SW837 and SW480 cell lines were purchased from the American Type Culture Collection (Manassas, VA, USA). Cells were cultured in Dulbecco's modified Eagle's medium (Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc.) at 37°C in an atmosphere containing 5% CO₂ (23). Tan IIA (1–20 µM; cat. no. 568-72-9; Sigma-Aldrich; Merck KGaA) was used to treat cells for 12 h and the PBS-treated cells were used as the control group. In order to activate mitophagy, cells were pretreated with carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP; 5 µM, cat. no. S8276; Selleck Chemicals, Houston, TX, USA) for ~5 min at 37°C; and to inhibit mitophagy, 3-methyladenine (MA) (10 nM) was used to treat cells for ~2 h at 37°C. To ac tivate and inhibit the AMPK pathways, cells were incubated for ~4 h at 37°C with 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR; 10 µM) and compound C (20 µM), respectively.

Firstly, SW837 cells (1×10⁶) were washed with PBS and fixed with 4% paraformaldehyde for 30 min at room temperature. Following this, 0.1% Triton X-100 was used to permeabilize the samples for ~15 min at room temperature. To perform the immunofluorescence assay, the following primary antibodies were incubated with the samples overnight at 4°C (24): Anti-translocase of outer mitochondrial membrane 20 (Tom20; 1:500; cat. no. ab78547), which was used to label mitochondria; anti-lysosomal-associated membrane protein 1 (1:500; cat. no. ab24170), which was used to label lysosomes; anti-cytochrome c (cyt-c; 1:500; cat. no. ab133504), anti-p-Parkin (1:250; cat. no. ab73016) and anti-Skp2 (1:250; cat. no. ab68455; all Abcam, Cambridge, UK). Subsequently, samples were incubated with Alexa Fluor 488 donkey anti-rabbit secondary antibodies (1:1,000; cat. no. A-21206; Invitrogen; Thermo Fisher Scientific, Inc.) for ~1 h at room temperature. DAPI was used to label the nuclei, and images were captured using an inverted microscope (magnification, ×40; BX51; Olympus Corporation, Tokyo, Japan).

SW837 cells were washed with PBS and lysed in Laemmli Sample Buffer (Bio-Rad Laboratories, Inc., Hercules, CA, USA), and further homogenized with a rotor-stator homogenizer. Proteins were isolated and concentrations were determined using the Bicinchoninic Acid Protein Assay kit (Thermo Fisher Scientific, Inc.) (25). Equal amounts of protein (20 or 30 µg) were resolved via 8–15% SDS-PAGE and then transferred to polyvinylidene difluoride membranes (EMD Millipore, Billerica, MA, USA) (26). Membranes were blocked with 5% nonfat dried milk in Tris-buffered saline containing 0.05% Tween-20 (TBST) for 2 h at room temperature and were incubated overnight at 4°C with primary antibodies. The primary antibodies used were as follows: Anti-pro-caspase-3 (1:1,000; cat. no. 9662; Cell Signaling Technology, Inc., Danvers, MA, USA), anti-microtubule-associated proteins 1A/1B light chain 3B (LC3)II (1:1,000; cat. no. 3868; Cell Signaling Technology, Inc.), anti-complex III subunit core (CIII-core2; 1:1,000; cat. no. 459220; Invitrogen; Thermo Fisher Scientific, Inc.), anti-complex II (CII-30; 1:1,000; cat. no. ab110410), anti-complex IV subunit II (CIV-II; 1:1,000; cat. no. ab110268), anti-p-Parkin (1:11,000; cat. no. ab73016), anti-Skp2 (1:11,000; cat. no. ab68455), anti-GAPDH (1:11,000; cat. no. ab9485), anti-p62 (1:11,000; cat. no. ab56416), anti-β-actin 1:11,000; cat. no. ab8226; all Abcam), anti-Beclin1 (1:1,000; cat. no. 3495; Cell Signaling Technology, Inc.), anti-B-cell lymphoma 2 (Bcl-2) associated agonist of cell death (Bad; 1:1,000; cat. no. ab90435; Abcam), anti-cleaved caspase-3 (1:1,000; cat. no. 9664; Cell Signaling Technology, Inc.), anti-caspase-9 (1:1,000; cat. no. ab32539), anti-Poly (ADP-ribose) polymerase 1 (PARP1; 1:1,000; cat. no. ab32138; both Abcam), anti-autophagy-related 5 (ATG5; 1:1,000; cat. no. 12994; Cell Signaling Technology, Inc.), anti-cellular inhibitor of apoptosis 1 (C-IAP1; 1:1,000; cat. no. ab25939), anti-survivin (1:1,000; cat. no. ab182132), anti-Bcl-2 (1:1,000; cat. no. ab196495), anti-AMPK (1:1,000; cat. no. ab32047), anti-phosphorylated (p)-AMPK (1:1,000; cat. no. ab133448) and anti-Parkin (1:1,000; cat. no. ab15954; all Abcam) (27). The membrane was subsequently washed with TBST (5 min; three times) and incubated with horseradish peroxidase-conjugated secondary antibodies (1:2,000; cat. nos. 7076 and 7074; Cell Signaling Technology, Inc.) for 1 h at room temperature. Following washing with TBST (5 min; three times), bands were detected using an enhanced chemiluminescence substrate (Applygen Technologies, Inc., Beijing, China). Band intensities were normalized to the respective internal standard signal intensity (β-actin or GAPDH) using Quantity One Software (version 4.6.2; Bio-Rad Laboratories, Inc.).

