Endoplasmic reticulum stress may induce apoptosis in gastrocnemius injury of obese mice with OSAHS

doi:10.21203/rs.3.rs-5302463/v1

Download PDF

Research Article

Endoplasmic reticulum stress may induce apoptosis in gastrocnemius injury of obese mice with OSAHS

https://doi.org/10.21203/rs.3.rs-5302463/v1

This work is licensed under a CC BY 4.0 License

You are reading this latest preprint version

Background:Obstructive sleep apnea-hypopnea syndrome (OSAHS) combined with obesity exacerbates clinical symptoms of OSAHS and has been associated with increased endoplasmic reticulum stress (ERS) leading to multi-organ damage including kidneys and liver. However, research on skeletal muscle injury in the context of OSAHS combined with obesity remains limited. This study aims to investigate the mechanisms and potential regulatory role of ERS in gastrocnemius muscle damage induced by OSAHS combined with obesity. Using a chronic intermittent hypoxia (CIH) combined with high-fat diet animal and cellular models, this research focuses on elucidating whether ERS plays a regulatory role in this process. Results from this study may contribute to understanding the pathophysiology of skeletal muscle damage in OSAHS patients with obesity and identify potential therapeutic targets.

Method:Forty male C57/BL6 mice aged 6-8 weeks and weighing 25-30g were randomly divided into four groups: Control group (RC), Chronic Intermittent Hypoxia group (RH), High-Fat Diet group (HC), and Chronic Intermittent Hypoxia combined with High-Fat Diet group (HH). The study aims to observe morphological changes in gastrocnemius muscle among these groups. Real-time quantitative PCR and Western blotting will be used to assess mRNA expression of MyHC1, MYH2, MYOG, MYOD, IRE1α, XBP1s, CHOP, and protein expression of MYH7, MYH2x, MYOG, MYOD1, IRE1α, XBP1s, CHOP, Caspase-3, TNF-α, IL-6 in mouse gastrocnemius muscle tissue. Additionally, a cellular model using C2C12 myotubes will be established combining Chronic Intermittent Hypoxia with High-Fat Diet, with groups including Control (CON), Chronic Intermittent Hypoxia (CIH), High-Fat Diet (PA), and Chronic Intermittent Hypoxia combined with High-Fat Diet (CIH+PA). Changes in cellular morphology post-differentiation and gene/protein expressions related to muscle fibers, endoplasmic reticulum stress, inflammation, damage, and apoptosis will be evaluated.

Results:1.In vivo experimental results: Chronic intermittent hypoxia combined with a high-fat diet leads to structural damage in the gastrocnemius muscle of mice, characterized by the transformation of type I muscle fibers to type II muscle fibers, inhibition of myotube formation, and enhanced endoplasmic reticulum stress. This condition further promotes inflammation and apoptosis within the gastrocnemius muscle tissue.2.In vitro cell culture results: Chronic intermittent hypoxia and high-fat treatment may alter the morphology of C2C12 myotube cells, leading to the transformation of type I muscle fibers to type II muscle fibers. This treatment inhibits myotube formation and differentiation, significantly enhances endoplasmic reticulum stress, and promotes cellular inflammation and apoptosis.

Conclusion :This study elucidates that chronic intermittent hypoxia combined with a high-fat diet may lead to structural damage in the gastrocnemius muscle of mice through the activation of endoplasmic reticulum stress, resulting in a transformation of muscle fiber types from type I to type II. Additionally, chronic intermittent hypoxia combined with a high-fat diet may induce excessive endoplasmic reticulum stress, activating apoptotic pathways and causing severe damage to the gastrocnemius muscle in mice.

chronic intermittent hypoxia

obesity

gastrocnemius

ER stress

injury

Obstructive sleep apnea hypopnea syndrome (OSAHS) is a sleep disorder characterized by recurrent upper airway obstruction or collapse during sleep, leading to restricted airflow or even breathing pauses. Clinical symptoms include snoring, daytime sleepiness, fragmented sleep, apnea episodes, cognitive impairment, and fatigue, posing significant health risks to affected individuals. Research has linked OSAHS closely with various diseases such as hypertension, stroke, coronary artery disease, heart failure, diabetes, metabolic disorders, and cognitive impairments [1].Obesity is widely recognized as the primary risk factor for OSAHS, with every 10% increase in body weight correlating to a six-fold increase in OSAHS risk [2]. Obesity exacerbates OSAHS symptoms, contributing to worsened hypoxia, increased sympathetic nervous system activity, oxidative stress, inflammation, and endothelial dysfunction [3]. Studies have also indicated that short sleep duration may increase the risk of obesity and abdominal obesity by 1.45 times, suggesting that insufficient sleep duration is an important factor in triggering obesity [4]. Therefore, OSAHS and obesity interact synergistically, further exacerbating damage to the body.

