Ulmus macrocarpa Extract and Catechin 7-O-β-D-apiofuranoside on Muscle Loss and Muscle Atrophy in C2C12 Murine Skeletal Muscle Cells

Abstract

Muscle atrophy is known to be one of the symptoms leading to sarcopenia, which significantly impacts the quality of life, mortality, and morbidity. Therefore, the development of therapeutics for muscle atrophy is essential. This study focuses on addressing muscle loss and atrophy using Ulmus macrocarpa extract and its marker compound, catechin 7-O-β-D-apiofuranoside, by investigating their effects on biomarkers associated with muscle cell apoptosis. Additionally, protein and gene expression in a muscle atrophy model were examined using Western blotting and RT-PCR. Ulmus macrocarpa has been used as food or medicine due to its safety, including its roots, barks, and fruit. Catechin 7-O-β-D apiofuranoside is an indicator substance of plants of the Ulmus genus and has been reported to have various effects such as antioxidant and anti-inflammatory effects. The experimental results demonstrated that catechin glycoside and Ulmus macrocarpa extract decreased the expression of the muscle-degradation-related proteins Atrogin-1 and Muscle RING-Finger protein-1 (MuRF1) while increasing the expression of the muscle-synthesis-related proteins Myoblast determination (MyoD) and Myogenin. Gene expression confirmation experiments validated a decrease in the expression of Atrogin and MuRF1 mRNA and an increase in the expression of MyoD and Myogenin mRNA. Furthermore, an examination of muscle protein expression associated with the protein kinase B (Akt)/forkhead box O (FoxO) signaling pathway confirmed a decrease in the expression of FoxO, a regulator of muscle protein degradation. These results confirm the potential of Ulmus macrocarpa extract to inhibit muscle apoptosis, prevent muscle decomposition, and promote the development of functional materials for muscle synthesis, health-functional foods, and natural-product-derived medicines.

Full text

Citation: Kim, M.S.; Park, S.; Kwon,
Y.; Kim, T.; Lee, C.H.; Jang, H.; Kim,
E.J.; Jung, J.I.; Min, S.; Park,
K.‑H.; et al. Eects of Ulmus
macrocarpa Extract and Catechin
7‑O‑β‑D‑apiofuranoside on Muscle
Loss and Muscle Atrophy in C2C12
Murine Skeletal Muscle Cells. Curr.
Issues Mol. Biol. 2024,46, 8320–8339.
hps://doi.org/10.3390/cimb46080491
Academic Editor: Claudia
Camerino
Received: 21 June 2024
Revised: 27 July 2024
Accepted: 29 July 2024
Published: 1 August 2024
Copyright: © 2024 by the authors.
Licensee MDPI, Basel, Swierland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Aribution (CC BY) license (hps://
creativecommons.org/licenses/by/
4.0/).
Article
Eects of Ulmus macrocarpa Extract and Catechin
7‑O‑β‑D‑apiofuranoside on Muscle Loss and Muscle Atrophy in
C2C12 Murine Skeletal Muscle Cells
Min Seok Kim 1, Sunmin Park 1, Yeeun Kwon 1, TaeHee Kim 1, Chan Ho Lee 2, HyeonDu Jang 2, Eun Ji Kim 3,
Jae In Jung 3, Sangil Min 4, Kwang‑Hyun Park 5and Sun Eun Choi 1,2,*
1Dr. Oregonin Inc., #802 Bodeum Hall, Kangwondaehakgil 1, Chuncheon 24341, Republic of Korea;
ms23217@naver.com (M.S.K.); dpkssm0929@naver.com (S.P.); kye0519@naver.com (Y.K.);
kth02120@naver.com (T.K.)
2Department of Forest Biomaterials Engineering, Kangwon National University, Chuncheon 24341,
Republic of Korea; lgh4107@naver.com (C.H.L.); wkdgusen98@naver.com (H.J.)
3Industry Coupled Cooperation Center for Bio Healthcare Materials, Hallym University, Chuncheon 24252,
Republic of Korea; myej4@hallym.ac.kr (E.J.K.); jungahoo@hallym.ac.kr (J.I.J.)
4Division of Transplantation and Vascular Surgery, Department of Surgery, Seoul National University
Hospital, Seoul 03080, Republic of Korea; surgeonmsi@gmail.com
5Department of Emergency Medical Rescue, Nambu University, Gwangju 62271, Republic of Korea;
pkh15129@gmail.com
*Correspondence: oregonin@kangwon.ac.kr; Tel.: +82‑10‑2352‑8496; Fax: +82‑33‑902‑9990
Abstract: Muscle atrophy is known to be one of the symptoms leading to sarcopenia, which signif‑
icantly impacts the quality of life, mortality, and morbidity. Therefore, the development of thera‑
peutics for muscle atrophy is essential. This study focuses on addressing muscle loss and atrophy
using Ulmus macrocarpa extract and its marker compound, catechin 7‑O‑β‑D‑apiofuranoside, by in‑
vestigating their eects on biomarkers associated with muscle cell apoptosis. Additionally, protein
and gene expression in a muscle atrophy model were examined using Western bloing and RT‑PCR.
Ulmus macrocarpa has been used as food or medicine due to its safety, including its roots, barks, and
fruit. Catechin 7‑O‑β‑D apiofuranoside is an indicator substance of plants of the Ulmus genus and
has been reported to have various eects such as antioxidant and anti‑inammatory eects. The ex‑
perimental results demonstrated that catechin glycoside and Ulmus macrocarpa extract decreased the
expression of the muscle‑degradation‑related proteins Atrogin‑1 and Muscle RING‑Finger protein‑1
(MuRF1) while increasing the expression of the muscle‑synthesis‑related proteins Myoblast deter‑
mination (MyoD) and Myogenin. Gene expression conrmation experiments validated a decrease
in the expression of Atrogin and MuRF1 mRNA and an increase in the expression of MyoD and
Myogenin mRNA. Furthermore, an examination of muscle protein expression associated with the
protein kinase B (Akt)/forkhead box O (FoxO) signaling pathway conrmed a decrease in the ex‑
pression of FoxO, a regulator of muscle protein degradation. These results conrm the potential of
Ulmus macrocarpa extract to inhibit muscle apoptosis, prevent muscle decomposition, and promote
the development of functional materials for muscle synthesis, health‑functional foods, and natural‑
product‑derived medicines.
Keywords: Ulmus macrocarpa; Catechin 7‑O‑β‑D apiofuranoside; apoptosis; muscle atrophy;
sarcopenia
1. Introduction
Aging is a signicant factor in demographic changes worldwide. With advances in
medical science leading to increased average lifespans and declining birth rates resulting in
a growing elderly population, there is heightened interest in issues related to aging, such as
anti‑aging and disease prevention [1]. Muscle atrophy, known to be one of the symptoms
Curr. Issues Mol. Biol. 2024,46, 8320–8339. https://doi.org/10.3390/cimb46080491 https://www.mdpi.com/journal/cimb

Curr. Issues Mol. Biol. 2024,46 8321
leading to sarcopenia, involves the loss of muscle mass, weakness, fatigue, and reduced
muscle contraction activity. These conditions contribute to the worsening of chronic dis‑
eases such as chronic heart failure, chronic kidney disease, and cancer. Current treatment
methods for muscle atrophy primarily include exercise and nutritional supplements [2].
However, exercise can be limited for the elderly and bedridden patients. Thus, there is
a pressing need for the development of safe and harmless therapeutic agents to prevent
muscle atrophy.
Ulmus macrocarpa belongs to the Ulmaceae family, and is known in traditional Korean
medicine as Yubaekpi (楡白皮), with the dried root bark known as Yugeunpi (楡根皮). It is
considered non‑toxic and has been traditionally used for conditions such as eczema, arthri‑
tis, and inammation [3]. In folk medicine, it has also been utilized to treat gastrointestinal
disorders, edema, and cancer [4]. Plants of the Ulmus genus contain (‑)‑catechin, triterpene,
and neolignan glycoside, and these are known to exert their eects through their metabo‑
lites after absorption and metabolism. [5–8]. In a recent study conducted by our research
team, various parts (leaves, stems, and bark) of four indigenous Ulmus genus plants were
subjected to phytochemical analyses using HPLC and NMR. Through this analysis, cate‑
chin 7‑O‑β‑D‑apiofuranoside was identied as a standard compound that can be used to
determine the chemotaxonomy of the Ulmus genus [6]. Numerous previous studies have
also conrmed that the characteristic compound of the Ulmus genus is catechin 7‑O‑β‑D‑
apiofuranoside [9,10]. This compound has been reported to exhibit various and remarkable
biological activities, including antioxidant eects [11], anti‑inammatory eects [5], im‑
munomodulatory eects [12], and the regulation of apoptosis in human papilla cells [13].
Therefore, our goal is to develop a therapeutic agent that utilizes natural compounds de‑
rived from Ulmus macrocarpa for the prevention of muscle loss and atrophy.
In this study, we investigated biomarkers related to the inhibition of muscle cell apop‑
tosis in an Ulmus macrocarpa extract and its active component, catechin 7‑O‑β‑D‑
apiofuranoside. Additionally, we employed a dexamethasone‑induced muscle cell model
to verify their anti‑atrophy eects and performed a detailed validation of the protein and
mRNA expression of muscle‑synthesis‑ and atrophy‑related biomarkers, such as atrogin1,
MuRF1, Myogenin, and MyoD. Our research team conducted an in vitro mechanism study
that covered four distinct categories—inhibiting oxidative stress, anti‑apoptosis eects,
promoting muscle protein synthesis, and inhibiting muscle protein degradation—to ex‑
plore the mechanisms through which Ulmus macrocarpa prevents muscle loss and atrophy
and its eectiveness.
2. Materials and Methods
2.1. Plant Extract Materials
This study was conducted using an Ulmus macrocarpa extract and isolated and puri‑
ed catechin 7‑O‑β‑D apiofuranoside from the extract. The Ulmus macrocarpa bark used in
this study was purchased from Seoul Yakryeong Market and was used after it was certi‑
ed by Professor Choi (Department of Forest Biomaterials Engineering, Kangwon National
University). The bark was cleaned and washed to remove impurities and used as an ex‑
perimental material. A sample of the Ulmus macrocarpa extract (UME‑2023‑04) is stored at
the Department of Forest Biomaterials Engineering, Kangwon National University.
2.2. Pilot‑Scale Extraction and Solvent Fractionation of Bark of Ulmus macrocarpa
In order to determine the optimal yield conditions and the standard component con‑
tent of Ulmus macrocarpa at the lab scale stage, Ulmus macrocarpa was extracted at dierent
edible ethanol ratios (25%, 50%, and 100%) and extraction times (4 h and 8 h). In the semi‑
pilot scale stage, dextrin, a forming agent, was added at concentrations of 0, 5, 10, 25, and
50% to the Ulmus macrocarpa extract. Subsequently, in the pilot scale stage, 300 kg of Ul‑
mus macrocarpa raw material was concentrated under a 50% edible ethanol concentration
and 80 ◦C, resulting in the acquisition of 60 kg of Ulmus macrocarpa concentrate (yield 20%,
Lot. No. DJTT‑22969). From this, 31 kg of Ulmus macrocarpa concentrate was freeze‑dried,