Cells were washed with cold PBS and incubated on ice in lysis buffer (cat. no. C3601; Beyotime Institute of Biotechnology, Haimen, China) for 30 min. The cells were subsequently scraped, and homogenates were spun at 800 × g for 5 min at 4°C. The supernatants were centrifuged at 10,000 × g for 20 min at 4°C to acquire the pellets, which were spun again. The final pellets were suspended in lysis buffer containing 1% Triton X-100 and were noted as mitochondrial-rich lysate fractions (28,29).

SW837 cells were used to analyze mROS, mitochondrial potential, ATP production and mPTP opening. mROS levels were detected using the MitoSOX red probe (Molecular Probes; Thermo Fisher Scientific, Inc.) (30). Cells (1×10⁶) were cultured with the MitoSOX red probe at 37°C for ~15 min. Subsequently, PBS was used to wash the cells three times. Finally, mROS production was detected via flow cytometric analyses using a BD FACSCalibur™ flow cytometer (BD Biosciences, San Jose, CA, USA) (31).

A JC-1 assay was used to investigate mitochondrial potential. Briefly, cells (1×10⁶) were treated with a MitoProbe™ JC-1 assay kit (Thermo Fisher Scientific Inc.) (10 mg/ml) at 37°C in the dark for 15–20 min. Subsequently, PBS was used to wash the cells three times. Finally, mitochondrial potential was determined using a fluorescence microscope, and the images were captured. In addition, mitochondrial function was determined via ATP production using a Celltiter-Glo Luminescent Cell Viability assay (Promega Corporation, Madison, WI, USA) according to the manufacturer's protocol (32). Furthermore, in the mPTP opening assay, calcein-acetoxymethyl ester (5 µM, cat. no. 148504-34-1; Sigma-Aldrich; Merck KGaA) was incubated with SW837 cells at room temperature in the dark for 30 min. Subsequently, the mPTP opening rate was determined according to a previous study (33).

MTT assay was used to determine cellular viability. SW837 cells and SW480 cells were treated with 50 µl MTT at 37°C for ~4 h. Subsequently, cells were incubated with 200 µl dimethyl sulfoxide for ~10 min at 37°C (34). The optical density at a wavelength of 570 nm was then determined. Furthermore, cellular viability was also investigated using LDH release ELISA kit (cat. no. C0016; Beyotime Institute of Biotechnology) according to the manufacturer's protocol (35).

Extracellular lactate levels were measured in the cell culture medium using a lactate assay kit (cat. no. K607-100; BioVision, Inc., Milpitas, CA, USA). Intracellular glucose levels were measured in the cell lysates using a glucose assay kit (cat. no. K606-100; BioVision, Inc.). The uptake of glucose and the production of lactate were measured according to the manufacturer's protocols, and as previously described (36,37). Mitochondrial respiration was initiated by the addition of glutamate/malate, at a final concentration of 5 and 2.5 mmol/l, respectively. State 3 respiration was initiated by the addition of ADP (150 nmol/l); state 4 was measured as the rate of oxygen consumption following ADP phosphorylation (38,39).

