Animal experiments were approved by the Jiangsu Provincial Laboratory Animal Management Committee (No. 20180305-004). All animal studies were conducted according to the AAALAC and the IACUC guidelines. Male ICR mice (25 g±2 g, 8–10 weeks) were provided by the Experimental Animal Center of Nantong University and housed in a temperature-controlled room at 22 C with a 12 h light/dark cycle, with free access to water and diets. The mice were divided into control group (Control) and streptozotocin (STZ)-induced diabetes group. The mice were fasted for 15 h overnight, and then were injected with STZ (#S0130, Sigma) at a single dose of 150 mg/kg (dissolved in 100 mmol/L citrate buffer, pH 4.5) in the diabetes group. The mice received an equal volume of citrate buffer in control group. Random blood glucose was measured on the third and seventh day, and if the levels of blood glucose were consistently above 16.7 mmol/L, the mice were considered diabetic mice and included in the study. Subsequently, the diabetic mice were randomly divided into two groups: the diabetes group (DM) and the diabetes group treated with celecoxib (at a dose of 5 mg/kg/d, Pfizer, NY, United States) (DM + CXB), given through intraperitoneal injection for 28 consecutive days. The mice in control and DM groups were given an equal volume of normal saline. The motor functions were used to initially assess the development of diabetic sarcopenia (diabetes-induced muscle atrophy). After 4 weeks of celecoxib treatment, the mice were euthanized with pentobarbital sodium injection, and muscle samples were immediately collected to further analysis. Subsequently, the samples were either frozen at −80°C for later experiments, fixed in 4% paraformaldehyde (PFA, #G1101, Servicebio), or embedded in pre-cooled OCT (#4583, Leica, Germany) for subsequent experiments.

(1) Rotarod Test

In the week before formal testing, each mouse was subjected to adaptive running training, with the speed gradually increasing from 5 r/min to 10 r/min to 15 r/min, for 5 min each time, three consecutive times. If the mouse was initially unable to run, tail assistance was provided. For formal testing, the mouse was placed on the lane of the rotarod apparatus (Panlab, LE8500) with the minimum speed at 4 r/min. After the mouse stabilized, the speed was gradually increased from 4 r/min to 40 r/min within 10 min. The time for the mouse to fall from the lane was recorded for three consecutive times, and the longest fall time was recorded.

(2) Hindlimb Grip Test

The grip strength meter (Shanghai HaoHao, SH-10) was placed flat on the cage cover. The tail of the mouse was held to control the mouse’s body parallel to the direction of force. The hind paw of the mouse clasped the hook of the grip strength meter. Simultaneously, force was applied to the tail of the mouse in the opposite direction of the grip strength meter, until the hind paw of the mouse released. The test was conducted six times at 1-min intervals, and the average muscle strength value was calculated.

The tibialis anterior fixed in 4% PFA was dehydrated by gradient alcohol and embedded in OCT. After he tibialis anterior was sectioned (transverse and longitudinal sections of 10 μm), they were dried at 37°C for 2 h and used for subsequent experiments. For hematoxylin-eosin (H&E) Staining: after washing with PBS to remove OCT, the sections were stained with hematoxylin and eosin, and images were captured using an optical microscope to observe inflammatory infiltration in tibialis anterior. For immunofluorescence staining: the sections were blocked by 5% BSA, then incubated with anti-laminin (#ab11575, Abcam, United Kingdom), anti-CD68 (#38005, Signalway Antibody, United States), anti-NF200 (#N2912, Sigma, United States) and anti-α-BTX (#MKCK3522, Sigma, United States). The secondary antibody was Alexa Fluor^®488 (#ab150073, Abcam, United Kingdom) or Alexa Fluor^®594 (ab150116, Abcam, United Kingdom). The sections were sealed after staining with DAPI (#P0131, Beyotime, China). Images were captured using an SP5 confocal microscope (Leica, Wetzlar, Germany).

The fresh tibialis anterior were quickly embedded in OCT, then sliced (10 μm). The entire process was performed as quickly as possible, and the sections were stained with SDH staining reagent (#G2000, Solarbio, China). Images were captured using Zeiss microscope (Germany), and the proportion of positive muscle fibers was calculated.