Curr. Issues Mol. Biol. 2024,46 8322
recovering 14.105 kg of powder (yield 45.5%). To this powder, 10% dextrin (1.411 kg) was
added, resulting in the obtainment of 15.516 kg of Ulmus macrocarpa extract powder (UME,
Lot. No. DJTT‑06789). The extraction process was conducted by DanjoungBio Co., Ltd.
(Wonju, Republic of Korea).
2.3. Separation and Purication of Catechin Glycoside
The purication and isolation of catechin 7‑O‑β‑D‑apiofuranoside (CAG) were per‑
formed using the method detailed in Kwon et al., 2022 [13] (Figure 1).
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 3
80 °C, resulting in the acquisition of 60 kg of Ulmus macrocarpa concentrate (yield 20%, Lot.
No. DJTT-22969). From this, 31 kg of Ulmus macrocarpa concentrate was freeze-dried, re-
covering 14.105 kg of powder (yield 45.5%). To this powder, 10% dextrin (1.411 kg) was
added, resulting in the obtainment of 15.516 kg of Ulmus macrocarpa extract powder (UME,
Lot. No. DJTT-06789). The extraction process was conducted by DanjoungBio Co., Ltd.
(Wonju, Republic of Korea).
2.3. Separation and Purification of Catechin Glycoside
The purification and isolation of catechin 7-O-β-D-apiofuranoside (CAG) were per-
formed using the method detailed in Kwon et al., 2022 [13] (Figure 1).
Figure 1. The structure of catechin 7-O-β-D apiofuranoside.
Catechin 7-O-β-D-apiofuranoside (1) Brown amorphous powder, Negative LC-MS/MS:
m/z 422.38 [M − H]−
1H-NMR (400 MHz, MeOH-d4): 6.82 (H-2′, 1H, d, J = 1.2 Hz), 6.76 (H-5′, 1H, d, J = 7.6 z),
6.72 (H-6′, 1H, dd J = 2, 8 Hz), 6.12 (H-8, 1H, d, J = 2.4 Hz), 6.07 (H-6, 1H, d, J = 2.4 Hz), 5.47
(H-1″, 1H, d, J = 3.2 Hz), 4.58 (H-2, 1H, d, J = 7.6 Hz), 4.13 (H-2″, 1H, d, J = 3.2 Hz), 4.06 (H-
4a″, 1H, d, J = 10 Hz), 3.99 (H-3, 1H, m), 3.84 (H-4b″, 1H, d, J = 10 Hz), 3.60 (H-5″, 2H, m),
2.82 (H-4a, 1H, dd, J = 5.6, 16.8 Hz), and 2.56 (H-4b, 1H, dd, J = 8, 16.4 Hz) [6].
13C-NMR (100 MHz, MeOH-d4): 158.3 (C-7), 157.7 (C-5), 157.0 (C-9), 146.4 (C-4′), 146.4 (C-
3′), 132.3 (C-1′), 120.1 (C-6′), 116.2 (C-5′), 115.3 (C-2′), 108.8 (C-1″), 103.4 (C-10), 97.0 (C-8),
97.4 (C-6), 83.1 (C-2), 80.4 (C-3″), 78.4 (C-2″), 75.6 (C-4″), 68.8 (C-3), 65.1 (C-5″), and 28.6
(C-4) [6].
2.4. Quantitative Chromatographic Analysis of Ulmus macrocarpa Extract Powder (UME)
An HPLC analysis of Ulmus macrocarpa extract powder (UME) was performed using
a Waters 2695 (Waters, Milford, MA, USA). Its column conditions were used in combina-
tion with a SkyPak C18 column (5 µm) and Phenomenex KJ0-4282 guard column. The
injected volume was 20 µL and the wavelength was 280 nm. The analysis was conducted
at a flow rate of 1 mL/min for 35 min using 0.9% acetic acid in water and acetonitrile as
the mobile phases. The relationship between the concentration and the peak area was
measured using the minimum square method (R2 value). A standard calibration curve and
the equation of that curve were obtained from six concentrations of catechin 7-O-β-D api-
ofuranoside (CAG) (Figure 2). This resulted in Y = 9982x + 68854 (R2 = 0.999) increments
of CAG, as shown in Figure 2. The calibration curve has good linearity (correlation coeffi-
cient ≥ 0.999). A chromatogram of CAG is shown in Figure 3, and the average content of
the UME was calculated as 133.36 ± 0.27 µg/mL using the formula above (Figure 4).
Figure 1. The structure of catechin 7‑O‑β‑D apiofuranoside.
Catechin 7‑O‑β‑D‑apiofuranoside (1) Brown amorphous powder, Negative
LC‑MS/MS: m/z422.38 [M −H]−
1H‑NMR (400 MHz, MeOH‑d4): 6.82 (H‑2′, 1H, d, J = 1.2 Hz), 6.76 (H‑5′, 1H, d, J = 7.6 z),
6.72 (H‑6′, 1H, dd J= 2, 8 Hz), 6.12 (H‑8, 1H, d, J= 2.4 Hz), 6.07 (H‑6, 1H, d, J= 2.4 Hz), 5.47
(H‑1″, 1H, d, J= 3.2 Hz), 4.58 (H‑2, 1H, d, J= 7.6 Hz), 4.13 (H‑2″, 1H, d, J= 3.2 Hz), 4.06
(H‑4a″, 1H, d, J= 10 Hz), 3.99 (H‑3, 1H, m), 3.84 (H‑4b″, 1H, d, J= 10 Hz), 3.60 (H‑5″, 2H,
m), 2.82 (H‑4a, 1H, dd, J= 5.6, 16.8 Hz), and 2.56 (H‑4b, 1H, dd, J= 8, 16.4 Hz) [6].
13C‑NMR (100 MHz, MeOH‑d4): 158.3 (C‑7), 157.7 (C‑5), 157.0 (C‑9), 146.4 (C‑4′), 146.4 (C‑
3′), 132.3 (C‑1′), 120.1 (C‑6′), 116.2 (C‑5′), 115.3 (C‑2′), 108.8 (C‑1″), 103.4 (C‑10), 97.0 (C‑8),
97.4 (C‑6), 83.1 (C‑2), 80.4 (C‑3″), 78.4 (C‑2″), 75.6 (C‑4″), 68.8 (C‑3), 65.1 (C‑5″), and 28.6
(C‑4) [6].
2.4. Quantitative Chromatographic Analysis of Ulmus macrocarpa Extract Powder (UME)
An HPLC analysis of Ulmus macrocarpa extract powder (UME) was performed using a
Waters 2695 (Waters, Milford, MA, USA). Its column conditions were used in combination
with a SkyPak C18 column (5 µm) and Phenomenex KJ0‑4282 guard column. The injected
volume was 20 µL and the wavelength was 280 nm. The analysis was conducted at a ow
rate of 1 mL/min for 35 min using 0.9% acetic acid in water and acetonitrile as the mobile
phases. The relationship between the concentration and the peak area was measured using
the minimum square method (R2value). A standard calibration curve and the equation
of that curve were obtained from six concentrations of catechin 7‑O‑β‑D apiofuranoside
(CAG) (Figure 2). This resulted in Y = 9982x + 68,854 (R2= 0.999) increments of CAG, as
shown in Figure 2. The calibration curve has good linearity (correlation coecient ≥0.999).
A chromatogram of CAG is shown in Figure 3, and the average content of the UME was
calculated as 133.36 ±0.27 µg/mL using the formula above (Figure 4).