Skeletal muscle is the body's largest organ for movement and oxygen consumption, primarily composed of two fiber types: slow-twitch fibers, known as type I fibers (MyHC 1 and MyHC 2a), and fast-twitch fibers, known as type II fibers (MyHC 2b and MyHC 2x). The development of skeletal muscle is mediated by transcription factors, with the myogenic regulatory factor (MRF) family (including MyoD, MyoG, Myf5) playing crucial roles in regulating specific gene transcription in muscle cells, as well as in cell growth, cycle, and differentiation [5]. Slow-twitch fibers possess strong oxidative metabolism and fatigue resistance, whereas fast-twitch fibers have lower mitochondrial content, promoting glycolytic metabolism with weaker fatigue resistance but stronger contraction force. Muscle fibers can undergo type transformation under different environmental and stimuli conditions. Chronic intermittent hypoxia (CIH) can lead to morphological and functional changes in skeletal muscle. Previous research indicates that OSAHS patients have a wider area of fast-twitch fibers [6], potentially linked to oxidative stress and glycolytic metabolism induced by chronic intermittent hypoxia.

The pathophysiological basis of OSAHS is CIH. Multiple studies have linked CIH with liver injury, insulin metabolism abnormalities, heart disease, and pulmonary fibrosis, suggesting mechanisms involving endoplasmic reticulum stress (ERS), inflammatory responses, and cell apoptosis [7, 8]. Additionally, research has shown activation of various ERS markers in adipose tissue of obese individuals, exacerbating inflammatory responses [9]. Therefore, we hypothesize that the ERS pathway may mediate skeletal muscle damage induced by chronic intermittent hypoxia combined with a high-fat diet.This study aims to explore the mechanisms and potential pathways underlying skeletal muscle damage in OSAHS patients with obesity by establishing animal and cell models of chronic intermittent hypoxia combined with high-fat diet.

1) Establishment of animal model

Forty male C57/BL6 mice, aged 6-8 weeks and weighing 25-30g, were randomly divided into four groups of 10 each: Control group (RC), Chronic Intermittent Hypoxia group (RH), High-Fat Diet group (HC), and Chronic Intermittent Hypoxia combined with High-Fat Diet group (HH).RC group: Mice were fed a standard diet under normal oxygen conditions for 14 weeks.RH group: Mice were exposed to normal oxygen conditions for the first 8 weeks, followed by 6 weeks of exposure to intermittent hypoxia (oxygen concentration fluctuating between 6% to 21% for 8 hours daily).HC group: Mice were fed a high-fat diet under normal oxygen conditions for the entire 14 weeks.HH group: Mice were exposed to normal oxygen conditions for the first 8 weeks, followed by 6 weeks of intermittent hypoxia exposure, while consuming a high-fat diet throughout the study period.

2) Cell culture and differentiation

C2C12 cells（Procell，Wuhan）derived from mouse myoblasts, were cultured in growth medium containing 10% fetal bovine serum, 1% penicillin-streptomycin, and high-glucose DMEM. The culture dishes were maintained in a cell culture incubator at 37°C with 5% CO2. Upon reaching 80%~90% confluence, the cells were induced to differentiate for 5 days in differentiation medium consisting of high-glucose DMEM supplemented with 2% horse serum, resulting in the formation of C2C12 myotube cells.

3) Establishment of the cell model

After achieving approximately 80% cell fusion, C2C12 cells were dissociated with trypsin, resuspended, and seeded into 6-well plates at a density of 1.1 x 10^6 cells per well. They were cultured under standard conditions at 37°C with 5% CO2 to allow for differentiation into C2C12 myotubes. Following differentiation, the cells were randomly divided into four groups:Control group (CON): Cells cultured conventionally in a standard cell culture incubator for 48 hours.Chronic intermittent hypoxia group (CIH): Cells placed in an intermittent hypoxia chamber for 48 hours.Palmitic acid group (PA): Cells treated with medium containing 200μmol/L palmitic acid (PA) and cultured in a standard incubator for 48 hours.Chronic intermittent hypoxia combined with palmitic acid group (CIH+PA): Cells treated with medium containing 200μmol/L PA and cultured in the intermittent hypoxia chamber for 48 hours.The chronic intermittent hypoxia condition involved cycles where cells were exposed to 21% O2, 5% CO2, and 74% N2 for 15 minutes, followed by 5% O2, 5% CO2, and 90% N2 for another 15 minutes, maintaining oxygen concentrations between 5% and 21% within the chamber.

4) CCK8 cell proliferation experiment

C2C12 cells were digested and resuspended to achieve a confluence of approximately 80%. They were then seeded at a density of 1×10^4 cells per well in a 96-well plate. The cells were cultured in high glucose DMEM containing 10% fetal bovine serum (FBS) at 37°C in a 5% CO2 atmosphere until reaching approximately 90% confluence. Subsequently, the cells were differentiated in high glucose DMEM supplemented with 2% horse serum for 5 days.After differentiation, the cells were treated with various concentrations (100, 150, 200, 250, 300, 350, 400 μmol/L) of sodium palmitate (PA) for 24 hours. Following treatment, the culture medium was removed, and the wells were washed with PBS. Then, each experimental well was replenished with 100 μL of DMEM culture medium and 10 μL of CCK8 Solution. The plate was gently shaken to mix the solutions and incubated at 37°C in a dark environment. After 1 hour of incubation, the optical density (OD) at 450 nm was measured for each well.

