1. Introduction
Skeletal muscle atrophy is a common complication associated with both aging and a number of health problems, including chronic obstructive pulmonary disease, chronic heart failure, cancer, and inflammation. The underlying mechanisms of skeletal muscle atrophy involve the upregulation of specific proteins and inflammation through the NF-κB signaling pathway, which indirectly or directly disrupts muscle regeneration, thereby leading to skeletal muscle atrophy [
1]. Although skeletal muscle atrophy is not life threatening, it can contribute to the development of complications such as osteoporosis, thrombosis, and pressure ulcers, all of which can indirectly increase morbidity and mortality rates.
Currently, the most common treatment methods for skeletal muscle atrophy are physical rehabilitation therapy and drug therapy. Drug therapy involves the use of medications such as anabolic drugs, enzyme inhibitors, and anti-inflammatory drugs, including androgen/androgen receptor modulators, ghrelin, β-adrenoceptor agonists, Cox2 inhibitors, histone deacetylase inhibitors, and phosphodiesterase inhibitors [
2]. However, some of these drugs can have side effects, such as prostatitis, blood embolism, gastrointestinal complications, dizziness, and chronic diseases. Therefore, exploring the potential of natural compounds for the treatment of skeletal muscle atrophy may be a promising approach. Numerous studies have demonstrated that natural compounds can prevent or treat skeletal muscle atrophy. Examples of such natural compounds are resveratrol, salidroside, imperatorin, and ursolic acid [
2].
The seed of the cumin plant is a widely used spice worldwide and constitutes the main ingredient in curry powder. The major volatile substances found in cumin seeds are terpenoids, cymene, and cumin aldehyde [
3]. Cumin seeds also contain flavonoids, which are known for their antioxidant activity [
4]. Miah et al. indicated that cumin seed powder supplementation in rats fed with a high-fat diet markedly suppressed oxidative stress in both their plasma and livers [
5]. Cumin oil, which can be extracted from cumin seeds, has been used as an ingredient in some cosmetic products [
6]. Cumin aldehyde, another extract from cumin seed, has demonstrated the ability to scavenge the superoxide anion [
7]. Cumin oil has been shown to possess anti-inflammatory properties and has been found to inhibit the mRNA expression of interleukin (IL)-1β and IL-6, which are inflammatory factors, in lipopolysaccharide (LPS)-induced inflamed RAW264.7 macrophages. This effect occurs through the modulation of the JNK, ERK, and NF-κB signaling pathways [
8]. Langen demonstrated that high levels of tumor necrosis factor (TNF)-α and IL-1β inhibited the differentiation of cultured myoblasts into myotubes through the activation of the NF-κB transcription factor [
9]. This was also found to have cardioprotective, hypotensive, and lipid-lowering activities [
5,
10]. In addition, cumin exerts a hypoglycemic effect on streptozotocin induced diabetic rats [
11,
12]. However, the low water solubility, volatility, and cytotoxicity of CE impede its potential pharmaceutical applications, particularly for treating chronic diseases such as diabetes.
Diabetes is a prevalent chronic disease that affects a large proportion of the global population. Physiologically, insulin plays a crucial role in regulating skeletal muscle protein via the AKT/mTOR pathway. In health conditions where blood glucose levels are high and where insulin levels are low, protein synthesis is inhibited and protein breakdown is promoted. This imbalance leads to skeletal muscle atrophy and rapid weight loss [
13]. Moreover, diabetes can force the immune system to secrete cytokines, which trigger various inflammatory responses. Prolonged chronic inflammation has been shown to increase the occurrence of catabolic diseases. Among the inflammatory cytokines, TNF-α is known to trigger excessive inflammation in the body, leading to skeletal muscle atrophy [
14]. Skeletal muscle atrophy is often characterized by a decrease in the cross-sectional area of muscle fiber, an increase in protein catabolism, and subsequent loss of muscle mass and strength [
15]. Skeletal muscle atrophy can be due to either increased protein degradation or decreased protein synthesis. Sustained inflammation, high levels of oxidative stress, and upregulated levels of pro-inflammatory cytokines such as TNF-α and IL-1β contribute to the degeneration of healthy muscle cells, resulting in skeletal muscle atrophy [
16]. Furthermore, the elevation of inflammatory factors such as TNF-α, IL-1β, and IL-6 has been observed in human and chronic disease animal models of skeletal muscle atrophy [
17,
18].
