PTE, 3-methyladenine (3-MA), monodansylcadaverine (MDC) and dimethyl sulfoxide (DMSO) were purchased from Sigma-Aldrich; Merck KGaA (Darmstadt, Germany). Compound C was purchased from Selleck Chemicals (Houston, TX, USA).

PTE was dissolved in DMSO to yield a 78 mM stock solution that was stored at −20°C. The different doses of PTE (10, 20, 30, 40, 50, 60 and 70 µM) for each treatment group were diluted with RPMI-1640 medium (Sigma-Aldrich; Merck KGaA). All experiments used a corresponding volume of DMSO as a control treatment.

RPMI-8226, NCI-H929, U266 and ARH-77 human MM cell lines were purchased from American Type Culture Collection (Manassas, VA, USA). The RPMI-8226, NCI-H929, U266 and ARH-77 cells were cultured in RPMI-1640 medium supplemented with 10% fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) and 100 U/ml penicillin/streptomycin (Beyotime Institute of Biotechnology, Haimen, China). The cells were incubated in a humidified incubator with 5% CO₂ at 37°C.

Cell viability was determined using a Cell Counting kit-8 (CCK-8; Dojindo Laboratories, Kumamoto, Japan) assay. In brief, 1×10⁴ cells/well were seeded into 96-well plates in a final volume of 100 µl of complete culture medium with the specified concentrations of PTE. The CCK-8 reagent was added and incubated for an additional 0.5-4 h at 37°C and the optical density (OD) was measured at 450 nm using a microplate reader.

Following PTE treatment with or without 1 µM compound C for 48 h, the MM cells were harvested and lysed in radioimmunoprecipitation assay lysis solution (Beyotime Institute of Biotechnology) containing phosphatase inhibitors and PMSF. A bicinchoninic acid protein assay kit (Beyotime Institute of Biotechnology) was used to determine the protein concentrations. Protein lysates (30-50 µg per sample) were separated with 6-12% SDS-polyacrylamide gels (Beyotime Institute of Biotechnology) and transferred to polyvinyl difluoride membranes (EMD Millipore, Billerica, MA, USA). The membranes were blocked in 5% skimmed milk in Tris-buffered saline for 1 h at room temperature and incubated at 4°C overnight with primary antibodies against cleaved caspase 9 (Asp330; cat. no. 9501T), caspase 9 (cat. no. 9502T), poly(ADP-ribose) polymerase (PARP; cat. no. 9532), phosphorylated (p)-AMPK (Thr172; cat. no. 2535T), AMPK (cat. no. 5832T), FASN (cat. no. 3180T), ACC (cat. no. 3676T), p-ACC (Ser79; cat. no. 11818T), p-mTOR (Ser2448; cat. no. 5536), mTOR (cat. no. 2983), 4E-BP1 (cat. no. 9644T), p4E-BP1 (Thr37/46) (cat. no. 2855T), p-eIF2α (Ser51; cat. no. 3398T), eIF2α (cat. no. 5324T), autophagy-related (ATG)5 (cat. no. 12994T), beclin1 (cat. no. 3495T) and light chain (LC)3B (cat. no. 3868T; all dilution, 1:1,000; Cell Signaling Technology, Inc., Danvers, MA, USA), and primary antibody against cleaved caspase 3 (cat. no. ab32042; dilution, 1:500; Abcam, Cambridge, MA, USA). The primary antibody against β-actin was from Antgene (cat. no. ANT010S; dilution, 1:2,000; Wuhan, China). The membranes were incubated with horseradish peroxidase-conjugated anti-rabbit (cat. no. ANT020; dilution, 1:5,000; Antgene) or anti-mouse (cat. no. ANT019; dilution, 1:5,000; Antgene) secondary antibodies for 60 min at room temperature. The blots were developed using an enhanced chemiluminescence reagent (Antgene). The protein expression levels were normalized to the level of β-actin in the corresponding lanes and Image Lab 3.0 software (Bio-Rad Laboratories, Inc., Hercules, CA, USA) was used for densitometric analysis.