Curr. Issues Mol. Biol. 2024,46 8323
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 4
Figure 2. Calibration curve and equation of catechin 7-O-β-D apiofuranoside (1000 µg/mL, 500
µg/mL, 250 µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL).
Figure 3. HPLC chromatogram of catechin 7-O-β-D apiofuranoside (1000 µg/mL, 500 µg/mL, 250
µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL of CAG).
Figure 4. HPLC chromatogram of Ulmus macrocarpa extract powder (1000 µg/mL).
2.5. Cell Culture and Treatments
C2C12 cells, myoblasts derived from mouse skeletal muscle, were purchased from
the American Type Culture Collection (ATCC). C2C12 cells were cultured in Dulbecco’s
Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100
units/mL of penicillin, and 100 µg/mL streptomycin and incubated at 37 °C in a humidi-
fied CO
2
incubator (5% CO
2
/95% air).
2.5.1. Apoptosis Induction and Treatment
Figure 2. Calibration curve and equation of catechin 7‑O‑β‑D apiofuranoside (1000 µg/mL,
500 µg/mL, 250 µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL).
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 4
Figure 2. Calibration curve and equation of catechin 7-O-β-D apiofuranoside (1000 µg/mL, 500
µg/mL, 250 µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL).
Figure 3. HPLC chromatogram of catechin 7-O-β-D apiofuranoside (1000 µg/mL, 500 µg/mL, 250
µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL of CAG).
Figure 4. HPLC chromatogram of Ulmus macrocarpa extract powder (1000 µg/mL).
2.5. Cell Culture and Treatments
C2C12 cells, myoblasts derived from mouse skeletal muscle, were purchased from
the American Type Culture Collection (ATCC). C2C12 cells were cultured in Dulbecco’s
Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100
units/mL of penicillin, and 100 µg/mL streptomycin and incubated at 37 °C in a humidi-
fied CO
2
incubator (5% CO
2
/95% air).
2.5.1. Apoptosis Induction and Treatment
Figure 3. HPLC chromatogram of catechin 7‑O‑β‑D apiofuranoside (1000 µg/mL, 500 µg/mL,
250 µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL of CAG).
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 4
Figure 2. Calibration curve and equation of catechin 7-O-β-D apiofuranoside (1000 µg/mL, 500
µg/mL, 250 µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL).
Figure 3. HPLC chromatogram of catechin 7-O-β-D apiofuranoside (1000 µg/mL, 500 µg/mL, 250
µg/mL, 125 µg/mL, 62.5 µg/mL, and 31.25 µg/mL of CAG).
Figure 4. HPLC chromatogram of Ulmus macrocarpa extract powder (1000 µg/mL).
2.5. Cell Culture and Treatments
C2C12 cells, myoblasts derived from mouse skeletal muscle, were purchased from
the American Type Culture Collection (ATCC). C2C12 cells were cultured in Dulbecco’s
Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100
units/mL of penicillin, and 100 µg/mL streptomycin and incubated at 37 °C in a humidi-
fied CO
2
incubator (5% CO
2
/95% air).
2.5.1. Apoptosis Induction and Treatment
Figure 4. HPLC chromatogram of Ulmus macrocarpa extract powder (1000 µg/mL).
2.5. Cell Culture and Treatments
C2C12 cells, myoblasts derived from mouse skeletal muscle, were purchased from the
American Type Culture Collection (ATCC). C2C12 cells were cultured in Dulbecco’s Modi‑
ed Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS),
100 units/mL of penicillin, and 100 µg/mL streptomycin and incubated at 37 ◦C in a hu‑
midied CO2incubator (5% CO2/95% air).
2.5.1. Apoptosis Induction and Treatment
C2C12 myoblasts were exposed to 100 µM of H2O2to induce apoptosis. To investigate
the eect of UME and CAG on H2O2‑induced apoptosis in C2C12 myoblasts, cells were

Curr. Issues Mol. Biol. 2024,46 8324
plated in multi‑well plates and incubated for 24 h. Afterwards, C2C12 myoblasts were
treated with 100 µM of H2O2with/without dierent concentrations of UME or CAG, as
indicated [14].
2.5.2. Dierentiation Induction and Treatment
To determine the eect of UME and CAG on cell viability in myotubes in the presence
of dexamethasone (Sigma‑Aldrich, Darmstadt, Germany), C2C12 cells were plated in 24‑
well plates at a density of 5 ×104cells/well and dierentiated into myotubes for 4 days,
as described above. After the induction of dierentiation, the myotubes were treated with
5µM of dexamethasone and various concentrations of UME or CAG for 24 h.
2.6. Eects of Ulmus Macrocarpa Extract Powder (UME) and Catechin 7‑O‑β‑D‑apiofuranoside
(CAG) on Muscle Apoptosis Biomarkers
2.6.1. Evaluation of Apoptosis in H2O2‑Induced Myoblasts
To evaluate the eects of the test substances on H2O2‑induced apoptosis in myoblasts,
C2C12 cells were seeded into 24‑well plates at a density of 5 ×104cells/well and cultured
for 24 h. After the initial 24 h culture period, the cells were treated with 100 µM of H2O2
to induce myoblast damage. To investigate the protective eects of the test substances
against myoblast damage, ve dierent test substances were administered at various con‑
centrations in conjunction with 100 µM of H2O2, and the cells were cultured for an ad‑
ditional 48 h. The extent of the apoptosis in myoblasts was measured using a Cellular
DNA Fragmentation ELISA kit (Sigma‑Aldrich), which detects 5′‑Bromo‑2′‑deoxy‑uridine
(BrdU)‑labeled DNA, following the manufacturer’s protocol.
2.6.2. Western Blot Analysis
C2C12 myoblasts were plated and treated with UME or CAG in the presence of 100 µM
of H2O2for 24 h, as described above. After that, the cells were lysed and Western blot anal‑
yses were conducted as described previously [15]. Antibodies against Bax, Bcl‑2, cleaved
caspase‑3, cleaved PARP, and β‑actin (Cell Signaling Technology, Beverly, MA, USA) were
used in this analysis. The protein bands were developed using the LuminataTM Forte West‑
ern HRP Substrate (Millipore, Billerica, MA, USA) and captured and analyzed for their
intensity using an ImageQuantTM LAS 500 imaging system (GE Healthcare Bio‑Sciences
AB, Uppsala, Sweden). Their relative protein expressions were normalized to β‑actin.
2.7. Eects of Ulmus Macrocarpa Extract (UME) and Catechin 7‑O‑β‑D apiofuranoside (CAG)
on Muscle‑Synthesis‑ and Muscle‑Degradation‑Related Proteins and Gene Expression
2.7.1. Measurement of Myotube Diameter
C2C12 cells were plated in 24‑well plates at a density of 5 ×104cells/well, dierenti‑
ated into myotubes for 4 days, and treated with UME or CAG and 5 µM of dexamethasone
for 24 h, as described above. After that, the cells were xed with 4% paraformaldehyde,
permeabilized with 0.1% Triton X‑100, and blocked with 5% BSA. The cells were immunos‑
tained with MYH7 antibody and Alexa Fluor 594 labeled goat anti‑mouse IgG antibody.
DAPI (Sigma‑Aldrich) was applied as a counterstain for the nuclei. Fluorescent cell im‑
ages (6 images per group) were captured using microscopy (AxioImager, Carl Zeiss, Jena,
Germany) at 20×magnication. Ten myotubes per image were chosen on a random basis
from each micrograph. The thickest portion of each myotube was analyzed for its my‑
otube diameter using ImageJ software (National Institutes of Health, Bethesda, MD, USA,
Version 1.54).
2.7.2. Real‑Time Reverse Transcription Polymerase Chain Reaction (RT‑PCR) Analysis
C2C12 cells were plated, dierentiated into myotubes for 4 days, and treated with
UME or CAG and 5 µM of dexamethasone for 24 h, as described above. Total RNA was ex‑
tracted using an RNeasy®Plus Mini kit (Qiagen, Valencia, CA, USA) according to the man‑
ufacturer’s instructions. The extracted total RNA was quantied using a micro‑volume

Curr. Issues Mol. Biol. 2024,46 8325
spectrophotometer (BioSpec‑nano, Shimadzu, Kyoto, Japan). Reverse transcription from
2µg of total RNA was performed using a HyperScriptTM RT master mix kit (GeneAll
Biotechnology). A real‑time PCR was performed using a QuantiNova SYBR Green PCR
kit (Qiagen) on a Rotor‑gene 300 PCR (Corbe Research, Mortlake, Australia), as described
previously [16]. The sequences of the primers used in this PCR are shown in Table 1. Data
analysis was conducted using the Rotor‑Gene 6000 series System Soft program version 6
(Corbe). The relative expression levels of the target mRNA were normalized to that of
the housekeeping protein glyceraldehyde 3‑phosphate dehydrogenase (GAPDH) [10].
Table 1. Specic primer sequences for PCR.
mRNA Primer Sequences
Atrogin‑1 Forward 5′‑GCCCTCCACACTAGTTGACC‑3′
Reverse 5′‑GACGGATTGACAGCCAGGAA‑3′
MuRf‑1 Forward 5′‑GAGGGCCATTGACTTTGGGA‑3′
Reverse 5′‑TTTACCCTCTGTGGTCACGC‑3′
MyoD1 Forward 5′‑GCACTACAGTGGCGACTCAGAT‑3′
Reverse 5′‑TAGTAGGCGGTGTCGTAGCCAT‑3′
Myogenin Forward 5′‑CCATCCAGTACATTGAGCGCCT‑3′
Reverse 5′‑CTGTGGGAGTTGCATTCACTGG‑3′
GAPDH Forward 5′‑TGGGTGTGAACCATGAGAAG‑3′
Reverse 5′‑GCTAAGCAGTTGGTGGTGC‑3′
2.7.3. Western Blot Analysis
C2C12 myoblasts were plated, dierentiated into myotubes for 4 days, and treated
with UME and 5 µM of dexamethasone for 24 h, as described above. After that, the cells
were lysed and Western blot analyses were conducted as described previously [16]. Anti‑
bodies against Atrogin‑1, MuRF‑1 (Santa Cruz, Santa Cruz, CA, USA), MyoD1, Myogenin
(Abcam, Cambridge, MA, USA), phospho‑Akt (Ser473), Akt, phospho‑mTOR (Ser253),
mTOR, phospho‑FoxO1 (Ser256), FoxO1, phospho‑FoxO3 (Ser253), FoxO3, and β‑actin
(Cell Signaling Technology) were used in this analysis. The protein bands were developed
using the LuminataTM Forte Western HRP Substrate (Millipore, Billerica, MA, USA) and
then captured and analyzed for their intensity using the ImageQuantTM LAS 500 imaging
system (GE Healthcare Bio‑Sciences AB, Uppsala, Sweden). Their relative protein expres‑
sions were normalized to β‑actin.
2.8. Statistical Analysis
All values from the analyses are expressed as mean ±S.E.M. The collected results
were analyzed using the GraphPad Prism 5.0 (GraphPad Software, San Diego, CA, USA)
program. Student’s t‑test and a one‑way analysis of variance (ANOVA) were used to
compare the dierences between the treatment group and the control group. These were
judged to be statistically signicant only when p< 0.05.
3. Results
3.1. The Impact of UME and CAG on the Viability of C2C12 Cells
Measurement of Cell Viability under Normal Conditions
To investigate cytotoxicity in C2C12 cells, dierent concentrations of UME and CAG
were added to the cell culture medium and cultured for 48 h, and then an MTT assay was
performed. Treatment with UME (Figure 5A) resulted in a decrease in cell viability at
concentrations of 400 µg/mL and above, while treatment with CAG (Figure 5B) did not
decrease cell viability to 80% or below at any treatment concentration (Figure 5). Based on
these results, the highest treatment concentrations for UME and CAG were set to 200 µg/mL
and 100 µg/mL, respectively, for further experiments.