PI is a popular red-fluorescent nuclear and chromosome counterstain. Since PI cannot permeate live cells, it is also commonly used to detect dead cells in a population (40). Cells were treated with 1 mg/ml PI (Invitrogen; Thermo Fisher Scientific, Inc.) for ~15 min at room temperature. Subsequently, samples were washed three times with PBS, and DAPI (cat. no. 28718-90-3; Sigma-Aldrich; Merck KGaA) was used for nuclear staining for 5 min at room temperature. The images were acquired following Tan IIA treatment using a fluorescence microscope with standard excitation filters (Olympus Corporation) (41).

To investigate cellular apoptosis, TUNEL assays and trypan blue staining were performed. A TUNEL assay was performed using a TUNEL assay kit (Roche Applied Science, Madison, WI, USA) according to the manufacturer's protocol (42). Images were captured using an inverted microscope (magnification, ×40; BX51; Olympus Corporation). For trypan blue staining, cells were treated with 0.4% trypan blue at 37°C for ~2 min. Subsequently, the cells were observed under a light microscope (magnification, ×100; BX51; Olympus Corporation). Furthermore, caspase-3/9 activity levels were determined, in order to investigate cellular apoptosis, via caspase-3/9 activity kits (cat. nos. C1158 and C1115; Beyotime Institute of Biotechnology) according to the manufacturer's protocols (43). SW837 cells underwent TUNEL staining and caspase-3/9 activity assays.

To induce the overexpression of Skp2, pCMV6-Kan/Neo Skp1 plasmids were purchased from OriGene Technologies, Inc. (Rockville, MD, USA) (44). Transfection of SW837 cells (1×10⁶) with the Skp2 plasmid (1,336 bp; pDC315-Skp2-NheI-F, 5′-ATCTGTGACCTTAGACCTGATCCGTA-3′ and pDC315-Skp2-HindIII-R, 5′-GGTACCGATAGGAACATATTACCAGT-3′) (3.0 µg per 1×10⁴ cells/well) was performed using Lipofectamine 2000^® (Invitrogen; Thermo Fisher Scientific, Inc.). Following 48 h of incubation at 37°C. Finally, the supernatant was filtered and isolated in order to obtain the adenovirus Skp2 (Skp2 OE). Subsequently, adenovirus Skp2 was transfected into the SW837 cells to overexpress Skp2. Skp2 infection was carried out via incubating SW837 cells with adenovirus Skp2 in Opti-MEM media supplemented with Lipofectamine^® 2000 (Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. Infection was performed for 48 h at 37°C and infection efficiency was confirmed via western blotting (45). Null vector transfection was used as the control group (Ad-ctrl).

Experiments were repeated three times. Data are presented as the means ± standard error of the mean. One-way analysis of variance followed by Bonferroni's multiple comparison test was performed to analyze data using SPSS software (version 17.0; SPSS, Inc., Chicago, IL, USA). P<0.05 was considered to indicate a statistically significant difference. Experiments were repeated in triplicate.

To investigate whether Tan IIA enhances CRC apoptosis, an MTT assay was performed to determine the viability of SW480 and SW837 cells. When compared with the control group, Tan IIA significantly reduced the viability of SW40 cells (Fig. 1A) and SW837 cells (Fig. 1B), thus suggesting that Tan IIA may suppress viability of CRC cells. Notably, for both SW480 and SW837 cells, viability progressively decreased following treatment with Tan IIA in a dose-dependent manner (Fig. 1A and B). Since no differences were observed in the levels of cellular viability between SW837 and SW480 cells following treatment with Tan IIA, the SW837 cell line was used in the following study. To further investigate whether Tan IIA could promote the apoptosis of cancer cells, trypan blue staining was performed. When compared with the control group, Tan IIA increased the number of trypan blue-positive cells (Fig. 1C and D) in a dose-dependent manner. In addition, the results of a TUNEL assay were in agreement with the results obtained from trypan blue staining. The results of the TUNEL assay demonstrated that treatment with Tan IIA significantly enhanced the apoptosis of SW837 cells (Fig. 1E and F). Similar results were revealed from the LDH release assay, which suggested that Tan IIA significantly enhanced CRC cell apoptosis in a dose-dependent manner (Fig. 1G). Furthermore, it was revealed that the maximum lethal concentration of Tan IIA tested was 20 µM, whereas 1 µM had no influence on cellular viability. Therefore, 1 and 20 µM were used in subsequent experiments.