Following PTE treatment, or PTE co-treatment with 1 µM compound C or 3 mM 3MA, cell apoptosis was assayed with Annexin V/propidium iodide staining (PI; BD Biosciences, San Jose, CA, USA), followed by fluorescence-activated cell sorting analysis. RPMI-8226 cells were treated with 0, 20, 40 or 60 µM PTE and ARH-77 cells were treated with 0, 10, 20 or 40 µM PTE for 24 h. Next, 1×10⁶ cells were resuspended in 100 µl Annexin V-binding buffer containing 5 µl Annexin V and PI. Following incubation at room temperature for 15 min, 400 µl binding buffer was added to the cells, which were analyzed using a FACSAriaII flow cytometer (BD Biosciences). The rate of apoptotic events was defined as the sum of the number of cells in the early (Annexin V⁺/PI^-) and late (Annexin V⁺/PI⁺) stages of apoptosis. FlowJo 7.6 software (FlowJo LLC, Ashland, OR, USA) was used to analyze the flow cytometry data.

MDC was used as an autophagolysosome-specific marker to analyze the autophagic process. Cells were seeded into 6-well plates and treated with 40 µM PTE for 24 h. Autophagic vacuoles were labeled with MDC by incubating cells with 50 µM MDC at 37°C for 30 min. The cells were washed with PBS three times in 5 min intervals. The cells were then immediately analyzed using a fluorescence microscope (magnification, ×200; Nikon Corporation, Tokyo, Japan).

The autophagosome ultrastructure of MM cells was observed by TEM. RPMI-8226 cells were exposed to 40 µM PTE for 0, 7 and 24 h. The cells were fixed with 2.5% glutaraldehyde at 4°C for 2 h, followed by incubation with 2% O_sO₄ for 2-3 h at 4°C. Following dehydration in a graded ethanol series, cells were infiltrated and embedded in EMbed-812 (Electron Microscopy Sciences, Hatfield, PA, USA) at 60°C for 48 h. Thin (60-100 nm) sections were stained at room temperature with uranyl acetate for 15 min and lead citrate for 10 min, prior to being examined using a Tecnai G2 20 TWIN electron microscope (magnification, ×1,700 and ×5,000; FEI, Eindhoven, the Netherlands).

A total of 10 female NOD/SCID mice (weight, 15.7±1.0 g; age, 3-4 weeks; Nanjing Bioscience Company, Nanjing, China) were individually maintained in a temperature-controlled (23±2°C with 50-60% relative humidity) room with a 12-h light/dark cycle and ad libitum access to water and food. All experimental procedures and protocols were approved by the Committee on Animal Handling of Huazhong University of Science and Technology (Wuhan, China). The mice were subcutaneously injected with 2×10⁷ RPMI-8226 cells in 200 µl serum-free RPMI-1640 medium. The treatment began when the tumor volume reached ~100 mm³ (~3 weeks). Tumor volumes were calculated using the following formula: V = A×B²/2, where A is the largest diameter and B the smallest. 10 mice were randomly divided into the control (5% DMSO) or PTE (50 mg/kg in 5% DMSO) groups (n=5). Intraperitoneal injections were performed 5 days a week for 21 days. Tumor sizes were evaluated daily. All mice were sacrificed on day 21 and their tumors were excised. Tumor tissues were fixed in 10% formalin at room temperature for 24 h for further analysis. Blood was also collected at sacrifice and serum samples were stored at −80°C until further analysis.

A TUNEL assay kit (cat. no. 11684817910; Roche Diagnostics, Ltd., Basel, Switzerland) was used to detect apoptosis in the tumor tissues, according to the manufacturer's protocol. Tumor tissues from mice were fixed in 10% formalin at room temperature for 24 h. Paraffin-embedded sections (4-µm) from tumor tissues were deparaffinized and hydrated by sequential immersion in xylene and a graded alcohol series (100, 100, 95, 90, 80 and 70%). Next, the sections were incubated with the TUNEL reaction mixture (TdT+dUTP, v/v=1:9) at 37°C for 1 h, and hematoxylin was used to stain the nuclei at room temperature for 5 min. The sections were sealed with neutral gum and visualized using a fluorescence microscope (magnification, ×100; Nikon Corporation). Five fields of view for each section were observed.