Curr. Issues Mol. Biol. 2024,46 8326
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 7
To investigate cytotoxicity in C2C12 cells, different concentrations of UME and CAG
were added to the cell culture medium and cultured for 48 h, and then an MTT assay was
performed. Treatment with UME (Figure 5A) resulted in a decrease in cell viability at con-
centrations of 400 µg/mL and above, while treatment with CAG (Figure 5B) did not de-
crease cell viability to 80% or below at any treatment concentration (Figure 5). Based on
these results, the highest treatment concentrations for UME and CAG were set to 200
µg/mL and 100 µg/mL, respectively, for further experiments.
Figure 5. Cytotoxicity of (A) Ulmus macrocarpa extract (UME) and (B) catechin 7-O-β-D apio-
furanoside (CAG) on C2C12 cells. Cell viability was calculated as described in the Materials and
Methods. Values are expressed as mean ± S.E.M. (n = 3). * p < 0.05, *** p < 0.001 are significantly
different from that of the 0 µg/mL group.
3.2. Effects of Ulmus Macrocarpa Extract (UME) and Catechin 7-O-β-D Apiofuranoside (CAG)
on Muscle Apoptosis Biomarkers
3.2.1. Effects on H
2
O
2
-Induced Apoptosis in Myoblasts
In models of muscle atrophy caused by diabetes, cancer, heart failure, AIDS, and sep-
sis, skeletal muscle atrophy, which is associated with oxidative stress, is induced [17–20].
Oxidative stress leads to the generation of reactive oxygen species, activating the ubiquitin
pathway. This results in increased protein degradation and decreased myosin expression,
and ultimately leads to muscle atrophy [21–23]. To investigate the protective effect of UME
and CAG against muscle cell damage, oxidative stress was induced in C2C12 cells using
H
2
O
2
and each substance was administered to determine cell viability (Figure 6) [24,25].
When normal cells were treated with 100 µM of H
2
O
2
, the cell viability rate decreased to
34.1 ± 0.9%, and, when treated with UME (Figure 6A), it increased in a concentration-de-
pendent manner, reaching 52.6 ± 1.6% at the highest treatment concentration of 100
µg/mL, about an 18.5% increase. When apoptosis was induced using H
2
O
2
, the concentra-
tion-dependent cell viability was significantly decreased when treated with CAG (Figure
6B), increasing by 8.3% at the highest treatment concentration of 100 µg/mL. Accordingly,
the protective effects of UME and CAG against H
2
O
2
-induced myoblast cell damage were
confirmed.
Figure 5. Cytotoxicity of (A)Ulmus macrocarpa extract (UME) and (B) catechin 7‑O‑β‑D apiofura‑
noside (CAG) on C2C12 cells. Cell viability was calculated as described in the Materials and Meth‑
ods. Values are expressed as mean ±S.E.M. (n = 3). * p< 0.05, *** p< 0.001 are signicantly dierent
from that of the 0 µg/mL group.
3.2. Eects of Ulmus Macrocarpa Extract (UME) and Catechin 7‑O‑β‑D Apiofuranoside (CAG)
on Muscle Apoptosis Biomarkers
3.2.1. Eects on H2O2‑Induced Apoptosis in Myoblasts
In models of muscle atrophy caused by diabetes, cancer, heart failure, AIDS, and sep‑
sis, skeletal muscle atrophy, which is associated with oxidative stress, is induced [17–20].
Oxidative stress leads to the generation of reactive oxygen species, activating the ubiquitin
pathway. This results in increased protein degradation and decreased myosin expression,
and ultimately leads to muscle atrophy [21–23]. To investigate the protective eect of UME
and CAG against muscle cell damage, oxidative stress was induced in C2C12 cells using
H2O2and each substance was administered to determine cell viability
(Figure 6) [24,25]. When normal cells were treated with 100 µM of H2O2, the cell viabil‑
ity rate decreased to 34.1 ±0.9%, and, when treated with UME (Figure 6A), it increased in
a concentration‑dependent manner, reaching 52.6 ±1.6% at the highest treatment concen‑
tration of 100 µg/mL, about an 18.5% increase. When apoptosis was induced using H2O2,
the concentration‑dependent cell viability was signicantly decreased when treated with
CAG (Figure 6B), increasing by 8.3% at the highest treatment concentration of 100 µg/mL.
Accordingly, the protective eects of UME and CAG against H2O2‑induced myoblast cell
damage were conrmed.
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 8
Figure 6. Protective effect of (A) Ulmus macrocarpa extract (UME) and (B) catechin 7-O-β-D apio-
furanoside (CAG) on cell viability in H
2
O
2
-treated C2C12 myoblasts. Values are expressed as mean
± S.E.M. (n = 3). *** p < 0.001 are significantly different from that of the [H
2
O
2
(−)/UME(−)],
[H
2
O
2
(−)/CAG(−)] group.
#
p < 0.05,
##
p < 0.01, and
###
p < 0.001 are significantly different from that of
the [(+)/(−)] group.
3.2.2. Effects of Catechin 7-O-β-D apiofuranoside on Bax, Bcl-2, Cleaved Caspase-3, and
Cleaved PARP Protein Expression
Apoptosis is essential for normal growth and homeostasis in all multicellular organ-
isms and it occurs via two pathways: the intrinsic pathway, via the mitochondria, and the
extrinsic pathway. In both pathways, the activation of a family of sequential cysteine pro-
teases, the caspase enzymes, occurs concomitantly [26,27] and continuously [28]. In re-
search to date, the mechanisms through which mitochondria work in the apoptosis path-
way can be broadly divided into two types. First, there are mechanisms such as the Bcl-2
family [29], which are muscle proteins that control the apoptosis pathway and protect
cells, and Bax [30] and Apaf-1 [31], which are genes that induce apoptosis. Bcl-2 interrupts
the release of cytochrome C from the mitochondria or binds to muscle proteins around
the mitochondria, preventing apoptosis [27]. On the other hand, Bax facilitates the release
of cytochrome C in response to stress such as DNA damage, triggering apoptosis [32].
Second, they allow for the activation of caspases. Caspases, a crucial type of protease in-
volved in various cellular functions, including differentiation and apoptosis, are activated
by cytochrome C being released into the cytoplasm. This activates caspases such as
caspase-8, caspase-9, and caspase-2, leading to the activation of downstream caspases like
caspase-3 and caspase-6. These caspases cleave cellular muscle proteins, inducing apop-
tosis [33]. Of these, the activation of caspase-3 is particularly crucial as it occurs just before
apoptosis [34].
We investigated the effect of CAG on apoptosis in H2O2-induced C2C12 cells (Figure
7). The results showed that the expression of Bax (Figure 7A,B) was not significantly dif-
ferent between the group without an H2O2 treatment and groups treated with varying
concentrations of CAG after H2O2 exposure. However, the expression of Bcl-2 (Figure
7C,D) significantly decreased when treated with H2O2, and particularly at a concentration
of 50 µg/mL, where there was a significant increase in Bcl-2 protein expression to 1.29 ±
0.04.
Figure 6. Protective eect of (A)Ulmus macrocarpa extract (UME) and (B) catechin 7‑O‑β‑D api‑
ofuranoside (CAG) on cell viability in H2O2‑treated C2C12 myoblasts. Values are expressed as
mean ±S.E.M. (n = 3). *** p< 0.001 are signicantly dierent from that of the [H2O2(−)/UME(−)],
[H2O2(−)/CAG(−)] group. #p< 0.05, ## p< 0.01, and ### p< 0.001 are signicantly dierent from that
of the [(+)/(−)] group.