5) Oil red O staining

After completion of differentiation of C2C12 myotubes, the culture medium was discarded, and the cells were gently washed twice with PBS. They were fixed with 4% paraformaldehyde at room temperature for 30 minutes, followed by removal of the fixative solution and two additional washes with PBS. Subsequently, the cells were rinsed with 60% isopropanol and then stained with Oil Red O working solution for 30 minutes.After discarding the staining solution, the cells were briefly rinsed with 60% isopropanol for 5 seconds to ensure clarity of the interstitial spaces. The isopropanol was then discarded, and the cells were washed three times with distilled water until no excess dye was visible. Hematoxylin staining solution was added for nuclear staining for 2 minutes, followed by removal of the staining solution and three additional rinses with distilled water. Finally, the cells were observed under a microscope (IX 51; Olympus Corporation, Japan), with distilled water covering them for examination.

6) Detection of cell triglyceride and total cholesterol

According to TG and T-CHO detection preparations (Nanjing Jiancheng Bioengineering Research Institute Co., LTD.) : 1. Cell collection: After digestion of the PA-treated C2C12 muscle tube cells, the cell suspension was centrifuged at 1000 RPM for 10min, the supernatant was discarded and cell precipitation was left, the PBS was cleaned 1-2 times, and the centrifuge was continued at 1000 RPM for 10min, the supernatant was discarded and cell precipitation was left. 2, cell crushing: Add 0.2ml homogenate medium PBS, 300W, 3-5 seconds/time ultrasonic homogenate, each interval of 30 seconds, repeat 3-5 times. After homogenization, a mixture of blank holes, standard holes and sample holes was prepared according to the test reagent requirements, and incubated at 37℃ for 10min. The absorbance value of 500nm was measured, and TG and T-CHO levels were calculated. TG/T-CHO level (mmol/gprot) = (determination group - blank group)

7) RNA extraction, reverse transcription, real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR)

Total RNA was extracted from freeze-dried mouse gastrocnemius and C2C12 muscle duct cells with total RNA extraction reagent (NCM Biotech, China). RNA concentration and purity were measured by nucleic acid analyzer. The cDNA was obtained by reverse transcription using reverse transcription reagent (HiScript III RT SuperMix for qPCR (+gDNA wiper), Vazyme, China) at low temperature. Real-time PCR (qRT-PCR) was performed using ChamQ SYBR qPCR Master Mix (Vazyme, China) suitable for SYBR Green I. The gene expression level was determined by GAPDH, and the relative expression level of target gene was calculated by 2-∆∆Ct method. The target gene and primer sequences are listed in the table (Table 1). Each experiment was repeated at least three times. The data for each sample were normalized using GAPDH and the multiple change values were calculated using the ΔΔCt method.

Table 1: Primer sequence table

Gene name	Primer sequences
GAPDH	F：GCCAAGGCTGTGGGCAAGGT R: TCTCCAGGCGGCACGTCAGA
MYHC1	F: GAATGGCAAGACGGTGACTGTG R: GGAAGCGTAGCGCTCCTTGAG
MYH2x	F: CCAATGAAACCAAGACTCCTGG R: TGCTATCGATGAACTGTCCCTC
MYOD	F: GGCTCTCTCTGCTCCTTTGA R: GTAGGGAAGTGTGCGTGCTC
MYOG	F: AGGAAGTCTGTGTCGGTGGA R: AGGCGCTCAATGTACTGGAT
IRE1α	F: GGATGAGGAAACCAGAATGG R: TGGGTGCTCGTCTGATTCTC
XBP1s	F: GAGTCCGCAGCAGGTG R: GTGTCAGAGTCCATGGGA
CHOP	F: GAGAATGAAAGGAAAGTGGCAC R: ATGTGTGCTGTGTGTGTGTTCC

8) Western Blot (WB)

Mouse gastrocnemius and C2-C12 muscle duct cells were fully lysed in RIPA buffer containing protease inhibitor and PMSF, and the tissue and cell proteins were extracted. Gel was prepared according to the instructions of One-Step PAGE Gel Fast Preparation Kit (12%) (Vazyme, China). The total amount of protein sampled was 30 µg. For electrophoresis (Bio-Rad, USA), a constant voltage of 80 V is applied for 25 minutes, and a constant voltage of 110 V is applied for 60-80 minutes. Fast transfer solution (NCM Biotech, China) 400 mA constant current 40 min transfer. Rapid closure solution (NCM Biotech, China) and small shock closure at room temperature for 15 minutes. The primary antibody was incubated at 4 ℃ overnight. The next day, TBST washed the film for 10 minutes, three times in total, and incubated the film with the second antibody for 1 hour. The target protein was visualized using the enhanced ECL chemiluminescence detection kit (Vazyme, China). The strip gray values were analyzed using ImageJ software. The antibodies used are listed in table 2.