In the treatment of diseases with drugs, the location, timing, and profile of drug release must be controlled to minimize toxicity and increase the efficacy of the drug. Nanoparticle encapsulation of drugs has emerged as a promising approach in the field of medicine [
19]. Nanoparticles, with sizes ranging from 1 to 100 nm, offer advantages such as high drug loading capacity and specific targeting of cells owing to their size-dependent effect and intracellular uptake [
20]. Biopolymeric nanoparticles—including nanosized hyaluronan, chitosan, and gelatin—are particularly favored as carriers in drug delivery systems because of their biocompatible and biodegradable properties [
21]. In our previous studies, we have demonstrated that encapsulating heteronemin by using chitosan and hyaluronan nanoparticles can enhance the antitumor effects of heteronemin [
22]. Chitosan, a natural cationic polymer, has demonstrated properties such as absorption enhancement through mucosa, controlled drug release, and bioadhesion [
23,
24]. By considering the presence of the substantial amounts of beneficial reports of cumin and the drug carrier using chitosan, the current study was designed to investigate the anti-inflammatory effects of CE and CE-encapsulated chitosan nanoparticles (CECNs, which were fabricated using the electrostatic field system method) on C2C12 cells and the ameliorative effect on skeletal muscle atrophy in STZ-induced diabetic rats.
2. Materials and Methods
The dried cumin used in this study was purchased from a local store and was authenticated by Prof. Dr. Lin Li-wei. A voucher sample with the identification number ISU-MCMM-201 was preserved for future reference in the School of Chinese Medicines for Post-Baccalaureate, I-Shou University. STZ was purchased from Sigma (St. Louis, MO, USA), and 3-4,5-Dimethylthiazol-2-yl-2,5-diphenyltetrazolium bromide (MTT), Dulbecco’s Modified Eagle Medium, fetal bovine serum, streptomycin, and penicillin were obtained from Gibco (Waltham, MA, USA). All chemicals used in the present study were of reagent grade. Animal experiments conducted in this study were approved by the Institutional Animal Care and Use Committee of I-Shou University, Kaohsiung, Taiwan (IACUC-ISU-110–052, approval date: 4 March 2022).
2.1. Isolation and Characterization of CE
CE was extracted using the water distillation method. This procedure involved grinding 200 g of cumin seeds into a powder and then placing this powder in a rotary bottle. To this powder, 600 mL of ddH
2O was then added. The extraction was conducted using a rotary evaporator (EYELA 1300VF, Tokyo, Japan) under the following conditions: vacuum pressure, 200 hPa; rotary speed, 20 rpm; heating temperature, 65 °C; condensation temperature, 4 °C; and extraction period, 3 h. The obtained CE was further treated with anhydrous sodium sulfate to remove residual water, and the resultant extract was stored at 4 °C for further experiments. Fourier-transform infrared (FTIR) spectroscopy and
1H nuclear magnetic resonance (NMR) analyses were performed to identify the CE, as compared with the pure cumin purchased from Sigma. FTIR spectroscopy was employed to characterize the characteristic peaks of CE (
Figure 1a). The yield of extraction was determined using the following formula:
High-performance liquid chromatography (HPLC, Agilent 1100, Santa Clara, CA, USA) equipped with a Zoxbax-C18 column was employed to determine the chemical composition of the CE. The mobile phase consisted of a mixture of sodium sulfate, acetonitrile, and methanol (90:190:20, v/v), and the flow rate was set at 1 mL/min. Detection was performed at 326 nm using an ultraviolet–visible detector. The characteristic peaks of CE were further analyzed through FTIR spectroscopy with potassium bromide disks (Jasco 4700, Tokyo, Japan), and the NMR spectra were recorded using a Bruker AM-400 spectrometer (Bruker Avance III HD 600 MHz, Karlsruhe, Germany).