Curr. Issues Mol. Biol. 2024,46 8327
3.2.2. Eects of Catechin 7‑O‑β‑D apiofuranoside on Bax, Bcl‑2, Cleaved Caspase‑3, and
Cleaved PARP Protein Expression
Apoptosis is essential for normal growth and homeostasis in all multicellular organ‑
isms and it occurs via two pathways: the intrinsic pathway, via the mitochondria, and
the extrinsic pathway. In both pathways, the activation of a family of sequential cysteine
proteases, the caspase enzymes, occurs concomitantly [26,27] and continuously [28]. In
research to date, the mechanisms through which mitochondria work in the apoptosis path‑
way can be broadly divided into two types. First, there are mechanisms such as the Bcl‑2
family [29], which are muscle proteins that control the apoptosis pathway and protect cells,
and Bax [30] and Apaf‑1 [31], which are genes that induce apoptosis. Bcl‑2 interrupts the
release of cytochrome C from the mitochondria or binds to muscle proteins around the
mitochondria, preventing apoptosis [27]. On the other hand, Bax facilitates the release of
cytochrome C in response to stress such as DNA damage, triggering apoptosis [32]. Sec‑
ond, they allow for the activation of caspases. Caspases, a crucial type of protease involved
in various cellular functions, including dierentiation and apoptosis, are activated by cy‑
tochrome C being released into the cytoplasm. This activates caspases such as caspase‑8,
caspase‑9, and caspase‑2, leading to the activation of downstream caspases like caspase‑
3 and caspase‑6. These caspases cleave cellular muscle proteins, inducing apoptosis [33].
Of these, the activation of caspase‑3 is particularly crucial as it occurs just before apopto‑
sis [34].
We investigated the eect of CAG on apoptosis in H2O2‑induced C2C12 cells
(Figure 7). The results showed that the expression of Bax (Figure 7A,B) was not signi‑
cantly dierent between the group without an H2O2treatment and groups treated with
varying concentrations of CAG after H2O2exposure. However, the expression of Bcl‑2
(Figure 7C,D) signicantly decreased when treated with H2O2, and particularly at a con‑
centration of 50 µg/mL, where there was a signicant increase in Bcl‑2 protein expression
to 1.29 ±0.04.
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 9
Figure 7. Anti-apoptotic effects of catechin 7-O-β-D apiofuranoside (CAG) on H
2
O
2
-induced oxida-
tive damage in C2C12. Western bloing was performed to analyze levels of (A,B) Bax, (C,D) Bcl-2,
(E,F) cleaved caspase-3, (G,H) cleaved PARP, and actin, as described in Materials and Methods.
Values are expressed as mean ± S.E.M. (n = 3). * p < 0.05, *** p < 0.001 are significantly different from
that of the untreated H
2
O
2
group.
#
p < 0.05 are significantly different from that of the H
2
O
2
-treated
group.
In the H
2
O
2
-treated control group, the expression of cleaved caspase-3 (Figure 7E,F)
significantly increased. However, when treated with CAG at concentrations of 10 µg/mL
and 50 µg/mL, its expression decreased significantly to 0.77 ± 0.05 and 0.69 ± 0.07, respec-
tively. Similarly, the expression of cleaved PARP (Figure 7G,H) significantly increased in
the H
2
O
2
-treated control group. However, when treated with 10–50 µg/mL of CAG, the
expression of cleaved PARP decreased significantly at all concentrations, to 0.61 ± 0.08,
0.60 ± 0.07, and 0.69 ± 0.06, respectively. This demonstrates that CAG effectively has an
inhibitory effect on muscle apoptosis.
3.3. Effects of UME and CAG on Muscle-Synthesis- and Muscle-Degradation-Related Proteins
and Gene Expression
3.3.1. Impact of Dexamethasone-Induced Myotube Damage
Dexamethasone (DEX) is a representative glucocorticoid that causes the degradation
of skeletal muscle and, based on this, it is widely used to induce muscle cell atrophy in in
vitro systems [35–38]. Previous studies have reported that dexamethasone increases pro-
tein degradation through the ubiquitin–proteasome pathway and regulates the expression
of related genes [39–41].
To investigate the effects of UME and CAG on myocyte damage, C2C12 cells were
treated with dexamethasone to induce myocyte atrophy, and the viability rate of the my-
otube cells was then confirmed via treatments with each substance. When the C2C12 cells
were treated with dexamethasone (Figure 8), their cell viability decreased to 89.3 ± 0.7%,
while treatments with UME (Figure 8A) or CAG (Figure 8B) led to a concentration-de-
pendent increase in cell viability. In particular, treatment with the highest concentration
of UME (200 µg/mL) resulted in an increase in cell viability to 94.5 ± 1.0%, while CAG led
Figure 7. Anti‑apoptotic eects of catechin 7‑O‑β‑D apiofuranoside (CAG) on H2O2‑induced ox‑
idative damage in C2C12. Western bloing was performed to analyze levels of (A,B) Bax, (C,D)
Bcl‑2, (E,F) cleaved caspase‑3, (G,H) cleaved PARP, and actin, as described in Materials and Meth‑
ods. Values are expressed as mean ±S.E.M. (n = 3). * p< 0.05, *** p< 0.001 are signicantly dierent
from that of the untreated H2O2group. #p< 0.05 are signicantly dierent from that of the H2O2‑
treated group.

Curr. Issues Mol. Biol. 2024,46 8328
In the H2O2‑treated control group, the expression of cleaved caspase‑3 (Figure 7E,F)
signicantly increased. However, when treated with CAG at concentrations of 10 µg/mL
and 50 µg/mL, its expression decreased signicantly to 0.77 ±0.05 and 0.69 ±0.07, respec‑
tively. Similarly, the expression of cleaved PARP (Figure 7G,H) signicantly increased in
the H2O2‑treated control group. However, when treated with 10–50 µg/mL of CAG, the
expression of cleaved PARP decreased signicantly at all concentrations, to 0.61 ±0.08,
0.60 ±0.07, and 0.69 ±0.06, respectively. This demonstrates that CAG eectively has an
inhibitory eect on muscle apoptosis.
3.3. Eects of UME and CAG on Muscle‑Synthesis‑ and Muscle‑Degradation‑Related Proteins
and Gene Expression
3.3.1. Impact of Dexamethasone‑Induced Myotube Damage
Dexamethasone (DEX) is a representative glucocorticoid that causes the degradation
of skeletal muscle and, based on this, it is widely used to induce muscle cell atrophy in
in vitro systems [35–38]. Previous studies have reported that dexamethasone increases
protein degradation through the ubiquitin–proteasome pathway and regulates the expres‑
sion of related genes [39–41].
To investigate the eects of UME and CAG on myocyte damage, C2C12 cells were
treated with dexamethasone to induce myocyte atrophy, and the viability rate of the my‑
otube cells was then conrmed via treatments with each substance. When the C2C12 cells
were treated with dexamethasone (Figure 8), their cell viability decreased to 89.3 ±0.7%,
while treatments with UME (Figure 8A) or CAG (Figure 8B) led to a concentration‑
dependent increase in cell viability. In particular, treatment with the highest concentra‑
tion of UME (200 µg/mL) resulted in an increase in cell viability to 94.5 ±1.0%, while CAG
led to an increase in cell viability at concentrations of 50 µg/mL or higher. This conrmed
that UME and CAG have a protective eect on dexamethasone‑induced myotube damage.
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 10
to an increase in cell viability at concentrations of 50 µg/mL or higher. This confirmed that
UME and CAG have a protective effect on dexamethasone-induced myotube damage.
Figure 8. Protective effect of (A) Ulmus macrocarpa extract (UME) and (B) catechin 7-O-β-D apio-
furanoside (CAG) on cell viability in DEX-treated C2C12 myotubes. Values are expressed as mean
± S.E.M. (n = 3). * p < 0.05, ** p < 0.01 are significantly different from that of the 0 µg/mL group.
##
p <
0.01,
###
p < 0.001 are significantly different from that of the [(+)/(−)] group.
3.3.2. Effect of UME and CAG on Dexamethasone-Induced Myotube Atrophy
To investigate the effects of UME and CAG on the dexamethasone-induced atrophy
of myotubes, the cells were cultured with UME and CAG treatments, and then the diam-
eter of the myotube cells was measured (Figure 9). When treated with 5 µM of dexame-
thasone, the diameter of the myocytes was significantly reduced; their diameter decreased
to 0.09~0.11 µm compared to the diameter of 0.36~0.39 µm observed in the control group.
Subsequently, when 50, 100, and 200 µg/mL of the UME (Figure 9A) were applied as treat-
ments, their diameter increased significantly, to 0.20 ± 0.01, 0.23 ± 0.01, and 0.26 ± 0.01 µm,
respectively, while, at the highest treatment concentration, it increased to 2.74 times that
of the dexamethasone-induced control group (Figure 9C). In addition, when CAG (Figure
9B) was applied, the diameter of the myotubes increased in a concentration-dependent
manner, increasing significantly at concentrations of 50 and 100 µg/mL to 0.25 ± 0.01 and
0.32 ± 0.01 µm, respectively (Figure 9D). At the highest treatment concentration, it in-
creased to 2.86 times that of the dexamethasone-induced control group and was similar to
that of the untreated control group.
Figure 8. Protective eect of (A)Ulmus macrocarpa extract (UME) and (B) catechin 7‑O‑β‑D api‑
ofuranoside (CAG) on cell viability in DEX‑treated C2C12 myotubes. Values are expressed as
mean ±S.E.M. (n = 3). * p< 0.05, ** p< 0.01 are signicantly dierent from that of the 0 µg/mL
group. ## p< 0.01, ### p< 0.001 are signicantly dierent from that of the [(+)/(−)] group.
3.3.2. Eect of UME and CAG on Dexamethasone‑Induced Myotube Atrophy
To investigate the eects of UME and CAG on the dexamethasone‑induced atrophy of
myotubes, the cells were cultured with UME and CAG treatments, and then the diameter
of the myotube cells was measured (Figure 9). When treated with 5 µM of dexametha‑
sone, the diameter of the myocytes was signicantly reduced; their diameter decreased
to 0.09~0.11 µm compared to the diameter of 0.36~0.39 µm observed in the control group.
Subsequently, when 50, 100, and 200 µg/mL of the UME (Figure 9A) were applied as treat‑
ments, their diameter increased signicantly, to 0.20 ±0.01, 0.23 ±0.01, and 0.26 ±0.01 µm,
respectively, while, at the highest treatment concentration, it increased to 2.74 times that of
the dexamethasone‑induced control group (Figure 9C). In addition, when CAG (Figure 9B)
was applied, the diameter of the myotubes increased in a concentration‑dependent man‑