Table 2: List of the experimentally-related antibodies

Antibody name	producer
MYH7 polyclonal antibody	Wuhan Sanying Biotechnology Co., LTD
Fast Myosin Skeletal Heavy chain Antibody（MYH2x）	Jiangsu Kinke Biological Research Center Co., LTD
MYOD1 antibody	Wuhan Sanying Biotechnology Co., LTD
Rabbit Anti-Myogenin（MYOG）	Beijing Boosen Biotechnology Co., LTD
Anti-rabbit second antibody	Bio-Rad Corporation
Rabbit anti-cleaved Caspase 3 antibody	Cell Signaling Technology Inc
Rabbit anti-mouse IL-6 antibody	Cell Signaling Technology Inc
Rabbit anti-mouse TNF- α antibody	Cell Signaling Technology Inc
Rabbit anti-mouse IRE 1 α antibody	Wuhan Doctor De Biological Engineering Co., LTD
Rabbit anti-mouse XBP 1s antibody	Cell Signaling Technology Inc
Rabbit anti-mouse CHOP antibody	abcam Inc
Rabbit anti-rat BAX antibody	Cell Signaling Technology Inc
Rabbit anti-mouse BCL-2 antibody	abcam Inc

9) Data processing and statistical analysis

The data are presented as mean ± standard deviation (x ± s). Paired t-tests were used to determine whether differences between groups were significant. Protein band grayscale values were analyzed using ImageJ software, and all statistical analyses were performed using GraphPad Prism 9. Significance levels were defined as *P < 0.05, **P < 0.01, and ***P < 0.001. All experiments were repeated ≥3 times.

3.1 CIH combined with a high-fat diet leads to structural damage in the gastrocnemius muscle tissue of mice.

Histological examination using hematoxylin and eosin (HE) staining was conducted to observe morphological changes in the gastrocnemius muscle of mice across different treatment groups: RC, RH, HC and HH. The results revealed that in the RC group (control), muscle fibers appeared well-organized in a sequential arrangement. In the RH group, mild morphological damage was observed with increased interstitial spaces between muscle fibers and a reduction in cross-sectional area. The HC group showed pronounced morphological abnormalities including thicker muscle fibers, occasional fiber fragmentation, irregular cross-sectional shapes, and significantly increased interstitial spaces compared to the control group. The HH group exhibited the most severe muscle fiber damage, characterized by increased fiber thickness, widespread fiber fragmentation, and markedly enlarged interstitial spaces.These findings demonstrate that CIH and HFD induce significant structural damage to mouse gastrocnemius muscle fibers(Fig.1).

3.2 CIH combined with a high-fat diet leads to a transition of type I muscle fibers to type II muscle fibers in mouse gastrocnemius muscle.

This study utilized WB and qPCR to assess the expression levels of type I muscle fiber markers MYH7 protein and MyHC1 gene, and type II muscle fiber markers MYH2x protein and MYH2x gene, in the gastrocnemius muscle of mice across four experimental groups: RC (control), RH (CIH), HC (HFD), and HH (CIH + HFD). WB and qPCR results indicated that compared to the RC group, both HC and HH groups exhibited a gradual decrease in MYH7 protein expression, suggesting suppression of type I fiber formation under conditions of HFD and CIH, with the HH group showing more significant effects (Fig.2B). Conversely, the RH group showed significantly increased MYH7 protein and MyHC1 gene expression compared to the control, indicating that CIH alone may promote type I fiber formation (Fig..2.B and D).Furthermore, both MYH2x protein and MYH2x gene expression showed an increasing trend in the HC and HH groups (Fig.2.C and E), with the HH group exhibiting statistically significant differences. These findings suggest that prolonged exposure to CIH combined with HFD leads to a reduction in type I muscle fibers and an increase in type II muscle fibers in the mouse gastrocnemius muscle. Thus, CIH combined with HFD appears to facilitate a transition of type I muscle fibers to type II muscle fibers in mice.

3.3 CIH combined with HFD inhibited the formation of muscle duct cells in gastrocnemius muscle of mice.

In this study, we investigated the expression of muscle tube formation-related genes and proteins MYOG and MYOD in the gastrocnemius muscle of mice subjected to CIH combined with HFD, compared to a control group (RC) and two experimental groups (RH, HC, HH). qPCR analysis revealed a significant decrease in MYOG gene expression in the RH, HC, and HH groups compared to the RC group (P<0.001), with the HH group showing the most pronounced reduction(Fig.3.A and B). Similarly, MYOD gene expression was significantly decreased in the RH and HH groups (P<0.001). Western blotting results indicated a notable reduction in MYOG and MYOD1 protein expression levels in all three experimental groups (RH, HC, HH) compared to the control group (RC)(Fig.3.C,D and E), suggesting that CIH combined with HFD significantly inhibits myotube differentiation and formation in mouse gastrocnemius muscle.