The proapoptotic effects of Tan IIA on CRC were subsequently investigated. Based on the results of a previous study (46), it was suggested that Tan IIA may regulate mitochondrial function, which is important for cell survival. Therefore, the present study investigated the levels of mitochondrial apoptosis. When compared with the control group, treatment with Tan IIA was revealed to significantly enhance the production of mROS (Fig. 2A and B). This effect was associated with a significant reduction in mitochondrial potential via JC-1 staining (Fig. 2C and D). Furthermore, Tan IIA was demonstrated to significantly increase the mPTP opening rate (Fig. 2E), which has previously been revealed to represent a feature of mitochondrial apoptosis activation (47). Following mPTP opening, mitochondria can release the proapoptotic factor cyt-c into the cytoplasm, thus resulting in cellular apoptosis (48). By determining the immunofluorescence levels of cyt-c, it was demonstrated that Tan IIA increased cyt-c leakage into the cytoplasm and the nucleus (Fig. 2F). To further investigate mitochondrial apoptosis, western blotting was performed (Fig. 2G-N). The results revealed that treatment with Tan IIA significantly upregulated the expression levels of proapoptotic proteins (caspase-3, PARP, caspase-9 and Bad), and significantly downregulated the expression levels of anti-apoptotic proteins (C-IAP1, survivin and Bcl-2).

Mitochondrial energy metabolism is important for cancer survival. In order to investigate mitochondrial energy metabolism following treatment with Tan IIA, ATP content was determined; the results revealed that Tan IIA significantly suppressed ATP production in SW837 cells compared with in the control group (Fig. 3A), which suggested that Tan IIA suppressed the mitochondrial ATP supply. Notably, mitochondrial ATP is primarily generated by the mitochondrial electron transfer respiratory chain (ETC) (48); however, as shown in Fig. 3B-E, Tan IIA suppressed the expression of ETCs when compared with the control group. ETCs are important factors for ATP production (49), and therefore the inhibitory effects of Tan IIA on ETC levels may be responsible for ATP suppression in SW837 cells. Furthermore, ETC-associated mitochondrial respiratory function, such as state 3 and state 4 respiratory rates, were also suppressed in Tan IIA-treated cells when compared with the control group (Fig. 3F and G). These results suggested that treatment with Tan IIA may suppress mitochondrial energy production. To investigate this further, alterations in glycometabolism were determined following treatment with Tan IIA. As revealed in Fig. 3H, Tan IIA was demonstrated to significantly suppress glucose uptake in SW837 cells compared with in the control group. Furthermore, levels of lactate production were significantly decreased in Tan IIA-treated cells compared with in the control group (Fig. 3I). These results suggested that treatment with Tan IIA may suppress mitochondrial energy metabolism in CRC.

A previous study revealed that mitophagy (48), which is an important mitochondrial self-protective mechanism, is responsible for CRC cell survival in response to radiotherapy and chemotherapy; therefore, in the present study, cellular mitophagy activity was determined following treatment with Tan IIA. Notably, when compared with the control group, treatment with Tan IIA (20 µM) markedly decreased mitochondria engulfment by lysosomes, which is indicative of mitophagy inactivation (Fig. 4A). Conversely, co-culture with FCCP, an activator of mitophagy, was revealed to enhance the fusion of lysosomes and mitochondria, which was demonstrated to subsequently block Tan IIA-inhibited mitophagy (Fig. 4A). Furthermore, western blotting was used to investigate mitophagy activity. Following treatment with Tan IIA, the expression levels of mitochondrial (mito)-LC3II, Beclin1, ATG5 and p62 were revealed to be significantly decreased compared with in untreated cells, thus suggesting that mitophagy was suppressed (Fig. 4B-F). Conversely, following treatment with FCCP, an activator of mitophagy, the expression levels of mito-LC3II, Beclin1, ATG5 and p62 were revealed to be attenuated compared with in cells treated with Tan IIA alone. Furthermore, the mitophagy inhibitor 3-MA was administered to cells to perform a loss of function assay regarding mitophagy. Following treatment with 3-MA in FCCP-treated cells, the expression levels of mito-LC3II, Beclin1, ATG5 and p62 were revealed to be significantly suppressed compared with cells treated with Tan IIA + FCCP, thus suggesting that mitophagy was inhibited (Fig. 4B-F).