Tumors from mice were fixed in 10% formalin at room temperature for 24 h. Paraffin-embedded sections (4-µm) of tumor tissues were deparaffinized and hydrated by sequential immersion in xylene and a graded alcohol series (100, 100, 95, 90, 80 and 70%). Following antigen retrieval with 10 mM citrate buffer (pH=6.0) at 95°C for 10 min, endogenous tissue peroxidase activity was blocked with 3% hydrogen peroxide at room temperature for 10 min. Next, non-specific sites were blocked with 5% bovine serum albumin (Roche Diagnostics) at room temperature for 20 min. The sections were incubated with antibodies against p-AMPK (Thr172; cat. no. 2535S; dilution, 1:100; Cell Signaling Technology, Inc.), p-mTOR (cat. no. bs-3494R; dilution, 1:200; Biosynthesis Biotechnology Co., Ltd., Beijing, China), FASN (cat. no. 3180S; dilution, 1:50; Cell Signaling Technology, Inc.) and cleaved caspase-3 (cat. no. RLC006; dilution, 1:50; Ruiying Biological, Suzhou, China) at 4°C overnight. Next, the sections were incubated with the following secondary antibodies: horseradish peroxidase-conjugated anti-rabbit or anti-mouse (cat. no. K5007; dilution, 1:200; Dako; Agilent Technologies, Santa Clara, CA, USA) for 50 min at room temperature. The labelled cells were observed using a fluorescence microscope (Nikon Corporation) for cleaved caspase-3 (magnification, ×100) and for p-AMPK, FASN and p-mTOR (magnification, ×200). Five fields of view for each section were observed.

Inner canthus blood samples (100 µl) were collected from the mice prior to sacrifice. Blood cells were isolated by density gradient centrifugation at 3,000 x g at room temperature for 10 min, and creatine kinase-MB (CK-MB), cardiac troponins T (cTnT), alanine aminotrans-ferase (ALT), aspartate amino transferase (AST), alkaline phosphatase (ALP), total bilirubin (TBIL), urea nitrogen (BUN) and serum creatinine (Scr) were detected in the serum. Concentrations of CK-MB and cTnT were determined quantitatively using the commercially available CK-MB ELISA kit (cat. no. FLSW-M1025; Kehua Bio-Engineering co., Ltd., Shanghai, China) and cTnT ELISA kit (cat. no. FLSW-M763; Kehua Bio-Engineering co., Ltd.) according to the manufacturer's protocol. The absorbance at 450 nm was measured using an automated microplate reader. The other serum indexes, including ALT, AST, ALP, TBIL, BUN and Scr, were measured by routine methods using a fully automatic biochemical analyzer (Automatic Analyzer AU2700; Beckman Coulter, Inc., Brea, CA, USA), according to the manufacturer's protocol.

SPSS 15.0 software (SPSS Inc, Chicago, IL, USA) was used to analyze all recorded data. Data are presented as the mean ± standard deviation of three independent experiments. Statistically significant differences between multiple group comparisons were determined using one-way analysis of variance followed by Dunnett's test. Comparisons between two groups were made using unpaired Student's t-tests. P<0.05 was considered to indicate a statistically significant difference.

PTE treatment has demonstrated antitumor activity in several types of human cancer cells (28,29). Therefore, the growth inhibition effect of PTE on MM cells was assessed. Different concentrations of PTE between 0 and 70 µM were applied to the MM RPMI-8226, ARH-77, U266 and NCI-H929 cell lines for 48 h. Consistent with other previous studies (30,31), a significant reduction in cell viability was observed following the PTE treatment of MM cells. PTE induced the concentration-dependent decrease in viability of RPMI-8226, ARH77, U266 and NCI-H929 cells, with IC₅₀ values at 48 h of 33.8, 16.6, 63.5 and 44.5 µM, respectively (Fig. 1A). The relatively sensitive RPMI-8226 and ARH-77 cells were used for subsequent experiments.