3.4 CIH combined with HFD enhances endoplasmic reticulum stress in gastrocnemius cells of mice

To investigate whether CIH combined with HFD induces ERS in mouse gastrocnemius muscle, we evaluated the expression of ER stress-related proteins and genes, including inositol-requiring enzyme 1 alpha (IRE1α), X-box binding protein 1 spliced form (XBP1s), and C/EBP homologous protein (CHOP). qPCR analysis revealed that in the HH group, the expression of IRE1α, XBP1s, and CHOP was significantly increased compared to the RC group. In the RH group, IRE1α and XBP1s expression decreased compared to the RC group, while in the HC group, IRE1α and CHOP expression increased with no significant change in XBP1s expression compared to the RC group(Fig.4.A,B and C). These results suggest that enhanced ERS in mouse skeletal muscle may be more closely associated with HFD, with a more significant effect observed when CIH and HFD are combined.Western blotting results corroborated these findings, demonstrating a significant increase in the expression of ER stress-related proteins (IRE1α, XBP1s, and CHOP) in the IRE1α pathway when CIH was combined with HFD, showing statistical significance(Fig.4.D,E,F and G).Overall, our findings indicate that CIH combined with HFD enhances ERS in mouse gastrocnemius muscle cells.

3.5 CIH combined with HFD resulted in significant inflammatory injury and apoptosis of gastrocnemius cells in mice

This study aimed to assess the impact of CIH combined with HFD on mouse gastrocnemius muscle damage, focusing on the expression levels of apoptosis-related Caspase-3, and inflammation-related TNF-α and IL-6 proteins. The results indicated that Caspase-3 and TNF-α expression levels were significantly increased only in the HH group compared to other groups, showing statistical significance. On the other hand, IL-6 expression was significantly elevated in all three treatment groups compared to the control group, with the HH group exhibiting the most pronounced increase (P < 0.05)(Fig.5).These findings suggest that CIH combined with HFD promotes inflammatory damage and apoptosis in mouse gastrocnemius muscle cells.

3.6 CIH combined with sodium palmitate (PA) resulted in morphological changes of C2C12 muscle duct cells

The effects of CIH combined with PA treatment on differentiated C2C12 myotubes cultured for 5 days were evaluated using CCK8 cell proliferation assays over a 48-hour period (Fig.6 A). The successful establishment of a high-fat cellular model was confirmed by Oil Red O staining (Fig.6 B). Compared to the control group (CON), the CIH group exhibited widened diameters of C2C12 myotubes, partial nuclear morphology damage, and uneven cluster distribution. The PA treatment group showed significantly reduced myotube formation compared to CON, characterized by narrower myotube cell diameters and disrupted continuity. In the CIH combined with PA treatment group, there was a notable decrease in C2C12 myotube formation, with irregular diameters, significantly increased nuclear fragmentation and dissolution, and pronounced morphological abnormalities(Fig.6 C). These cell experiments suggest that CIH combined with PA treatment may lead to certain impairments and inhibitions in the formation and differentiation of C2C12 myotubes, resulting in morphological changes.

3.7 CIH combined with PA resulted in the transformation of type I muscle fibers to type II muscle fibers in C2C12 muscle tube cells.

As depicted in Fig.7, Western blotting results showed a significant decrease in the expression level of type I muscle fiber protein MYH7 following combined CIH and PA treatment in the experimental groups (P<0.05). Conversely, the expression of type II muscle fiber protein MYH2x markedly increased in the experimental groups compared to the control group, with the CIH+PA group showing the most pronounced effect. These findings suggest that CIH with PA-induced high-fat treatment leads to a transition from type I to type II muscle fibers in C2C12 myotube cells. Notably, in cellular experiments, the CIH group alone did not show an increase in MYH7 expression, contrasting with animal experiment results. This discrepancy may be attributed to more complex influencing factors in animal experiments, potentially promoting growth in both type I and type II muscle fibers in mouse gastrocnemius tissue under CIH conditions. Experimental errors cannot be ruled out as contributing to these observations.

3.8 CIH combined with PA inhibited the formation of C2C12 muscle duct cells

The gene and protein expressions of myogenin (MYOG) and myogenic differentiation factor (MYOD) in C2C12 myotube cells were evaluated using q-PCR and Western blot (WB) assays. Results showed that both MYOG and MYOD gene and protein expressions were significantly reduced in the experimental groups compared to the CON group. Specifically, MYOD1 protein expression was markedly decreased in the CIH+PA group (P < 0.001)(Fig.8). Thus, it can be concluded that CIH combined with PA treatment suppresses the formation and differentiation of C2C12 myotube cells.

3.9 CIH combined with PA resulted in enhanced endoplasmic reticulum stress in C2C12 muscle duct cells

To investigate the effects of CIH combined with PA treatment on endoplasmic reticulum (ER) stress at the cellular level, this experiment used WB to assess the expression levels of ER S-related proteins GRP78, CHOP, and the IRE1 pathway proteins IRE1α and XBP1s in C2C12 myotube cells under different treatment conditions.In vitro cell experiments revealed that the expression levels of ERS-related proteins GRP78, CHOP, IRE1α, and XBP1s were significantly elevated in the experimental groups, particularly in the CIH+PA group, with statistical significance. These findings indicate that CIH combined with PA treatment markedly enhances ERS in C2C12 myotube cells(Fig.9).The results suggest that the combination of CIH and PA treatment induces significant ERS in C2C12 myotube cells at the cellular level, as evidenced by the upregulation of key ERS markers.

