All animals were treated according to the guidelines for the Care and Use of Laboratory Animals (23), and the study was approved by The Committee on The Ethics of Animal Experiments of Xinjiang Medical University (approval no. XM2017MD). In total, 40 male C57BL/6 mice (weight, 18–22 g; age, 6 weeks) were purchased from The Experimental Animal Center of Nanjing Medical University (certificate of conformity no. SCXK JS 2016-0002). Mice were acclimatized for 7 days prior to the start of the study. Mice were provided access to drinking water and standard rodent chow ad libitum. The temperature of housing environment was set as 22±2°C. The relative humidity was set at 60±10% under a 12-h light/dark cycle. The mice were randomly divided into four groups: i) Control mice (Con); ii) HFD mice (Veh); iii) HFD mice treated with 10 mg/kg CA (CA-L); and iv) HFD mice treated with 20 mg/kg CA (CA-H). The HFD protocol was performed according to a previous study (24). CA was purchased from Changsha Yaying Biotechnology Co., Ltd. CA solution was prepared according to a previous study (18). Mice were treated with CA via gavage for 9 weeks following a 6-week period of HFD. The caloric intake was calculated by subtracting the fecal caloric excretion (fecal caloric content × fecal excretion) from the dietary caloric intake (diet caloric content × food intake). After 15 weeks, all mice were fasted for 12 h. The blood of anesthetized mice was collected via retro-orbital puncture. Subsequently, body and liver weights were measured, and the brain, pancreas and whole liver tissues were collected at 4°C. The tissues were frozen in liquid nitrogen and stored at −80°C. For histology, the tissues were fixed in 10% neutral buffered formalin (saturated aqueous formaldehyde from Thermo Fisher Scientific, Inc., buffered to pH 6.8–7.2 with 100 mM phosphate buffer) for 24 h at 25°C.

According to the metabolic syndrome diagnostic criteria provided by the World Health Organization (25), insulin resistance (IR) is required to diagnose metabolic syndrome. In addition, to diagnose metabolic syndrome, two other criteria among the following five are required: Obesity (waist/hip ratio: >0.90 for males, >0.85 for females; or body mass index >30 kg/m²), hyperglycemia, dyslipidemia [triglycerides (TG): >150 mg/dl or high-density lipoprotein cholesterol: <35 mg/dl for males, <39 mg/dl for females], hypertension (≥140/90 mmHg) and microalbuminuria. In the present study, IR, TG and hyperglycemia were selected to assess metabolic syndrome in mouse models. After 15 weeks, blood was collected by retro-orbital puncture method. Additionally, the serum was obtained from blood samples via centrifugation with 2,000 RPM (800 × g) for 15 min at 4°C. Then, the serum was transferred to a 5-ml centrifuge tube, which was placed on the glacial table. Serum levels of TG (cat. no. A110-2-1) and total cholesterol (TC; cat. no. A111-2-1) were tested with biochemical kits (Nanjing Jiancheng Taihao Biotechnology Co., Ltd.). Glucose (cat. no. F006-1-1) and insulin (cat. no. H203) levels were also measured using biochemical kits (Nanjing Jiancheng Taihao Biotechnology Co., Ltd.).

After 15 weeks, the serum was obtained via centrifugation of blood samples with 2,000 RPM (800 × g) for 15 min at 4°C. ELISA kits, including TNF-α (cat. no. MTA00B), IL-1β (cat. no. MLB00C) and IL-6 (cat. no. D6050) was used to identify the serum concentrations of these inflammatory cytokines. The detailed steps were followed according to the manufacturer's protocols (R&D System, Inc.). The standard curve for each cytokine was calculated; the concentration of the antigens was determined at 450 nm.

Histopathological evaluation was performed on the liver, pancreas and brain tissues of mice. Samples were fixed with 10% formalin buffer for 24 h at 25°C. Subsequently, the pretreated samples were embedded in paraffin and sliced (3–5 µm). H&E staining was performed according to the previous method (26). Briefly, samples were stained with hematoxylin for 10 min and with eosin for 1 min (both at room temperature) to establish the diagnosis areas. After H&E staining, histopathological alterations were observed using a light microscope. For each sample, three randomly selected microscopic fields of view were used to calculate the pathological scores. The microscopic fields were scored blindly using a scale from 0 (normal) to 5 (highly destructive pathology). The mean of the three values was used to calculate an overall pathological score, as previously described (27).