Curr. Issues Mol. Biol. 2024,46 8329
ner, increasing signicantly at concentrations of 50 and 100 µg/mL to 0.25 ±0.01 and
0.32 ±0.01 µm, respectively (Figure 9D). At the highest treatment concentration, it in‑
creased to 2.86 times that of the dexamethasone‑induced control group and was similar
to that of the untreated control group.
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 11
Figure 9. Effects of (A,C) Ulmus macrocarpa extract (UME) and (B,D) catechin 7-O-β-D apio-
furanoside (CAG) on dexamethasone-induced muscle atrophy in C2C12 cells. Values are expressed
as mean ± S.E.M. (n = 3). Scale bar is 100 µm. *** p < 0.001 are significantly different from that of the
[(−)/(−)] group. # p < 0.05, ### p < 0.001 are significantly different from that of the [(+)/(−)] group.
Therefore, it was confirmed that UME and CAG significantly protect against dexa-
methasone-induced myotube atrophy.
3.3.3. Expression of Atrogin-1, MuRF1, Myogenin, and MyoD1 Protein in DEX-Treated
C2C12 Myotubes
MuRF1 and atrogin-1 act as ubiquitin E3 ligases and are expressed by the sub-signal
system of GDF8, which involves smad 2/3. Additionally, they are stimulated by MAPK to
enhance muscle protein degradation, contributing to muscle atrophy at the cellular level.
This process is triggered by various environmental factors, such as increased glucocorti-
coid levels and reduced muscle usage. In summary, mechanisms of action involving Akt,
mTOR, FoxO, etc., are directly associated with skeletal muscle atrophy and the synthesis
and degradation of muscle proteins [42–45].
The expression of Atrogin1 was investigated after UME (Figure 10A,B) and CAG (Fig-
ure 10C,D) treatments; the increased expression of Atrogin1 that was a result of the dexa-
methasone treatment decreased to 2.15 ± 0.07 and 2.21 ± 0.05 at concentrations of 100 and
200 µg/mL of UME, respectively. Treatment with CAG led to a significant decrease in
Atrogin1 to 2.47 ± 0.13 and 2.17 ± 0.10 at concentrations of 50 µg/mL and 100 µg/mL, re-
spectively. Additionally, the expression of MuRF1 increased to 1.96 ± 0.07 with the dexa-
methasone treatment alone, but decreased in a concentration-dependent manner when
subsequently treated with UME (Figure 10E,F), falling to 1.35 ± 0.03 at a concentration of
100 µg/mL. Similarly, treatment with CAG (Figure 10G,H) led to a concentration-depend-
ent decrease in MuRF1 expression, with significant decreases to 2.01 ± 0.07 and 1.52 ± 0.14
at concentrations of 50 µg/mL and 100 µg/mL, respectively.
Figure 9. Eects of (A,C)Ulmus macrocarpa extract (UME) and (B,D) catechin 7‑O‑β‑D apiofura‑
noside (CAG) on dexamethasone‑induced muscle atrophy in C2C12 cells. Values are expressed as
mean ±S.E.M. (n = 3). Scale bar is 100 µm. *** p< 0.001 are signicantly dierent from that of the
[(−)/(−)] group. #p< 0.05, ### p< 0.001 are signicantly dierent from that of the [(+)/(−)] group.
Therefore, it was conrmed that UME and CAG signicantly protect against
dexamethasone‑induced myotube atrophy.
3.3.3. Expression of Atrogin‑1, MuRF1, Myogenin, and MyoD1 Protein in DEX‑Treated
C2C12 Myotubes
MuRF1 and atrogin‑1 act as ubiquitin E3 ligases and are expressed by the sub‑signal
system of GDF8, which involves smad 2/3. Additionally, they are stimulated by MAPK to
enhance muscle protein degradation, contributing to muscle atrophy at the cellular level.
This process is triggered by various environmental factors, such as increased glucocorti‑
coid levels and reduced muscle usage. In summary, mechanisms of action involving Akt,
mTOR, FoxO, etc., are directly associated with skeletal muscle atrophy and the synthesis
and degradation of muscle proteins [42–45].
The expression of Atrogin1 was investigated after UME (Figure 10A,B) and CAG
(Figure 10C,D) treatments; the increased expression of Atrogin1 that was a result of the
dexamethasone treatment decreased to 2.15 ±0.07 and 2.21 ±0.05 at concentrations of 100
and 200 µg/mL of UME, respectively. Treatment with CAG led to a signicant decrease
in Atrogin1 to 2.47 ±0.13 and 2.17 ±0.10 at concentrations of 50 µg/mL and 100 µg/mL,
respectively. Additionally, the expression of MuRF1 increased to 1.96 ±0.07 with the dex‑
amethasone treatment alone, but decreased in a concentration‑dependent manner when
subsequently treated with UME (Figure 10E,F), falling to 1.35 ±0.03 at a concentration of

Curr. Issues Mol. Biol. 2024,46 8330
100 µg/mL. Similarly, treatment with CAG (Figure 10G,H) led to a concentration‑dependent
decrease in MuRF1 expression, with signicant decreases to 2.01 ±0.07 and 1.52 ±0.14 at
concentrations of 50 µg/mL and 100 µg/mL, respectively.
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 12
On the other hand, the expression of Myogenin decreased to 0.84 ± 0.03 with the dex-
amethasone treatment alone but showed a concentration-dependent increase when sub-
sequently treated with UME (Figure 10I,J), rising to 1.10 ± 0.01 at a concentration of 50
µg/mL. Treatment with CAG (Figure 10K,L) also led to a concentration-dependent in-
crease in myogenin expression, particularly to 1.02 ± 0.01 at a concentration of 50 µg/mL,
surpassing that of the untreated control group. The expression of MyoD1 decreased with
the dexamethasone treatment alone, but showed a concentration-dependent increase
when subsequently treated with UME (Figure 10M,N), reaching 0.91 ± 0.02 and 0.89 ± 0.03
at concentrations of 50 and 100 µg/mL, respectively. Similarly, treatment with CAG (Fig-
ure 10O,P) also led to a concentration-dependent increase, reaching 1.11 ± 0.02 and 1.14 ±
0.03 at concentrations of 50 and 100 µg/mL, surpassing that of the untreated control group
(Figure 10). Therefore, it was confirmed that UME and CAG reduce the expression of
Atrogin1 and MuRF1 while promoting the expression of myogenin and MyoD1.
Figure 10. Effects of Ulmus macrocarpa extract (UME) and catechin 7-O-β-D apiofuranoside (CAG)
on protein expression levels of (A–D) Atrogin-1, (E–H) MuRF1, (I–L) Myogenin, and (M–P) MyoD1
Figure 10. Eects of Ulmus macrocarpa extract (UME) and catechin 7‑O‑β‑D apiofuranoside (CAG)
on protein expression levels of (A–D) Atrogin‑1, (E–H) MuRF1, (I–L) Myogenin, and (M–P) MyoD1
in DEX‑treated C2C12 myotubes. UME and catechin 7‑O‑β‑D were added to DEX‑treated C2C12
myotubes and cultured for 24 h. Protein expression levels were determined using Western blot assay.
Values are expressed as mean ±S.E.M. (n = 3). * p< 0.05, ** p< 0.01, and *** p< 0.001 are signicantly
dierent from that of the [(−)/(−)] group. #p< 0.05, ## p< 0.01, and ### p< 0.001 are signicantly
dierent from that of the [(+)/(−)] group.
On the other hand, the expression of Myogenin decreased to 0.84 ±0.03 with the dex‑
amethasone treatment alone but showed a concentration‑dependent increase when subse‑
quently treated with UME (Figure 10I,J), rising to 1.10 ±0.01 at a concentration of 50 µg/mL.
Treatment with CAG (Figure 10K,L) also led to a concentration‑dependent increase in myo‑