3.10 CIH in combination with PA treatment promoted inflammatory damage and apoptosis in C2C12 myotube cells

To observe the effects of CIH combined with PA treatment on cellular damage in C2C12 myotube cells, this study utilized WB to assess the expression of cell apoptosis and inflammation-related proteins across different treatment groups.The results showed that apoptotic markers Caspase-3 and Bax were significantly upregulated in the CIH+PA group compared to the CON group, while the expression of the anti-apoptotic marker Bcl-2 was markedly decreased in the CIH+PA group, all with statistical significance (P<0.05). These findings suggest that CIH combined with PA treatment promotes apoptosis in C2C12 myotube cells(Fig.10 A,B,C,D and E). Additionally, inflammation marker IL-6 protein expression was measured, revealing a significant increase during CIH+PA treatment, also with statistical significance (P<0.05). This indicates that CIH combined with PA treatment promotes an inflammatory response in C2C12 myotube cells(Fig.10 A and F).The findings highlight that the combined treatment of CIH and PA induces apoptosis and inflammation in C2C12 myotube cells at the cellular level, as evidenced by altered expression levels of apoptotic and inflammatory markers.

The prevalence of OSAHS significantly increases in obese populations. Some studies suggest that the accumulation of visceral fat in obese patients leads to respiratory restrictions, with additional accumulation of fat around the neck contributing to upper airway compression [10]. The close relationship between OSAHS and obesity suggests that obesity may exacerbate clinical symptoms and risk factors associated with OSAHS. Their synergistic interaction may significantly worsen organ function damage, although the specific mechanisms require further investigation. Currently, there is limited research on how OSAHS combined with obesity affects skeletal muscle in the body. Studies have indicated that CIH may increase fast-twitch muscle fibers and enhance skeletal muscle fatigue [11, 12]. Conversely, other research suggests that isolated OSAHS may have minimal impact on skeletal muscle changes, with obesity potentially playing a larger role in muscle damage [13].This study reveals that CIH combined with HFD results in the conversion of type I muscle fibers to type II muscle fibers. In contrast, the CIH-only group exhibited an increase in both type I and type II muscle fibers, suggesting that CIH alone may lead to increased proliferation of skeletal muscle fibers without significantly affecting fiber type transformation. The underlying mechanisms of this phenomenon remain unclear.

The development of skeletal muscle is a highly complex process involving differentiation and proliferation of muscle cells, formation of myotubes, maturation of muscle fibers, and regulation of gene expression through signaling pathways and transcription factors. The muscle regulatory factor (MRF) family plays a crucial role in promoting skeletal muscle formation. Key members of MRF include myogenic regulatory factor 4 (MRF4), myogenin, myogenic differentiation factor (MYOD), and myogenic factor 5 (Myf5). MYF5, MYOD, and MRF4 act as myogenic determination factors guiding myogenic progenitor cells to establish the skeletal muscle lineage [14，15], primarily responsible for directed differentiation and proliferation of muscle cells. Myogenin primarily participates in the differentiation, proliferation of myocytes, and myotube fusion, forming mature muscle fibers.

In obese individuals, muscle cells can secrete high levels of pro-inflammatory molecules, leading to muscle inflammation and inhibition of myogenic differentiation. Obesity also increases leptin production, causing leptin resistance, which suppresses respiratory centers, alters normal respiratory dynamics of the upper airway, exacerbates hypoxia, increases sympathetic nervous system activity, oxidative stress, inflammation, and impairs endothelial function [16]. Our experiments have revealed that either CIH or HFD alone can reduce the expression of myotube formation-related proteins MYOG and MYOD, with the most significant downregulation observed when CIH and HFD act synergistically. Experimental results suggest that the combined effect of CIH and HFD may lead to prolonged intermittent hypoxia and increased lipotoxicity, resulting in elevated reactive oxygen species, mitochondrial damage, and abnormal autophagy. This further inhibits the formation and differentiation of gastrocnemius muscle fibers in mice.

CIH combined with HFD contributes to tissue damage primarily through the induction of ERS, a focus of our study. ERS initiates the unfolded protein response (UPR) through three signaling pathways: inositol-requiring enzyme 1 (IRE1), protein kinase RNA-like ER kinase (PERK), and activating transcription factor 6 (ATF6). These pathways aid in the degradation of unfolded or misfolded proteins to maintain ER homeostasis. Prolonged ERS can induce cell apoptosis via the IRE1 and PERK pathways [17].Our research has found that CIH combined with HFD leads to excessive activation of endoplasmic reticulum stress, evidenced by significantly increased expression of IRE1α and XBP1s. Additionally, studies by Liu et al. have shown that OSAHS is associated with elevated levels of various inflammatory cytokines, particularly IL-1, IL-6, and CRP, with changes in cytokine levels closely correlated with age, body mass index (BMI), and apnea-hypopnea index (AHI) [18].