Additionally, immunohistochemical staining was performed on 3–5-µm thick paraffin-embedded tissue sections. Sections were deparaffinized in xylene, hydrated using a graded alcohol series, and washed with TBS for 10 min and distilled water for a further 10 min. Endogenous peroxidase activity was blocked with 3% v/v H₂O₂ in water for 5 min. Antigen retrieval was performed for all antibodies by placement of the sections in citrate buffer and heating in a microwave oven for 15 min. Sections were treated with 25 ml blocking buffer for 1 h at room temperature. The treated sections were separately incubated with primary antibodies for TNF-α (1:200; cat. no. 11948) and IL-1β (1:400; cat. no. 12703) at 4°C overnight (both Cell Signaling Technology, Inc.). Then, all treated sections were incubated with the biotin-conjugated secondary antibody (1:800; cat. no. ab6720; Abcam) for 30 min at room temperature. The standard streptavidin-biotin-peroxidase complex method was performed using an LSAB System Universal kit (Dako; Agilent Technologies, Inc.) for 10 min, 3,3′-diaminobenzidine solution was used as a chromogen for 5 min, and all sections were counterstained with Mayer's haematoxylin for 1 min and mounted; all reactions were performed at room temperature. The species were observed under a light microscope (Eclipse 80i; Nikon Corporation). The percentage of IL-1β- and TNF-α-positive cells was calculated using the inForm cell analysis software (version 2.0.4743.16069; PerkinElmer, Inc.). The histopathological examination method was performed as previously described (28).

Total protein from different groups was extracted using the T-PER Tissue Protein Extraction Reagent kit (Thermo Fisher Scientific, Inc.). Protein concentration was determined using a bicinchoninic protein assay kit (Thermo Fisher Scientific, Inc.). Proteins (50 µg/lane) were separated by 10% SDS-PAGE. Subsequently, proteins were transferred to PVDF membranes, which were blocked with TBS containing 0.05% Tween-20 (TBS-T), supplemented with 5% skim milk (Sigma-Aldrich; Merck KGaA) at room temperature for 2 h on a rotating shaker. Membranes were subsequently washed with TBS-T. The primary antibodies used were as follows: IL-1β (1:100; cat. no. PA1351; Boster Biological Technology), IL-18 (1:500; cat. no. RP1017; Boster Biological Technology), TNF-α (1:500; cat. no. RP1000; Boster Biological Technology), IKKα (1:1,000; cat. no. 11930; Cell Signaling Technology, Inc.) and phosphorylated (p)-IKKα (1:1,000; cat. no. 2697; Cell Signaling Technology, Inc.), IκBα (1:500; cat. no. 4814; Cell Signaling Technology, Inc.) and p-IκBα (1:500; cat. no. 9246; Cell Signaling Technology, Inc.), NF-κB (1:1,000; cat. no. 8242; Cell Signaling Technology, Inc.) and p-NF-κB (1:1,000; cat. no. 3033; Cell Signaling Technology, Inc.), glial fibrillary acidic protein (GFAP; 1:2,000; cat. no. 3670; Cell Signaling Technology, Inc.), neuronal nuclei (Neu-N; 1:1,500; cat. no. 24307; Cell Signaling Technology, Inc.), ionized calcium-binding adapter molecule 1 (Iba-1; 1:2,000; cat. no. ab15690; Abcam), Bax (1:1,000; cat. no. ab32503; Abcam), Bcl-2 (1:1,500; cat. no. ab182858; Abcam), MMP-9 (1:100; cat. no. AF909; R&D Systems), Caspase-3 (1:400; cat. no. ab13585; Abcam), and GAPDH (1:1,000; cat. no. ab8245; Abcam). The antibodies were diluted in TBS-T and were incubated with the membranes at 4°C overnight. Subsequently, the membranes were washed with TBS-T followed by incubation with ImmPRESS^® HRP Universal Antibody Polymer Detection Kit (1:2,000; cat. no. MP-7500-15; Maravai LifeSciences) for 1 h at room temperature. The protein bands were detected using an ECL western blot detection kit (Thermo Fisher Scientific, Inc.). Western blot bands were observed with an ECL Western Blotting Analysis system (cat. no. RPN2108; GE Healthcare Life Sciences) and exposed to X-ray films (Kodak). Image Studio Lite Western Blot Analysis Software version 3.1 (LI-COR Biosciences) was chosen to perform pixel quantification of the images.