2.2. Fabrication of CECNs
Chitosan nanoparticles were prepared by following the methodology described in a previous study [
25]. In brief, an EFS was established using two parallel copper plates (18 cm × 6 cm) spaced 2 cm apart. The electrical field strength between the plates was generated using a direct-current power supply. The EFS was constructed in a thermal chamber for temperature control and safety purposes. To prepare the CECNs, 1 mL of chitosan solution (0.2 mg/mL) was carefully mixed with 150 μL of CE (0.5 M). This mixture was then transferred onto a petri dish and placed between the two plate electrodes in the EFS. The following conditions were set for CECNs fabrication in the EFS: temperature, 17 °C; applied electrical field strength, 2.5 kV/cm; reaction time, 5 h; and crosslinking reagent, 0.001 N NaOH. The sizes of the chitosan nanoparticles and CECNs were determined by randomly sampling approximately 60 individual nanoparticles through transmission electron microscopy (TEM). To assess the entrapment efficiency (EE) of CE, 1 mL of the solution containing the prepared CECNs was centrifuged at 10,000 rpm for 10 min. The amount of nonencapsulated CE in the supernatant was measured through HPLC. A standard concentration curve of CE was established to determine the amount of CE encapsulated in CECNs. Finally, the EE was calculated using the following formula: [(total amount of CE − amount of nonencapsulated CE)/total amount of CE] × 100%. Next, the in vitro release of CE from CECNs was evaluated by adding 100 μL of CECNs to a 1.5 mL microcentrifuge tube, which was then placed in a shaker set at 37 °C and 40 rpm. At predetermined time points, the sample was centrifuged at 10,000 rpm for 10 min. The amount of nonencapsulated CE in the supernatant was determined through HPLC. Independent experiments were performed three times in triplicate. The in vitro release rate was calculated as follows: [(total amount of CE − amount of residual CE)/total amount of CE] × 100%.
To determine the volatility of CE and CECNs, 1 mL of CE or CECNs solution was added to a 48-well plate, and this plate was placed in a 37 °C, 5% CO2 incubator. At predetermined time points, the sample was centrifuged at 10,000 rpm for 10 min. The amount of residual CE in the supernatant was then determined through HPLC. The volatility of CE or CECNs was calculated as follows: 1 − [(total amount of CE − amount of residual CE)/total amount of CE] × 100%.
2.3. Cytotoxicity of CE and CECNs in Relation to C2C12 Cells
To assess the cytotoxicity of CE and CECNs in relation to C2C12 cells (ATCC, VA, USA), the MTT assay was performed. The C2C12 cells were seeded on a 96-well plate at a density of 1.2 × 104 cells/well and were allowed to attach for 24 h. Subsequently, these cells were treated with multiple concentrations of CE (20–100 μM) and CECNs (0.6–1.4 mM). After 24 h of treatment, 10 μL of MTT solution (5 mg/mL) was added to each well, and the plate was then incubated for an additional 4 h. To dissolve the formazan precipitate, DMSO was added to each well. The absorbance of the formazan solution was measured at 450 nm using a multiplate reader (Thermo Scientific, Waltham, MA, USA).
2.4. Evaluation of the Anti-Inflammatory Effects of CE and CECNs on LPS-Induced C2C12 Cells
To evaluate the anti-inflammatory effects of CE and CECNs on LPS-induced C2C12 cells, C2C12 cells (1.2 × 104 cells/well) were seeded on a 24-well plate for 24 h and preincubated with 0.3 mg/mL LPS for 6 h. These cells were then treated with multiple concentrations of CE (20, 30, and 40 μM) and CECNs (0.6, 0.7, and 0.8 mM). After incubation for 24 h, the medium was collected and centrifuged at 1000× g for 20 min to obtain cell-free supernatant, which was then used to perform the enzyme-linked immunosorbent assay (ELISA) for IL-6, IL-1β, and TNF-α. The levels of IL-6, IL-1β, and TNF-α were quantified using the Elabscience® Mouse IL-6, IL-1β, and TNF-α ELISA Kit (Minneapolis, MN, USA), and absorbance was measured at 450 nm according to the manufacturer’s protocol.
2.5. Establishment of the STZ-Induced Diabetic Rat Model
Animal experiments were performed over 10 weeks (
Figure 1b). Diabetes was induced in rats by using STZ. A 1% STZ working solution (Sigma-Aldrich, St. Louis, MO, USA) was prepared in 50 mmol citrated buffer solution (pH 4.2). A total of 10 male rats were used to establish the diabetic rat model, whereas two rats served as the healthy control group. The rats were intraperitoneally injected with 200 μL of STZ solution (65 mg/kg body weight). After the injection, each rat was left untreated for 4 weeks. During this 4-week period, the rats’ blood glucose levels were measured at p.m. 14:00 every 3 days using EasyTouch
® (Yuan Yu Bio-Tech Co., Ltd., Taoyuan, Taiwan). Blood glucose levels of >16.0 mmol/L were considered indicative of diabetes [
26]. The rats were anesthetized using Zoletil
® (intraperitoneal administration of 40 mg/kg of tiletamine with 50 mg/kg of zolazepam,) and xylazine (10 mg/kg) and randomly allocated into the following groups (n = 2 per group): Group A: STZ-induced diabetic rats without treatment, serving as a negative control group; Group B: rats treated with 200 μL of CE (2.5 mM) every 2 days for 3 or 6 weeks; and Group C: rats treated with 200 μL of CECNs (2.5 mM) every 2 days for 3 or 6 weeks. At weeks 3 and 6 after treatment, one rat was randomly selected from each group and sacrificed. The rectus femoris muscle, liver, pancreas, kidney, and serum of the sacrificed rats were harvested for histopathological and insulin analyses.