Through animal and cell experiments, we observed that CIH combined with HFD induces apoptosis in gastrocnemius muscle cells of mice and upregulates levels of apoptosis-related proteins Caspase-3, BAX, inflammatory cytokines IL-6, TNF-α. This combined treatment may enhance oxidative stress in tissues and cells, leading to sustained UPR, exacerbating hypoxia, protein accumulation, calcium metabolism disorders, decreased cellular ATP levels, and oxidative stress [19].Activation of IRE1α processes unspliced XBP1 mRNA to generate active XBP1, which translocates to the nucleus and upregulates CHOP expression. This activation may initiate apoptotic cell death through CHOP. Moreover, sustained UPR may activate NF-κB through the IRE1 pathway, promoting inflammatory responses [20], aligning with our experimental findings. However, further investigation is needed to fully elucidate the specific mechanisms. Currently, our research primarily focuses on the IRE1 pathway, while other pathways remain to be explored, and whether additional pathways are jointly regulated remains unknown.

This study conducted research from both a macroscopic perspective in animal experiments and a microscopic perspective in in vitro cell experiments. The results indicate that CIH combined with HFD leads to abnormal damage in the morphological structure of the gastrocnemius muscle fibers in mice, with a shift from type I to type II muscle fibers. Additionally, it inhibits the formation and differentiation of myotubes and promotes inflammation and apoptosis in the gastrocnemius muscle cells. This conclusion may be closely related to the ERS apoptotic pathways. However, the study currently lacks experiments involving the knockout of ERS-related pathway genes, and further research is needed to exclude the influence of other pathways. Future research might explore specific therapeutic strategies to mitigate CIH and HFD-induced cellular damage, offering potential treatment targets for patients with OSAHS and obesity.

Acknowledgments

Thanks to Jiangsu University and the Central Laboratory of Affiliated Hospital of Jiangsu University for providing the experimental platform, and thanks to the members of the experimental team for their support.

Author contributions

H.P.H and Q.S. contributed significantly to the conception and design of this work. H.P.H and Q.S. contributed equally to this manuscript. H.P.H was responsible for guiding and arranging the experimental process, and Q.S. was responsible for the implementation of the experiment and data collection and analysis. Q.S. processed the data and participated in writing and revising the manuscript. H.P.H strictly revised the content of the manuscript and finally approved the manuscript for publication.

Funding

City "169" project Huang Hanpeng YLJ202105,Grant ID:000408720005

Availability of data and materials

All data generated or analysed during this study are included in this published article .

Ethical approval and consent to participate

All experimental activities will comply with relevant experimental animal management regulations and be approved by the Experimental Animal Ethics Committee of Jiangsu University.(Zhenjiang, China).

Consent for publication

All the authors agreed to publish it.

Competing interests

The authors declare no competing interests.