Total RNA was isolated from individual mouse brains using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.). The total RNA was used to evaluate the relative mRNA expression levels of IL-1β, IL-18, TNF-α, Bax, Bcl-2, MMP-9, Caspase-3 and GAPDH. The reverse transcribed cDNA was synthesized using the SuperScript First-Strand Synthesis kit (Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. The primers used for qPCR are listed in Table I. Primer sequences were verified with NCBI primer blast tool to avoid non-specific annealing (https://www.ncbi.nlm.nih.gov/tools/primer-blast/). All primers were obtained from Sangon Biotech Co., Ltd. Reactions were performed in a total volume of 20 µl, and consisted of 10 µl 2X SYBR Green PCR master mix (cat. no. 4309155; Applied Biosystems; Thermo Fisher Scientific, Inc.), 1 µl forward primer (10 pmol), 1 µl reverse primer (10 pmol), 1 µl cDNA template and 7 µl double distilled water. The thermocycling conditions were as follows: 94°C for 3 min, then 40 cycles of 95°C for 15 sec and 60°C for 25 sec. BioRad iCycler iQ detection system was used in this study (Bio-Rad Laboratories, Inc.). GAPDH was used as the reference gene. Normalization and fold change for each gene were calculated using the 2^−∆∆Cq method (29).

All experiments were repeated three times. Data are presented as the means ± SEM. Treated cells, tissues and the corresponding controls were compared using GraphPad Prism (version 6.0; GraphPad Software, Inc.). Multiple groups were compared using one-way ANOVA. Differences between groups were calculated using the Student-Newman-Keuls post hoc test. P<0.05 was considered to indicate a statistically significant difference.

Metabolic disease has previously been reported to be associated with neurodegenerative disease via the inflammatory response (30). A previous study suggested that HFD could induce metabolic diseases in rodents, causing type II diabetes characterized by IR (31). In the present study, animal models of metabolic syndrome were established by feeding mice a HFD. Mouse body weight in the Veh group was significantly higher than in the Con group (Fig. 1A; P<0.01). Conversely, high concentrations of CA could significantly reduce body weight compared with the Veh group (P<0.05). The present results suggested that CA induced body weight loss in a dose-dependent manner. Similarly, the liver weight in different groups was also measured. Liver weight increased significantly in the Veh group compared with in the Con group (P<0.05). Moreover, high CA was sufficient to significantly reduce the liver weight compared with the Veh group (P<0.05). The present results suggested that CA induced liver weight loss in a dose-dependent manner (Fig. 1B). In addition, the daily food intake in the Veh group was significantly lower than in the Con group (P<0.05). However, the daily food intake in the Veh group was restored following treatment with a high dosage of CA (P<0.05; Fig. 1C). Moreover, the daily caloric intake in the Veh group was significantly higher compared with that in the Con group (P<0.01). This effect was reversed by CA treatment (P<0.05; Fig. 1D). In addition, serum levels of TG, TC and insulin were analyzed. The present results suggested that TG, TC and insulin in the Veh group were significantly higher than in the Con group (P<0.001; Fig. 1E-G). Notably, CA treatment was able to significantly decrease the serum levels of TG, TC and insulin. In addition, the serum glucose level in the Veh group was significantly higher than the Con group (P<0.01). By contrast, serum glucose level was decreased following CA administration (Fig. 1H). The present results suggested that HFD could induce metabolic syndrome in mice, and the effects of HFD were reversed by CA administration. The present results indicated that CA may be used to treat metabolic syndrome.

In order to investigate whether CA could improve metabolic disease-associated inflammatory response, the relative expression levels of key factors associated with the inflammatory response were examined. HFD induced the inflammatory response in mouse liver tissue (Fig. 2A). The pathological score in the Veh group (pathological score, 3.2) was significantly higher than the Con group (pathological score, 0; P<0.001). By contrast, mice in the CA-L and CA-H groups exhibited a significant decrease in the pathological score (pathological scores, 1 and 0.8, respectively) compared with the Veh group (pathological score, 3.2; P<0.01). Moreover, pancreas injury was observed in the Veh group (Fig. 2B; pathological score, 3.0). Pancreas injury was significantly decreased following CA administration (pathological scores, 0.8 and 0.5 in the CA-L and CA-H groups, respectively). TNF-α and IL-1β are important pro-inflammatory cytokines, with a role in various diseases (32). Immunohistochemistry was performed to investigate the protein expression levels of TNF-α and IL-1β in mouse livers (Fig. 2C and D). The protein expression levels of TNF-α and IL-1β were significantly upregulated in the Veh group compared with in the Con group (P<0.001). Treatment with CA was able to significantly decrease the expression levels of both factors (P<0.001). The present results suggested that CA effectively inhibited the inflammatory response. Moreover, the serum levels of pro-inflammatory cytokines, including IL-1β, IL-6 and TNF-α, were investigated. The concentrations of IL-1β, IL-6 and TNF-α in the Veh group were significantly higher than in the Con group (P<0.001). The concentrations of these three pro-inflammatory cytokines were significantly reduced following treatment with CA (Fig. 2E-G). Collectively, the present results suggested that HFD promoted metabolism-associated inflammatory response in mice, and treatment with CA effectively inhibited the secretion of pro-inflammatory cytokines in mice fed a HFD.