To investigate the mechanisms of the growth inhibitory effect of PTE on MM cells, the effects of PTE on the rate of apoptosis were analyzed by flow cytometry. PTE treatment for 24 h induced early and late apoptosis to a greater extent compared with the DMSO-treated controls in RPMI-8226 and ARH-77 cells in a dose-dependent manner (Fig. 1B). Compared with the control group (2.3+7.9%), apoptosis was increased at 24 h in the 20 µM (6.2+12.5%), 40 µM (13.2+17.5%) and 60 µM (23.2+20.2%) PTE-treated groups in RPMI-8226 cells. Similar findings were observed in ARH77 cells, in which apoptosis was increased at 24 h in the 10 µM (6.8+9.9%), 20 µM (12.5+12.9%) and 40 µM (18.3+14.1%) PTE-treated groups, compared with the control group (3.9+6.9%). Taken together, these data indicated that PTE treatment increased the rate of apoptosis of MM cells.

These results coincided with a marked increase in the expression of the pro-apoptotic proteins, cleaved PARP and cleaved caspase 3/9, following PTE exposure for 48 h in RPMI-8226 and ARH-77 cells (Fig. 1C and D). Taken together, these results suggested that PTE treatment attenuates MM cell viability and induces apoptosis.

As the survival of MM cells depends on high rates of anabolic metabolism, and AMPK is a critical energy sensor in cellular metabolism (23,32), it was assessed whether PTE treatment activated the AMPK pathway in MM cells. RPMI-8226 and ARH-77 cells were exposed to different concentrations of PTE for 48 h, and the western blot data from these cells demonstrated that PTE induced the phosphorylation of AMPK in a dose-dependent manner (Fig. 2A).

The activity of the de novo fatty acid synthesis key enzymes FASN and ACC is negatively regulated by AMPK (33). Therefore, it was then investigated whether PTE decreased lipid synthesis by decreasing FASN expression or inhibiting ACC activity. It was observed that the FASN protein expression level was decreased, and ACC was phosphorylated in a dose-dependent manner when RPMI-8226 and ARH-77 cells were treated with PTE (Fig. 2B). The inhibition of lipogenic key enzymes may induce MM cells into a low de novo lipogenesis state.

As the survival of MM cells is dependent on extensive protein synthesis and the activation of AMPK inhibits the mTOR protein synthesis pathway (34,35), it was next examined whether the activation of AMPK by PTE affected the mTOR signaling pathway. It was demonstrated that PTE treatment for 48 h could inhibit mTOR phosphorylation in a dose-dependent manner in RPMI-8226 and ARH-77 cells. The best understood roles of mTOR in mammalian cells are regarding the control of mRNA translation by phosphorylating 4E-BP1 (20), it was observed that the inhibition of mTOR phosphorylation was accompanied by the decreased phosphorylation of 4E-BP1. Furthermore, eIF2α phosphorylation was increased in a dose-dependent manner (Fig. 2C). These results indicated that PTE activates AMPK phosphorylation to suppress the expression level and activity of lipogenesis- and mRNA translation-associated enzymes, potentially driving MM cells into a low nutrient state.

To more definitively establish the role of AMPK in mediating the apoptosis-promoting effect of PTE, a well-established pharmacological inhibitor of AMPK activity, compound C, was utilized. RPMI-8226 cells were treated with 60 µM PTE and ARH-77 cells with 40 µM PTE, in the absence or presence of compound C (1 µM). As expected, treatment with PTE alone activated AMPK phosphorylation. In addition, the expression levels of the lipogenesis-associated key enzyme FASN were decreased, and the activation of the mRNA translation-associated 4E-BP1 was suppressed in RPMI-8226 and ARH-77 cells after 48 h. However, the FASN expression level and 4E-BP1 activity were restored in the presence of compound C (Fig. 3A-C). Furthermore, pre-treating RPMI-8226 and ARH-77 cells with compound C for 1 h and then exposing the cells to PTE for 24 h partially reduced the ability of PTE to induce apoptosis (Fig. 3D). These results coincided with the expression of the pro-apoptotic protein, cleaved PARP. Following treatment with PTE for 48 h, the protein expression levels of cleaved PARP were increased, which were restored by co-treatment with compound C (Fig. 3E). Taken together, these results demonstrated that PTE decreased the expression level and activity of lipogenesis- and mRNA translation-associated proteins through the activation of the AMPK pathway, and the activation of AMPK at least in part, induced MM cells apoptosis.