Arnaud C, Bochaton T, Pépin JL, Belaidi E. Obstructive sleep apnoea and cardiovascular consequences: Pathophysiological mechanisms. Arch Cardiovasc Dis. 2020;113(5):350–358. 10.1016/j.acvd.2020.01.003. Epub 2020 Mar 26. PMID: 32224049.
Madkikar N, Pandey S, Ghaisas V. Multi Level Single Stage: Barbed Reposition Pharyngoplasty and Nasal Surgery in Treatment of OSA-Our Experience. Indian J Otolaryngol Head Neck Surg. 2019;71(3):309–14. 10.1007/s12070-019-01694-y. Epub 2019 Jun 27. PMID: 31559196; PMCID: PMC6737204.
Salzano G, Maglitto F, Bisogno A, Vaira LA, De Riu G, Cavaliere M, di Stadio A, Mesolella M, Motta G, Ionna F, Califano L, Salzano FA. Obstructive sleep apnoea/hypopnoea syndrome: relationship with obesity and management in obese patients. Acta Otorhinolaryngol Ital. 2021;41(2):120–30. 10.14639/0392-100X-N1100. PMID: 34028456; PMCID: PMC8142730.
Muscogiuri G, Barrea L, Annunziata G et al. Obesity and sleep disturbance: the chicken or the egg? Crit Rev Food Sci Nutr. 2019;59(13):2158–2165. doi: 10.1080/10408398.2018.1506979. Epub 2018 Oct 18. PMID: 30335476.
Wu H, Ballantyne CM. Skeletal muscle inflammation and insulin resistance in obesity. J Clin Invest. 2017;127(1):43–54. 10.1172/JCI88880. Epub 2017 Jan 3. PMID: 28045398; PMCID: PMC5199705.
Matsumoto T, Tanizawa K, Tachikawa R, et al. Associations of obstructive sleep apnea with truncal skeletal muscle mass and density. Sci Rep. 2018;8(1):6550. 10.1038/s41598-018-24750-z. PMID: 29695811; PMCID: PMC5916913.
Shi Z, Xu L, Xie H, Ouyang R, et al. Attenuation of intermittent hypoxia-induced apoptosis and fibrosis in pulmonary tissues via suppression of ER stress activation. BMC Pulm Med. 2020;20(1):92. 10.1186/s12890-020-1123-0. PMID: 32299413; PMCID: PMC7161195.
Hou Y, Yang H, Cui Z, Tai X, Chu Y, Guo X. Tauroursodeoxycholic acid attenuates endoplasmic reticulum stress and protects the liver from chronic intermittent hypoxia induced injury. Exp Ther Med. 2017;14(3):2461–8. 10.3892/etm.2017.4804. Epub 2017 Jul 19. PMID: 28962181; PMCID: PMC5609300.
Dasgupta A, Bandyopadhyay GK, Ray I, Bandyopadhyay K, et al. Catestatin improves insulin sensitivity by attenuating endoplasmic reticulum stress: In vivo and in silico validation. Comput Struct Biotechnol J. 2020;18:464–81. 10.1016/j.csbj.2020.02.005. PMID: 32180905; PMCID: PMC7063178.
Bozkurt NC, Beysel S, Karbek B, Unsal İO, Cakir E, Delibasi T. Visceral Obesity Mediates the Association Between Metabolic Syndrome and Obstructive Sleep Apnea Syndrome. Metab Syndr Relat Disord. 2016;14(4):217–21. 10.1089/met.2015.0086. Epub 2016 Mar 22. PMID: 27003688.
Sauleda J, García-Palmer FJ, Tarraga S, Maimó A, Palou A, Agustí AG. Skeletal muscle changes in patients with obstructive sleep apnoea syndrome. Respir Med. 2003;97(7):804 – 10. 10.1016/s0954-6111(03)00034-9. PMID: 12854630.
Wåhlin Larsson B, Kadi F, Ulfberg J, Piehl Aulin K. Skeletal muscle morphology and aerobic capacity in patients with obstructive sleep apnoea syndrome. Respiration. 2008;76(1):21 – 7. 10.1159/000126492. Epub 2008 Apr 11. PMID: 18408358.
Matsumoto T, Tanizawa K, Tachikawa R, Murase K, Minami T, Inouchi M, Handa T, Oga T, Hirai T, Chin K. Associations of obstructive sleep apnea with truncal skeletal muscle mass and density. Sci Rep. 2018;8(1):6550. 10.1038/s41598-018-24750-z. PMID: 29695811; PMCID: PMC5916913.
Comai G, Tajbakhsh S. Molecular and cellular regulation of skeletal myogenesis. Curr Top Dev Biol. 2014;110:1–73. 10.1016/B978-0-12-405943-6.00001-4. PMID: 25248473.
Weintraub H, Davis R, Tapscott S, Thayer M, Krause M, Benezra R, Blackwell TK, Turner D, Rupp R, Hollenberg S et al. The myoD gene family: nodal point during specification of the muscle cell lineage. Science. 1991;251(4995):761-6. 10.1126/science.1846704. PMID: 1846704.
Salzano G, Maglitto F, Bisogno A, Vaira LA, De Riu G, Cavaliere M, di Stadio A, Mesolella M, Motta G, Ionna F, Califano L, Salzano FA. Obstructive sleep apnoea/hypopnoea syndrome: relationship with obesity and management in obese patients. Acta Otorhinolaryngol Ital. 2021;41(2):120–30. 10.14639/0392-100X-N1100. PMID: 34028456; PMCID: PMC8142730.
Nadeem R, Molnar J, Madbouly EM, Nida M, Aggarwal S, Sajid H, Naseem J, Loomba R. Serum inflammatory markers in obstructive sleep apnea: a meta-analysis. J Clin Sleep Med. 2013;9(10):1003–12. 10.5664/jcsm.3070. PMID: 24127144; PMCID: PMC3778171.
Liu X, Ma Y, Ouyang R, Zeng Z, Zhan Z, Lu H, Cui Y, Dai Z, Luo L, He C, Li H, Zong D, Chen Y. The relationship between inflammation and neurocognitive dysfunction in obstructive sleep apnea syndrome. J Neuroinflammation. 2020;17(1):229. 10.1186/s12974-020-01905-2. PMID: 32738920; PMCID: PMC7395983.
Namgaladze D, Khodzhaeva V, Brüne B. ER-Mitochondria Communication in Cells of the Innate Immune System. Cells. 2019;8(9):1088. 10.3390/cells8091088. PMID: 31540165; PMCID: PMC6770024.
Sikkeland J, Lindstad T, Nenseth HZ, Dezitter X, Qu S, Muhumed RM, Ertunc ME, Gregor MF, Saatcioglu F. Inflammation and ER stress differentially regulate STAMP2 expression and localization in adipocytes. Metabolism. 2019;93:75–85. 10.1016/j.metabol.2019.01.014. Epub 2019 Jan 30. PMID: 30710574; PMCID: PMC6460919.

No competing interests reported.

WB.doc

Download PDF

Editorial decision: Revision requested
24 Oct, 2024
Editor assigned by journal
23 Oct, 2024
Submission checks completed at journal
23 Oct, 2024
First submitted to journal
21 Oct, 2024

You are reading this latest preprint version

Endoplasmic reticulum stress may induce apoptosis in gastrocnemius injury of obese mice with OSAHS

Status:

Version 1

Abstract

Figures

1. Background

2. Materials and methods

3. Results

Discussion

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1