Curr. Issues Mol. Biol. 2024,46 8331
genin expression, particularly to 1.02 ±0.01 at a concentration of 50 µg/mL, surpassing that
of the untreated control group. The expression of MyoD1 decreased with the dexametha‑
sone treatment alone, but showed a concentration‑dependent increase when subsequently
treated with UME (Figure 10M,N), reaching 0.91 ±0.02 and 0.89 ±0.03 at concentrations of
50 and 100 µg/mL, respectively. Similarly, treatment with CAG (Figure 10O,P) also led to
a concentration‑dependent increase, reaching 1.11 ±0.02 and 1.14 ±0.03 at concentrations
of 50 and 100 µg/mL, surpassing that of the untreated control group (Figure 10). Therefore,
it was conrmed that UME and CAG reduce the expression of Atrogin1 and MuRF1 while
promoting the expression of myogenin and MyoD1.
3.3.4. Eects on Muscle‑Degradation‑ and Muscle‑Synthesis‑Related Gene Expression
To ensure the accuracy of this study, experiments were conducted on the expression of
genes related to muscle degradation and synthesis. When treated with dexamethasone to
induce muscle atrophy, the expression of the muscle‑degradation‑related genes Atrogin‑1
mRNA and MuRF1 mRNA increased. When treated with 200 µg/mL of UME, the elevated
expression of Atrogin‑1 induced by dexamethasone decreased to 0.40 ±0.04. Additionally,
the expression of MuRF‑1 signicantly decreased at concentrations of 100 and 200 µg/mL to
0.56 ±0.04 and 0.31 ±0.06, respectively. However, the expression of MyoD and Myogenin,
which are genes related to muscle synthesis, decreased when treated with dexamethasone
and increased in a concentration‑dependent manner when treated with UME. MyoD’s ex‑
pression levels increased to 2.02 ±0.26 and 2.26 ±0.29 at UME concentrations of 50 and
100 µg/mL, respectively, while those of myogenin signicantly increased to 1.51 ±0.10
and 1.76 ±0.17 (Table 2).
Table 2. Eect of Ulmus macrocarpa extract on muscle‑degradation‑ and muscle‑synthesis‑related
gene expression.
DEX
(5 µM)
UME
(µg/mL)
mRNA
Atrogin‑1 MuRF‑1 Myo D Myogenin
− − 0.07 ±0.02 0.10 ±0.18 3.24 ±0.27 2.47 ±0.24
+−1.00 ±0.22 ** 1.00 ±0.10 *** 1.00 ±0.11 *** 1.00 ±0.07 ***
+ 50 1.03 ±0.12 0.90 ±0.06 1.47 ±0.21 1.27 ±0.09 #
+ 100 0.60 ±0.04 0.56 ±0.04 ## 2.02 ±0.26 ## 1.51 ±0.10 ##
+ 200 0.40 ±0.04 #0.31 ±0.06 ### 2.26 ±0.29 ## 1.76 ±0.17 ##
DEX: dexamethasone; UME: Ulmus macrocarpa extract. Values are expressed as mean ±S.E.M. (n = 3). The target
mRNA’s expression was normalized to that of GAPDH. ** p< 0.01, *** p< 0.001 are signicantly dierent from
that of the [(−)/(−)] group. #p< 0.05, ## p< 0.01, and ### p< 0.001 are signicantly dierent from that of the
[(+)/(−)] group.
In addition, when treated with CAG, the expression of Atrogin‑1 and MuRF1 mRNA,
which was increased by the dexamethasone treatment, showed a signicant concentration‑
dependent decrease. Atrogin‑1 decreased to 0.39 ±0.04 and 0.16 ±0.02 at 50 µg/mL and
100 µg/mL of CAG, and MuRF1 decreased to 0.45 ±0.05 and 0.20 ±0.03, respectively.
The expression of MyoD1 mRNA increased to 1.56 ±0.08 and 2.12 ±0.13 at 50 µg/mL and
100 µg/mL, respectively. Myogenin’s mRNA expression showed a signicant concentration‑
dependent increase, increasing signicantly to 1.44 ±0.14, 2.10 ±0.17, and 2.26 ±0.16 at
each concentration (Table 3). As a result, it was conrmed that UME and CAG have a sig‑
nicant eect on inhibiting muscle degradation and promoting muscle synthesis by reduc‑
ing the expression of Atrogin‑1 and MuRF1 mRNA, which are muscle‑degradation‑related
genes, and increasing the expression of MyoD and Myogenin, which are muscle‑synthesis‑
related genes.

Curr. Issues Mol. Biol. 2024,46 8332
Table 3. Eect of catechin 7‑O‑β‑D apiofuranoside on the expression of muscle‑degradation‑ and
muscle‑synthesis‑related genes.
DEX
(5 µM)
CAG
(µg/mL)
mRNA
Atrogin‑1 MuRF‑1 Myo D Myogenin
− − 0.06 ±0.01 0.11 ±0.02 2.98 ±0.29 3.34 ±0.55
+−1.00 ±0.16 ** 1.00 ±0.08 *** 1.00 ±0.09 *** 1.00 ±0.10 **
+ 10 0.75 ±0.05 0.75 ±0.06 #1.16 ±0.12 1.44 ±0.14 #
+ 50 0.39 ±0.04 ## 0.45 ±0.05 ### 1.56 ±0.08 ## 2.10 ±0.17 ##
+ 100 0.16 ±0.02 ### 0.20 ±0.03 ### 2.12 ±0.13 ### 2.26 ±0.16 ###
DEX: dexamethasone. CAG: Catechin 7‑O‑β‑D apiofuranoside. Values are expressed as mean ±S.E.M. (n = 3).
The target mRNA’s expression was normalized to that of GAPDH. ** p< 0.01, *** p< 0.001 are signicantly dierent
from that of the [(−)/(−)] group. #p< 0.05, ## p< 0.01, and ### p< 0.001 are signicantly dierent from that of the
[(+)/(−)] group.
3.3.5. Akt and mTOR Signaling Pathway‑Related Muscle Protein
Expression Investigation
Akt, also known as Protein kinase B, is known to phosphorylate various muscle pro‑
teins in skeletal muscles, inuencing the growth and proliferation of muscle cells. The sig‑
naling of Akt, activated by IGF‑1, has been reported to enhance muscle protein synthesis
by suppressing the expression of apoptotic muscle proteins, such as caspase‑9 [46]. When
phosphorylation occurs in proteins like IGF‑1 and Tuberous Sclerosis Complex 2 (TSC2),
they become inactivated, translocating to the cytoplasm outside the nucleus and increas‑
ing the activity of mTOR [47]. mTOR is a crucial factor involved in the growth, metabolic
action, and proliferation of cells, functioning as a serine/threonine protein kinase [48,49].
The eect of UME and CAG on Akt signaling activity in dexamethasone‑induced my‑
otube cell atrophy was investigated (Figure 11). The expression of p‑Akt signicantly in‑
creased to 0.59 ±0.02 and 0.58 ±0.02 at UME (Figure 11A,B) concentrations of 100 and
200 µg/mL, respectively. When treated with CAG (Figure 11C,D), the highest treatment
concentration of 100 µg/mL resulted in an increase to 0.95 ±0.09, which was similar to
that of the untreated control group.
Akt expression showed no signicant dierence with the addition of either UME
or CAG. The expression of p‑mTOR increased at all concentrations when treated with
UME (Figure 11E,F), and was signicantly increased to 0.73 ±0.04 at 100 µg/mL. CAG
(Figure 11G,H) led to a signicant increase from 0.64 ±0.07 at 50 µg/mL to 0.90 ±0.09 at
100 µg/mL. The expression of mTOR did not dier signicantly when treated with UME,
but decreased when treated with CAG. These results conrmed that UME and CAG eec‑
tively promote muscle protein synthesis.
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 15
Figure 11. Cont.

Curr. Issues Mol. Biol. 2024,46 8333
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 15
Figure 11. Eect of Ulmus macrocarpa extract (UME) and catechin 7‑O‑β‑D apiofuranoside (CAG)
on protein expression levels of (A–D) phospho‑Akt and Akt in DEX‑treated C2C12 myotubes and
(E–H) phosphor‑mTOR and mTOR in DEX‑treated C2C12 myotubes. UME and CAG were added
to DEX‑treated C2C12 myotubes and cultured for 24 h, respectively. Protein expression levels were
determined using Western blot assay. Values are expressed as mean ±S.E.M. (n = 3). *** p< 0.001 are
signicantly dierent from that of the [(−)/(−)] group. #p< 0.05, ## p< 0.01 are signicantly dierent
from that of the [(+)/(−)] group.
3.3.6. FoxO Signaling Pathway‑Related Muscle Protein Expression Investigation
The FoxO subgroup consists of FoxO1, FoxO3a, FoxO4, and FoxO6, which are located
within muscles. This location protects the DNA‑binding region called FoxO, which is a
transcription factor located in the cell nucleus that regulates various muscle protein sig‑
naling. When FoxO is activated in the nucleus, it decomposes muscle proteins by regulat‑

Curr. Issues Mol. Biol. 2024,46 8334
ing the ubiquitin proteasome and autophagy lysosome systems, which are muscle protein
degradation pathways [47].
We examined the eects of UME and CAG on FoxO signaling activity in
dexamethasone‑induced myotube cell atrophy (Figure 12). The expression of phospho‑
FoxO1 increased in a concentration‑dependent manner when treated with UME
(Figure 12A,B) and signicantly increased when treated with all concentrations of CAG
(Figure 12C,D), 10, 50, and 100 µg/mL, rising to 0.74 ±0.01, 0.77 ±0.04, and 0.87 ±0.04,
respectively. The expression of FoxO1 also increased when treated with dexamethasone
alone, but signicantly decreased when treated with all concentrations of UME. In par‑
ticular, at the highest treatment concentration of 200 µg/mL, it decreased to 0.95 ±0.05,
which was lower than that in the untreated control group. It was shown that the expres‑
sion of phospho‑FoxO3αdecreased when treated with dexamethasone and signicantly
increased when treated with 100 µg/mL of UME (Figure 12E,F) or higher, leading to val‑
ues of 0.79 ±0.04 and 0.76 ±0.04. When treated with CAG (Figure 12G,H), the expression
of phospho‑FoxO3αincreased in a concentration‑dependent manner, leading to a higher
expression, especially at 100 µg/mL, than that seen in the untreated control group. The
expression of FoxO3a signicantly decreased to 1.12 ±0.13 when treated with 200 µg/mL
of UME. It was shown that treatment with CAG resulted in its decreased expression at all
concentrations, particularly at 100 µg/mL, where the expression of FoxO3 was signicantly
reduced, to 0.75 ±0.12, compared to the untreated control group. These results show that
UME and CAG eectively inhibit the expression of muscle‑degrading proteins (atrogin‑1
and MuRF1).
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 17
Figure 12. Cont.