The level of reactive oxygen species (ROS) in the tibialis anterior was determined using the DHE, a superoxide anion fluorescence probe (#S0063, Beyotime, China). The sections of fresh tibialis anterior (10 μm) were washed in PBS for 10 min, then incubated with DHE (10 μM) for 30 min at room temperature in the dark. The sections were washed three times with PBS for 5 min and sealed. All images were captured using a Zeiss microscope (Germany). The fluorescent intensity was analyzed using Image J software.

The tibialis anterior was prepared as a 1*1*3 mm³ cubic specimen and immersed in 2.5% glutaraldehyde followed by re-fixation in 1% osmium tetroxide. The section underwent rapid dehydration and was embedded in polymeric epoxy resin. Finally, the section was stained with 1% uranyl acetate and 0.3% lead citrate. The electron microscope section was photographed using a transmission electron microscope (HT7700, Hitachi, Tokyo, Japan).

Muscle tissue was lysed in a tissue lysis buffer containing proteinase inhibitor and phosphatase inhibitor. After centrifugation, the supernatant was collected and the protein concentration was determined using the BCA Protein Assay Kit (#P0010, Beyotime). An appropriate amount of protein was loaded onto 8%, 10%, or 12% one-step PAGE gels (#E303-11, Vazyme) according to the instructions. The separated proteins were then transferred to PVDF membranes activated by methanol. After blocking with 5% skim milk, the membranes were incubated with primary antibodies overnight at 4°C, followed by incubation with secondary antibodies at room temperature for 2 h on a horizontal shaker. The membranes were then washed and developed. The antibodies included: β-Tubulin (#ab6046), Cox2 (#ab15191), IL-1β(#ab254360), Nox2(#ab129068), Nox4(#ab109225), GPX1(#ab22604), Nrf2(#ab137550), Fbx32(#ab168372), MHC(#ab91506), Beclin1(#ab207612), ATG7(#ab133528), PGC1α(#ab191838), IL-6 (#ab9731), Sirt1 (#ab110304), BNIP3 (#ab10433), Goat Anti-Rabbit IgG H&L (HRP) (#ab205718), Goat Anti-Mouse IgG H&L (HRP) (#ab205719) from Abcam; CD68 (#38005), MuRF1 (#38580) from SAB; CD86 (#26903-1-AP), PTGES2 (#10881-1-AP), TNF-α (#60291-1-lg) from Proteintech; p-NF-κB (#3033S), p-Stat3 (Tyr705, #9145S), NLRP3(#30835S), Caspase1 (E9R2D, #83383S), Perk (#3192S), EIF-2α (#9722S), p-EIF-2α (#9721S), ATF-4 (#11815S), Chop (#2895S), Foxo3a (#12829S) from Cell Signaling Technology; p-Perk (#PA5-40294) from Invitrogen; LC3Ⅱ (#M186-3) from MBL.

All data are presented as mean ± standard deviation (SD). Statistical analysis was performed using GraphPad Prism 8.0.2 software. One-way ANOVA and Tukey’s analysis were used to detect differences and statistical significances. A p-value <0.05 was considered statistically significant.

After successful induction of diabetes using streptozotocin (STZ) in mice, we administered celecoxib treatment for 4 weeks. To evaluate the effects of celecoxib on motor function of diabetic mice, we measured the times to fall of rotarod test and hindlimb grip strength (Zhang et al., 2022). The results showed that compared to the control group, diabetic mice exhibited a significant decreased minute of rotarod test (Figure 1A) and a weakened hindlimb grip strength (Figure 1B), indicating impaired motor function and the possibility of diabetic sarcopenia (diabetes-induced skeletal muscle atrophy) in the muscle of diabetic mice. However, celecoxib showed a significant improvement in rotarod test and hindlimb grip strength in mice (Figures 1A,B). These results suggested that celecoxib significantly improves motor function of muscle in diabetic mice.