2.6. Histopathological Analyses
At weeks 3 and 6 after treatment, the sacrificed rats’ rectus femoris muscle, liver, pancreas, and kidney tissues were harvested and fixed in 10% neutral-buffered formalin. The resulting samples were then dehydrated in graded ethanol solution, cleared in xylene, embedded in paraffin blocks, and cut into 5 μm thick sections. Histopathological examination of the tissue samples was performed through hematoxylin and eosin (H&E) staining.
2.7. Forelimb Grip Strength Measurement
As shown in
Figure 1c, the grip strength of the rats was assessed using a standardized method, where each rat was lifted by the tail and encouraged to grasp a rigid metal bar attached to a digital force gauge (AMETEK
® Measurement & Calibration Technologies, Inc., Miami, FL, USA). Subsequently, the rat was gently pulled backward by the tail until it released the metal bar. The tension reading displayed on the digital force gauge just before the rat released the metal bar was recorded as the grip strength. This test was performed three consecutive times with a 3 min rest after each round. Each rat’s grip strength measurement is expressed as mean ± standard error in this paper.
2.8. Insulin Assay
To assess the function of the rat pancreas, a blood biochemical parameter, namely insulin, was assayed. Rat serum samples were isolated from whole blood sample and subjected to insulin biochemical analysis according to the insulin manufacturer’s instructions (Elabscience® Insulin ELISA kit, Houston, TX, USA).
2.9. Immunohistochemistry (IHC) Analysis
For (IHC) analysis, the sections were deparaffinized and rehydrated using graded concentrations of ethanol and then deionized water. The sections were incubated in a hydrogen peroxide blocking solution for 10 min and then washed with PBS. The sections were subjected to heat treatment (at 95 °C) in 0.01 M sodium citrate buffer with Tween 20. Each section was incubated with ImmunoBlock (PBS, pH 7.6, with 0.5% bovine serum albumin) at room temperature for 20 min, followed by washing with PBS. Next, they were incubated with rabbit anti-mouse IL-6 (Novus Biologicals, LLC, Centennial, CO, USA) at 4 °C overnight and then with Mouse/Rabbit Probe HRP Labeling solution at 25 °C for 30 min. Finally, 3,3′-diaminobenzidine was applied for 10 min. The intensity of the brown color, indicating the presence of IL-6 was semi-quantified using ImageJ software (Version 1.50; National Institutes of Health, Bethesda, MD, USA).
2.10. Statistical Analysis
In this paper, all data are expressed as mean ± standard error of the mean. Differences among groups were analyzed using a one-way analysis of variance followed by Tukey’s multiple comparison test. A p value of <0.05 was considered statistically significant. All statistical analyses were performed using SPSS (version 20.0).
4. Discussion
The presence of elevated extracellular matrix levels in diabetic tissue samples has been reported in a number of studies. Abnormal extracellular matrix (ECM) turnover can interfere with growth factor signaling and thus can impede myogenesis. In addition, remodeling of the ECM in diabetic mice is advised on the basis of evidence of increased collagen presence, which affects satellite cell functionality and migration, both of which in turn influence muscle growth, maintenance, and repair [
28,
29,
30]. Consistent with the findings of previous studies, in the present study, increased collagen deposition was observed between the muscle fibers of STZ-induced diabetic rats, as was the aggregation of muscle cells and inflammatory cells located between muscle. These changes are associated with muscle atrophy and a decrease in grip strength, as evidenced by the narrowed perimysium and endomysium space between muscle fibers in this study. Importantly, after treatment with CE and CECNs, both the area of the perimysium and endomysium between muscle fibers and the area of muscle fibers (size) had significantly increased, indicating the amelioration of muscle atrophy. Furthermore, the significant recovery of grip strength to more than 80% of that of the healthy control rats indicated the partial recovery of muscle function (
Figure 7). However, interpreting muscle biopsies can be challenging, because morphological changes alone may not be sufficient to assign a specific diagnosis; diagnosing muscle changes or diseases often requires a combination of multiple changes observed in a biopsy.