Curr. Issues Mol. Biol. 2024,46 8335
Curr. Issues Mol. Biol. 2024, 46, FOR PEER REVIEW 17
Figure 12. Eect of Ulmus macrocarpa extract (UME) and catechin 7‑O‑β‑D apiofuranoside on pro‑
tein expression levels of (A–D) phospho‑FoxO1 and FoxO1 in DEX‑treated C2C12 myotubes and
(E–H) phospho‑FoxO3a and FoxO3a in DEX‑treated C2C12 myotubes. UME and CAG were added
to DEX‑treated C2C12 myotubes and cultured for 24 h, respectively. Protein expression levels were
determined using Western blot assay. Values are expressed as mean ±S.E.M. (n = 3). * p< 0.05,
** p< 0.01, and *** p< 0.001 are signicantly dierent from that of the [(−)/(−)] group. #p< 0.05,
## p< 0.01, and ### p< 0.001 are signicantly dierent from that of the [(+)/(−)] group.
4. Discussion
In existing research on plants belonging to the Ulmus genus, various outstanding bi‑
ological activities have been reported, including antioxidant, anti‑inammatory, immune‑
modulating eects; the inhibition of angiogenesis; and the regulation of cell apoptosis
in human papilla cells [5,11,12,50]. Plants of the Ulmus genus contain (‑)‑catechin, triter‑
pene, and neolignan glycoside, and these are known to exert their eects through their
me‑tabolites after absorption and metabolism [5–8]. Many of the biological eects of Ulmus
genus are believed to be mediated by its polyphenol catechins [51]. However, research on
muscle loss and muscle atrophy is lacking. Therefore, our goal is to develop a therapeutic
agent that utilizes natural compounds derived from Ulmus macrocarpa for the prevention of
muscle loss and atrophy. Our research team conducted an in vitro mechanism study that
covered four distinct categories—inhibiting oxidative stress, anti‑apoptosis, promoting
muscle protein synthesis, and inhibiting muscle protein degradation—to explore the eec‑
tiveness and mechanisms by which Ulmus macrocarpa prevents muscle loss
and atrophy.
Reactive oxygen species (ROS) are formed within cells, generating free radicals that
increase oxidative stress and trigger muscle cell apoptosis [52]. Bcl‑2 family proteins play
a crucial role in the inhibition and induction of cell apoptosis, with Bcl‑2 acting as an in‑

Curr. Issues Mol. Biol. 2024,46 8336
hibitor and Bax as an inducer [27]. The disruption of this balance leads to mitochondrial
dysfunction, causing the release of cytochrome c from the inner mitochondrial membrane
into the cytoplasm, stimulating various genes to induce cell apoptosis at low levels [32].
In H2O2‑induced C2C12 cells, Bax was not signicantly expressed, indicating that H2O2
does not inuence Bax expression in these cells. However, treatment with catechin 7‑O‑β‑
D apiofuranoside signicantly increased the expression of Bcl‑2. These ndings suggest
that catechin 7‑O‑β‑D apiofuranoside prevents the loss of mitochondrial function due to
ROS in H2O2‑treated C2C12 cells, inhibiting the activation of the apoptosis pathway and
enhancing cell viability. Our results demonstrate that the increased expression of Bcl‑2
in C2C12 cells treated with catechin 7‑O‑β‑D apiofuranoside could potentially block mito‑
chondrial dysfunction, thereby suppressing the induction of cell apoptosis. Additionally,
caspase‑9, a key player in the induction of cell apoptosis, is presumed to induce the activ‑
ity of caspases such as caspase‑3, leading to the expression of PARP and DNA fragmen‑
tation [34]. PARP plays a crucial role in maintaining DNA repair and genetic stability in
normal cells. Our results also indicate that catechin 7‑O‑β‑D apiofuranoside inhibits the ex‑
pression of caspase‑3 and PARP induced by H2O2in C2C12 cells. This outcome highlights
the inhibitory eect of catechin 7‑O‑β‑D apiofuranoside on the induction of caspase‑3 and
PARP in response to H2O2treatments.
Myogenesis becomes imperative when muscle regeneration is required due to the
reduced muscle mass caused by atrophy. It involves the dierentiation of cells into my‑
otubes, regulated by myogenic regulatory factors (MRFs) such as myogenin and
myoD [53,54]. Atrogin‑1 and MuRF1 are recognized as ubiquitin–proteasome protein
degradation factors contributing to muscle atrophy [42,55]. Dexamethasone (DEX),
a prominent glucocorticoid, induces skeletal muscle degradation when inappropriately
used in clinical seings, making it a widely employed agent to simulate muscle cell atrophy
in in vitro systems [35,36]. To assess the impact of Ulmus macrocarpa extract and catechin 7‑
O‑β‑D apiofuranoside on muscle degradation and synthesis in a dexamethasone‑induced
muscle atrophy model, this study investigated the protein and gene expression of key
biomarkers: myoD, myogenin, atrogin‑1, and MuRF1. Treatment with Ulmus macrocarpa
extract or catechin 7‑O‑β‑D apiofuranoside resulted in the decreased expression of atrogin‑
1 and MuRF1, while the expression of myogenin and myoD increased. This conrmed
that catechin 7‑O‑β‑D apiofuranoside mitigates muscle atrophy by downregulating mus‑
cle proteins and genes involved in the muscle synthesis pathway, thereby promoting mus‑
cle dierentiation. Muscle protein synthesis is facilitated through the insulin‑like growth
factor 1/PI3K/Akt signaling pathway, while muscle protein degradation is governed by a
signaling pathway comprising forkhead box O (FoxO) and the ubiquitin–proteasome sys‑
tem [56]. Catechin 7‑O‑β‑D apiofuranoside was found to inhibit muscle atrophy through
the Akt/FoxO signaling pathway. Fluorescent staining under a microscope revealed the
inhibitory eect of catechin 7‑O‑β‑D apiofuranoside on muscle atrophy, with myotube
cell diameters reduced by dexamethasone and returning to normal post‑treatment. This
observation conrmed that the restoration was to a level nearly identical to that of the
control group.
In conclusion, this study demonstrates that Ulmus macrocarpa extract, including cate‑
chin compounds known to have antioxidant, anti‑diabetic [57], and anti‑obesity eects [58],
can prevent and inhibit muscle loss and atrophy. In addition, since the muscle loss preven‑
tion eect of Ulmus macrocarpa extract was veried in a recent in vivo experiment [10], it
is thought that Ulmus macrocarpa extract can be utilized as a key ingredient in the develop‑
ment of functional foods and health supplements for preventing muscle loss and atrophy
and sarcopenic obesity in diabetic patients in the future.
Author Contributions: M.S.K.: Formal analysis and writing—original draft; S.P.: project administra‑
tion; Y.K.: project administration; T.K.: project administration; C.H.L.: project administration; H.J.:
project administration; E.J.K.: methodology and visualization; J.I.J.: methodology and visualization;
S.M.: conceptualization and validation; K.‑H.P.: methodology and visualization; S.E.C.: formal anal‑

Curr. Issues Mol. Biol. 2024,46 8337
ysis, funding acquisition, investigation, project administration, supervision, and writing—review
and editing. All authors have read and agreed to the published version of the manuscript.
Funding: This study was supported by the R&D Program for Forest Science Technology (Project No.
2023469A00‑2325‑EE01) provided by the Korea Forest Service (Korea Forestry Promotion Institute),
partially supported by the Starting Growth Technological R&D Program (TIPS Program, RS‑2023‑
00222349) funded by the Ministry of SMEs and Startups (MSS, Korea) in 2023, and also partially
supported by the Commercialization Promotion Agency for R&D Outcomes (COMPA) funded by
the Ministry of Science and ICT (MSIT) (No. RS‑2024‑00417542).
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not applicable.
Data Availability Statement: The original contributions presented in the study are included in the
article, further inquiries can be directed to the corresponding author.
Conicts of Interest: Sun Eun Choi reports that nancial support was provided by the Korea Forestry
Promotion Institute. Sun Eun Choi reports that nancial support was provided by the Korea Ministry
of Small and Medium Enterprises and Startups. Sun Eun Choi reports that nancial support was
provided by the Ministry of Science and ICT. Sun Eun Choi has patent #10‑2022‑0021852 licensed to
Sun Eun Choi. Sun Eun Choi has patent #PCT/KR2022/006674 licensed to Sun Eun Choi. Sun Eun
Choi has patent #3202787 licensed to Sun Eun Choi. Sun Eun Choi has patent #18/266174 licensed
to Sun Eun Choi. Sun Eun Choi has patent #1‑2023‑03567 licensed to Sun Eun Choi. Sun Eun Choi
has patent #2022395063 licensed to Sun Eun Choi. If there are other authors, they declare that they
have no known competing nancial interests or personal relationships that could have appeared
to inuence the work reported in this paper. Authors Min Seok Kim, Sunmin Park, Yeeun Kwon,
TaeHee Kim were employed by the company Dr. Oregonin Inc. The remaining authors declare that
the research was conducted in the absence of any commercial or nancial relationships that could be
construed as a potential conict of interest.
Abbreviations
Akt protein kinase B
Bax bcl‑2‑associated X protein
Bcl‑2 B‑cell leukemia/lymphoma 2 protein
CAG catechin 7‑O‑β‑D apiofuranoside
DAPI 4′, 6‑diamidino‑2‑phenylindole
DEX dexamethasone
FoxO forkhead box O
HPLC high‑performance liquid chromatography
mTOR mammalian target of rapamycin
MuRF1 muscle RING‑nger protein‑1
MyoD myoblast determination
PARP Poly (ADP‑ribose) polymerases
RT‑PCR real‑time polymerase chain reaction
UME Ulmus macrocarpa extract
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