To investigate the consequences of mitophagy inhibition following Tan IIA treatment, caspase-9 activity and cellular death were investigated. As revealed in Fig. 4G, caspase-9 activity was significantly increased following treatment with Tan IIA compared with in the control group. However, treatment with FCCP significantly attenuated this effect. Similar results were also revealed using PI staining (Fig. 4H), which is a marker of cell death. In conclusion, these results suggested that mitophagy may be suppressed following treatment with Tan IIA, which potentially contributed to mitochondria-dependent apoptosis.

To determine the function of Tan IIA in mitophagy inactivation, Parkin-dependent mitophagy was investigated. In response to mitochondrial damage, Parkin is activated, which contributes to the fusion between mitochondria and lysosomes. Notably, numerous studies have demonstrated the regulatory signaling associated with Parkin-mediated mitophagy, including c-Jun N-terminal kinase, AMPK and ROS (50–53). Therefore, whether Parkin is activated by AMPK and subsequently contributes to mitophagy activation following treatment with Tan IIA was investigated in the present study (Fig. 5). Firstly, as revealed in Fig. 5A, Tan IIA treatment resulted in a marked decrease in the levels of p-Parkin compared with in the control group, thus suggesting that a strong association may exist between Tan IIA and Parkin-mediated mitophagy. Subsequently, it was demonstrated that AMPK activity was markedly suppressed following treatment with Tan IIA, as demonstrated by reduced p-AMPK levels; however, this effect was attenuated by treatment with FCCP (Fig. 5A). Notably, following treatment with the AMPK activator AICAR, the phosphorylation levels of AMPK and Parkin were markedly increased compared with in the Tan IIA treatment group (Fig. 5A-C). In addition, Compound C, an inhibitor of AMPK, was used as a positive control. Treatment with compound C markedly suppressed the expression levels of p-AMPK and p-Parkin, which was similar to the results exhibited by the Tan IIA treatment group.

Skp2 represents a novel regulator of autophagy; however, little is known about its involvement in mitophagy (20). In the present study, the results demonstrated that treatment with Tan IIA suppressed the expression of Skp2 compared with in the control group via immunofluorescence analysis (Fig. 5F). However, re-activation of AMPK was revealed to attenuate Skp2 downregulation. Conversely, inhibition of AMPK via treatment with compound C was able to markedly suppress the expression levels of Skp2 (Fig. 5F). These results suggested that Skp2 may function downstream of the AMPK pathway.

To determine the function of Skp2 in Parkin regulation, Skp2 OE was performed, the efficiency of which was confirmed by western blotting (Fig. 5D and E). Notably, in Skp2 OE cells treated with Tan IIA, the expression levels of Skp2 and p-Parkin were enhanced compared with in the Tan IIA-treated cells that did not possess Skp2 OE (Fig. 5F). These results demonstrated that Parkin was suppressed by Tan IIA-induced downregulation of Skp2 and AMPK. To establish the association between AMPK/Skp2/Parkin and mitophagy, the expression levels of mitophagy markers (ATG5, Beclin1, p62 and mito-LC3II) were investigated. The inhibitory effects of Tan IIA on the expression levels of mitophagy markers were markedly attenuated following treatment with AICAR or Skp2 OE (Fig. 5G-J). Furthermore, caspase-3 activity was determined, in order to investigate the effects of AMPK/Skp2/Parkin on cell death. Increased caspase-3 activity following treatment with Tan IIA was significantly decreased following treatment with AICAR and in Skp2 OE cells (Fig. 5L). In conclusion, these results suggested that Parkin-mediated mitophagy was markedly suppressed following treatment with Tan IIA via the AMPK/Skp2 pathway.