Prolonged exposure to inflammation creates a state of chronic inflammation which involve complicated processes of cytokine release and the emergence of a high-oxidative-stress status, which can cause irreversible cellular and tissue damage. In turn, excessive oxidative stress caused by reactive oxygen species (ROS) can trigger an inflammatory response to lead to further oxidative stress and chronic inflammation. Studies indicated that persistent inflammation caused extreme pro-inflammatory cytokines production and linked chronic inflammation to such as type 2 diabetes and arthritis diseases [
31]. Atrophy is a condition that affects skeletal muscle and is characterized by the loss of muscle mass and muscle function. The mechanisms underlying muscle atrophy are complex and multifaceted. TNF-α is the dysregulation of inflammatory cytokines playing a crucial role behind muscle atrophy. IL-6 is also a classic pro-inflammatory cytokine, which possess ancillary function of influencing metabolism. In a rodent study, IL-6 was reported to induce atrophy via downregulation of ribosomal S6 kinase phosphorylation [
32,
33]. This study revealed that CE and CECNs was successful in decreasing the pro-inflammatory cytokines IL-6, TNF-α and IL-1β levels in in vitro cellular tests (
Figure 4c,d). Furthermore, treatment with CE and CECNs decreased the IL-6 level to ameliorate skeletal muscle atrophy and, significantly improved the grip strength in STZ-induced diabetic rats (
Figure 7).
Streptozotocin (STZ) is a naturally occurring compound that specifically targets insulin-containing β cells, causing pancreatic β-cell destruction and is widely used to produce an animal model of diabetic mellitus. In this present study, we utilized a multiple, low-dose STZ approach only partially damaged pancreatic islets, triggering inflammatory responses that caused gradual loss of β-cell activity that resulted in insulin deficiency and hyperglycaemia. This response more closely resembled type I diabetes in pathogenesis and morphologic changes than the single, high-dose STZ protocol. In addition, different sensitivity to STZ-induced diabetes has been reported in male and female mice [
34]. Kim et al. indicated that fasting blood glucose was about 50% higher in C57BL/6J male mice than female mice at 5 weeks after STZ injection [
35]. Another study in C57BL/6 mice reported similar results, i.e., blood glucose was about 90% higher in male mice than female mice [
36]. These data indicated that male mice were more sensitive to STZ and tended to develop greater hyperglycaemia. In other words, male mice were more susceptible to STZ to induce diabetes. Indeed, both sexes should be studied to elucidate the determinants of these fundamental biological sex differences from genes to hormones. Further, the identification of sex differences in metabolic function and disease would provide knowledge to allow the development of relevant sex-based therapeutic avenues for diabetes. When considering the efficacy of STZ-induction to diabetes, we utilized male rat as animal target in the present study. As shown in
Figure 9, the abnormal pathological conditions originated in STZ-induced diabetic rats, such as shrank pancreatic islet and decreases in the number and size of pancreatic islet in the pancreas, were ameliorated by oral administration of CE and CECNs. Furthermore, serum insulin secretion was slightly increased, and blood glucose levels were lowered under the action of increased insulin levels. These findings indicated that CE and CECNs exerted a reparative effect on the pancreas in the STZ-induced diabetic rats.
In our previous studies, we have successfully produced chitosan and hyaluronan nanoparticles to encapsulate or aggregate poorly soluble drugs in order to enhance the therapeutic efficacy of these drugs [
37]. In particular, the positive [H
+] charge of chitosan nanoparticles, generated in an acid solution environment prepared using NaOH gelation agent during the preparation process, plays a crucial role in protecting the encapsulated drug from gastric acid degradation during oral administration. Aggregated chitosan nanoparticles have the ability to disintegrate in the alkaline environment of the intestine, thereby facilitating the controlled release of the encapsulated drug (
Figure 10). On the basis of these unique characteristics of chitosan nanoparticles, CECNs were observed to exhibit a more marked therapeutic effect than CE alone. This effect was confirmed on the basis of histological analysis, significant changes in biochemical test results and improvements in muscle grip strength.
This study revealed many novel aspects of cumin extract effect on restoration of muscle grip strength and protection of pancreas organ in STZ-induced diabetic rats. However, some tests of this research, such as measurement of the anti-oxidative capability of cumin extract, were not performed, as we believed that the antioxidant properties of cumin extract had already been elucidated and proved [
38]. This study suggests that the oral administration of cumin extract (CE and CECNs) may be responsible for the lowering of inflammation response, improved muscle grip strength, a reduction of damage to the pancreas, restoring its morphological phenotype, increasing serum insulin levels, and lowering blood glucose levels in STZ-induced diabetic rats. However, further research is suggested in order to obtain the benefits of cumin supplementation in a clinical setup.