We further examined the effects of celecoxib on diabetic sarcopenia. The results showed that compared to the control group, the muscle weight of the tibialis anterior (TA) and the index of TA weight/body weight were significantly reduced in DM group (Figures 2A, B). However, the reduces were restored in the diabetic mice treated with celecoxib (Figures 2A, B). Subsequently, we conducted laminin staining on TA muscle and observed a significant reduction in cross-sectional area (CSA) of myofibers in diabetic mice compared to the control group, indicating occurrence of evident atrophy in STZ-induced diabetic muscle (Figures 2C, D). However, the decrease in CSA of myofibers was inhibited by celecoxib (Figures 2C, D). The CSA distribution of myofibers in diabetic muscle was shifted to the left, which was inhibited by celecoxib (Figure 2E), consistent with the laminin staining. Moreover, under conditions of diabetes, fragmentation and a decrease in the quantity and size of the NMJs could be observed (Francis et al., 2011). To examine the restoration of diabetic neuropathy by celecoxib, we performed immunofluorescent staining with neurofilament 200 (NF200) to label nerve fibers and α-bungarotoxin (α-BTX) to label motor endplates in TA. The results showed that compared to the control group, the morphology and number of motor endplates were overall smaller and fewer in DM group, whereas they improved in the celecoxib group (Figure 2F). These indicated that NMJs were damaged during diabetic-induced muscle atrophy, and celecoxib had a certain protective effect in NMJs of skeletal muscles affected by diabetes. The above results demonstrated that diabetic sarcopenia was successfully constructed, and celecoxib effectively alleviated STZ-induced diabetic sarcopenia and protected the NMJs damaged by diabetes.

Celecoxib, an anti-inflammatory drug (Bertagnolli et al., 2006), may alleviate diabetic sarcopenia by exerting anti-inflammatory effects. We validated the anti-inflammatory effect of celecoxib during diabetes-induced muscle atrophy. The results showed that compared with control group, diabetes-induced atrophic muscle showed more infiltration of inflammatory cells, which were improved by celecoxib (Figure 3A). Macrophages are the main effector cells in inflammation (Ji et al., 2022), so we performed immunofluorescence staining for the marker CD68 of macrophage. The results showed that the levels of CD68 was significantly enhanced in diabetes-induced atrophic muscle, and it was reduced after celecoxib treatment (Figure 3A). Western blotting results of CD68 were consistent with immunofluorescence staining (Figure 3B). Moreover, compared with control group, the level of CD86, M1 macrophage marker, was significantly upregulated in atrophic muscle, and it was inhibited in the celecoxib group (Figures 3B–D). Celecoxib can inhibit the production of inflammatory PGE2 by inhibiting COX2, thus exerting anti-inflammatory effects (Cruz et al., 2022). Next, we measured the levels of COX2 and PGE2. The results showed that, compared with control group, the levels of COX2 and PGE2 were significantly upregulated in atrophic muscle, while celecoxib inhibited their increases (Figure 3E). The occurrence of diabetes-related inflammation is usually accompanied by the release of a large number of pro-inflammatory factors, which activate inflammatory signaling pathways (Shen et al., 2022). We verified that, compared with the control group, the levels of pro-inflammatory factors, IL-6 and TNF-α, and the phosphorylation levels of NF-κB and Stat3 were significantly increased in atrophic muscle, indicating activation of the inflammatory signaling pathways during diabetes-induced muscle atrophy. After celecoxib treatment, the levels of IL-6, TNF-α, p-NF-κB, and p-Stat3 were significantly reduced, indicating that the inflammation were significantly inhibited by celecoxib (Figures 3E–K). In addition, the NLRP3-mediated inflammasome pathway plays an important role in muscle atrophy (Zhang et al., 2022). The results showed that compared with the control group, the levels of NLRP3, Caspase1, and IL-1β were significantly upregulated in atrophic muscle, while celecoxib inhibited the increases in NLRP3-mediated inflammasome pathway (Figures 3L–O). These results indicated that celecoxib inhibited inflammation during diabetes-induced muscle atrophy by inhibiting the TNF-α/NF-κB, IL-6/Stat3, and NLRP3 inflammasome pathways.

Chronic inflammation in metabolic diseases can further increase the production of reactive oxygen species (ROS), leading to oxidative stress (Newsholme et al., 2016). To investigate whether celecoxib has antioxidant effects during diabetes-induced muscle atrophy, we first measured the levels of ROS in the tibialis anterior of mice using DHE probe. The results showed that the level of ROS was significantly increased in atrophic tibialis anterior, while it was significantly reduced after celecoxib treatment (Figures 4A, B). To understand how celecoxib affects the levels of ROS, we further performed western blotting to detect the levels of NADPH oxidases 2 (Nox2) and NADPH oxidases 4 (Nox4), which promote ROS production, as well as the levels of glutathione peroxidase 1 (GPX1) and antioxidant-related transcription factor Nrf2, which protect the body from oxidative stress (Zorov et al., 2014). The results showed that the levels of Nox2 and Nox4 were significantly increased, while GPX1 and Nrf2 were significantly downregulated in atrophic muscles. Celecoxib significantly reversed these phenomena (Figures 4C–G). The above results indicated that celecoxib alleviated oxidative stress during diabetes-induced muscle atrophy.