Autophagy is activated when cellular energy is insufficient. The AMPK/mTOR pathway is involved in the regulation of autophagy in cancer cells (36,37). Therefore, the effect of PTE on autophagy was assessed. Using TEM, the gold standard for autophagy assessment, the majority of the RPMI-8226 cells in the control group were structurally complete, whereas PTE treatment resulted in an increase in autophagosomes (Fig. 4A). MDC is a specific label for autophagic vacuoles that was used to further identify PTE-induced autophagy (38). The PTE-treated RPMI-8226 cells displayed a greater fluorescence intensity and a greater number of MDC-labeled particles compared with the untreated control group (Fig. 4B). Furthermore, the western blotting analysis of cell lysates revealed the increased accumulation of autophagy markers, including LC3-II, beclin1 and ATG5, in a dose-dependent manner at 48 h (Fig. 4C and D). Autophagy can serve either a pro-survival or pro-death function (39). Its role in the effect of PTE intervention on MM cells was then demonstrated. The pro-apoptotic effect of PTE was further enhanced subsequent to blocking autophagic flux by co-treatment with 3 mM 3MA for 24 h (Fig. 4E), indicating that PTE-induced autophagy is a cell protective mechanism, as the inhibition of autophagy aggravated PTE-induced apoptosis. These data confirmed that PTE induced autophagy in MM cells. The co-presence of PTE and an autophagy inhibitor improved the anti-MM activity compared with PTE alone.

The in vivo antitumor effect of PTE was then examined using a mouse model. RPMI-8226 cells were subcutaneously injected into 3-week-old female NOD/SCID mice. As demonstrated in Fig. 5A, the xenograft tumors in the mice receiving PTE treatment exhibited a reduced tumor volume compared with the mice receiving DMSO. The tumor xenografts were harvested following the final treatment, and TUNEL assays and IHC were performed. From the in situ TUNEL assays, it was observed that tumors in mice treated with PTE had increased levels of apoptotic cells compared with control mice (Fig. 5B). IHC was performed to investigate the molecular mechanisms involved in the anti-tumorigenic effects of PTE. An increase in cleaved-caspase 3 staining confirmed the induction of apoptosis in the treatment group (Fig. 5B). Similarly, an increase in p-AMPK-positive cells and a decrease in FASN- and p-mTOR-positive cells were observed with IHC in the xenograft tumors of the PTE treatment group (Fig. 5C). The in vivo data demonstrated the promising anti-tumor effect of PTE for the treatment of MM, associated with metabolic dependence, implying that PTE may be an effective MM treatment.

Mice treated with PTE exhibited no notable change in body weight, food and water intake, or activity. Chemotherapy-induced heart disease, liver dysfunction or kidney malformation are common side effects that may restrict the clinical application of certain chemotherapeutic agents. Therefore, it was assessed whether the administration of PTE to tumor-bearing mice stimulated cardiac, nephritic or hepatic toxicity. Mouse plasma was collected at sacrifice, and mouse plasma myocardial damage and liver/kidney disease markers were quantified. The results revealed that the administration of PTE did not increase liver damage markers, including ALT, AST, ALP and TBIL (Fig. 6A; P>0.05). No significant increase in the kidney dysfunction indicators in BUN and Scr was observed (Fig. 6B; P>0.05). PTE treatment also did not result in cardiac damage, as no increase in cTnT or CK-MB was observed (Fig. 6C; P>0.05). Taken together, these results indicated that PTE treatment had no evident toxic effects on the heart, liver or kidneys.