Central nervous system injury caused by metabolic disease has been previously described (33). However, to the best of our knowledge, no effective therapeutic strategies are currently available. In the present study, brain injury was observed following HFD, as indicated by high protein expression levels of TNF-α and IL-1β in the brain (Fig. 3A). The present results suggested that HFD induced metabolic syndrome and brain injury in mice. Additionally, CA was able to inhibit the expression levels of TNF-α and IL-1β. Western blot analysis was performed to examine the protein expression levels of mature IL-1β and IL-18 in mouse brain tissues following different treatments. The protein expression levels of mature IL-1β and IL-18 were significantly higher in the Veh group compared with in the Con group (P<0.001). CA treatment decreased the protein expression levels of mature IL-1β and IL-18 compared with in the Veh group (P<0.001; Fig. 3B and C). Furthermore, the protein expression levels of TNF-α were upregulated in brain tissue collected from Veh mice (P<0.001). Notably, the protein expression levels of TNF-α were reduced following CA administration (P<0.001; Fig. 3B). In addition, RT-qPCR was used to investigate the mRNA expression levels of pro-inflammatory cytokines, including IL-1β, IL-6, IL-18 and TNF-α. The present results suggested that IL-1β, IL-6, IL-18 and TNF-α were upregulated in the Veh group compared with in the Con group (P<0.001). Notably, the expression levels of these genes were reduced following CA administration (P<0.001). The present qPCR results were consistent with the aforementioned western blotting results (Fig. 3C). Collectively, the present results suggested that CA inhibited the secretion of inflammatory cytokines in a mouse model of brain injury induced by HFD.

The NF-κB signaling pathway has been reported to be associated with the secretion of pro-inflammatory cytokines (34). The protein expression levels of p-IKKα, p-IκBα and p-NF-κB in the Veh group were increased compared with in the Con group (P<0.01; Fig. 4A). Notably, the protein expression levels of p-IKKα, p-IκBα and p-NF-κB were significantly reduced following CA administration (P<0.05). The present results suggested that brain injury caused by HFD involved the NF-κB signaling pathway, and CA was a negative regulator of the NF-κB signaling pathway. As a biomarker of nerve injury, GFAP is expressed in astrocytes (35). In the present study, GFAP was highly expressed in mouse brain tissues. High CA dosage decreased the expression levels of GFAP (Fig. 4A). However, low CA dosage did not affect the protein expression levels of GFAP. The present data suggested that CA influenced the expression levels of GFAP in a dose-dependent manner. Additionally, the protein expression levels of Neu-N and Iba1 were decreased following treatment with CA at low and high concentrations (Fig. 4B and C). Notably, the protein expression levels of Neu-N were significantly upregulated in the Veh group compared with in the Con group (P<0.001). Collectively, it was suggested that HFD caused nerve injury by activating the NF-κB signaling pathway. In addition, HFD increased the protein expression levels of GFAP, Neu-N and Iba-1. Treatment with CA may attenuate brain injury by decreasing the protein expression levels of GFAP, Neu-N, Iba-1 and p-NF-κB.

Neurodegeneration has been suggested to be associated with inflammatory response and cell apoptosis (36). Notably, Caspase-3 is an important regulator of apoptosis (37). In the present study, the Caspase-3 apoptotic pathway was investigated. The pro-apoptotic factors Bax and MMP-9 were significantly upregulated in mouse brain tissues from the Veh group compared with in the Con group (P<0.01; Fig. 5A and B). CA treatment significantly decreased the protein expression levels of both factors (P<0.01). Additionally, the protein expression levels of Caspase-3 were increased in the Veh group, but were reduced following CA treatment (P<0.001; Fig. 5C). By contrast, Bcl-2, an anti-apoptotic factor, was significantly downregulated in the Veh group compared with in the Con group (P<0.05), and treatment with CA was sufficient to upregulate the protein expression of Bcl-2 in mouse brain (P<0.05; Fig. 5D). In order to further investigate the expression levels of these factors, RT-qPCR was performed to examine the mRNA expression levels of Bax, MMP-9, Caspase-3 and Bcl-2 in different groups. The qPCR results were consistent with the aforementioned protein expression results (Fig. 5E). Collectively, the present results suggested that CA attenuated nerve injury caused by apoptosis in the mouse brain.