Based on our above results, diabetes-induced atrophic muscles showed inflammation (Figure 3). The endoplasmic reticulum (ER) stress can trigger inflammatory pathways via the NF-κB pathway (Keestra-Gounder et al., 2016). To understand the effects of celecoxib on ER stress during diabetes-induced muscle atrophy, we used western blotting to detect the levels of p-Perk, p-EIF-2α, ATF4 and Chop, which related to ER stress. The results showed that the levels of p-PerK, p-EIF-2a, ATF4, and Chop were significantly upregulated in atrophic tibialis anterior, while celecoxib inhibited the upregulation of these (Figure 5). These results indicated that celecoxib inhibited the excessive activation of ER stress, possibly due to its anti-inflammation, thereby protecting against diabetes-induced muscle atrophy.

The inflammation can activate the ubiquitin-proteasome system by FoxO and downstream E3 ubiquitin ligases, and eventually leading to muscle atrophy in diabetes (Shen et al., 2022). To investigate the effect of celecoxib on the ubiquitin-proteasome system during diabetes-induced muscle atrophy, we identified the levels of Foxo3a, two E3 ubiquitin ligases (Fbx32 (also named Atrogin-1) and MuRF1) and myosin heavy chain (MHC) by western blotting. The results showed that compared with the control group, the levels of the Foxo3a, Fbx32 and MuRF1 were significantly upregulated in atrophic tibialis anterior, while celecoxib significantly reversed these phenomena (Figures 6A–D). Moreover, the levels of MHC were significantly downregulated in atrophic tibialis anterior, and celecoxib inhibited the downregulation (Figures 6A, E). These results indicated that celecoxib attenuated the activation of the ubiquitin-proteasome system during diabetes-induced muscle atrophy and has a protective effect on muscle.

The autophagy lysosome system is also crucial for maintaining skeletal muscle homeostasis, but excessive activation of autophagy promotes muscle atrophy (Yin et al., 2021). FoxO3 regulates a number of essential autophagy genes, including Bcl2 family member BNIP3, LC3 II and ATG7 (Yin et al., 2021; Zhang et al., 2022). Beclin1, a component of VPS34 complex that regulates autophagy process, plays an important role in the autophagy lysosome system (Zhang et al., 2022). To determine the effects of celecoxib on autophagy lysosome system in diabetes-induced muscle atrophy, we examined the levels of BNIP3, Beclin1, LC3 II and ATG7 in tibialis anterior. Compared to control group, the levels of BNIP3, Beclin1, ATG7, and LC3 II were significantly upregulated in atrophic muscle, while celecoxib inhibited the upregulation of these autophagy-related proteins (Figures 7A–E). These results suggested that celecoxib can alleviate diabetes-induced muscle atrophy by attenuating the excessive activation of autophagy lysosome system.

Diabetes affects the mass, strength, and function of skeletal muscles through disturbances in energy metabolism and mitochondrial dysfunction (Bianchi and Volpato, 2016). To evaluate the effects of celecoxib on mitochondria and metabolism during diabetes-induced muscle atrophy, we first observed the morphological changes of mitochondria using electron transmission microscopy. Compared to control group, the atrophic tibialis anterior showed disordered muscle filament arrangement and increased number of mitochondrial vacuoles, consistent with the autophagy results (Figure 7). However, celecoxib treatment group exhibited relatively orderly muscle filament arrangement and relatively intact mitochondria (Figure 8A). Furthermore, to understand the effects of celecoxib on mitochondrial biogenesis, we performed western blotting to detect the levels of Sirt1 and PGC1α. The results showed that compared to control group, the atrophic muscle exhibited significant downregulation of the levels of Sirt1 and PGC1α, while celecoxib significantly inhibited their downregulation (Figures 8B–D). Succinate dehydrogenase (SDH), one of the markers of mitochondrial function involved in the tricarboxylic acid cycle, provides energy for the body (Porporato et al., 2018). The results showed that compared to control group, the number of SDH-positive fibers had a significantly reduced in atrophic muscle, which was inhibited by celecoxib (Figures 8E, F). These results suggested that celecoxib could protect mitochondrial integrity, enhance mitochondrial biogenesis, maintain mitochondrial function during diabetes-induced muscle atrophy.