The HCT116, SW480 and LoVo colorectal cancer cell lines were supplied by the Laboratory of Gastrointestinal Surgery, Union Hospital (Wuhan, China). The cells were grown in high glucose Dulbecco's modified Eagle's medium (DMEM; Hyclone; GE Healthcare Life Sciences, Logan, UT, USA) supplemented with 10% fetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA). SW620 cells (supplied by the Laboratory of Gastrointestinal Surgery, Union Hospital) were grown in RPMI-1640 (Hyclone; GE Healthcare Life Sciences) and supplemented with 10% FBS (16). All cells were maintained in a humidified 37°C incubator containing 5% CO₂. The HCT116, SW480 and LoVo cell lines were grown in DMEM with varying concentrations of JZL184 (10, 25 or 50 µM; Sigma-Aldrich, St. Louis, MO, USA) or dimethyl sulfoxide (DMSO; 0.1%), representing the control group, for 48 h. These cells were then used in the following assays, except the migration assay.

Cells (1×10⁴) of the HCT116, SW480 and LoVo cell lines were grown in 96-well plates for 48 h, as described. Medium was then replaced in the cell cultures with DMEM only (control group) or DMEM supplemented with 10 µM JZL184 (experimental group). Cells were then treated with varying concentrations of 5-fluorouracil (10, 100, 200 or 500 µM) for 48 h. Subsequently, 20 µl (5 mg/ml) MTT was added to each well for 4 h. The MTT formazan precipitate was dissolved in DMSO to 150 µl. The absorbance of each well was measured at a wavelength of 490 nm, and the cell proliferation inhibition rate was calculated using the following formula: Proliferation inhibition rate = [1 - value of the experimental group/average value of the control (5-fluorouracil only) group] × 100% (17).

The cells were collected and washed with phosphate-buffered saline (PBS). According to the manufacturer's protocol, samples were stained with Annexin-V APC (Nanjing KeyGen Biotech Co., Ltd., Nanjing, China) and PI (Sigma-Aldrich) for 30 min at room temperature in the dark. The cells were then examined using a BD FACSCanto II flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA).

Total mRNA was extracted from the cells using TRIzol^® (Takara Bio, Inc., Otsu, Japan). According to the manufacturer's protocols the RNA was reverse transcribed to cDNA using Primescript RT Master mix (Takara Bio, Inc.). RT-qPCR was used to detect the levels of mRNA in each sample, with glyceraldehyde-3-phosphate dehydrogenase (GAPDH) serving as an internal control. SYBR Green Master mix (Takara Bio, Inc.) and the StepOnePlus™ Real-time PCR system (Applied Biosystems; Thermo Fisher Scientific, Inc.) were used to conduct qPCR, according to the manufacturers' protocols. The qPCR cycling conditions were as follows: Denaturing the cDNA at 95°C for 30 sec; annealing at 60°C for 60 sec; and extension at 95°C for 5 sec. The reaction proceeded for 40 cycles, gathering the data of each sample, and the mRNA expression levels were evaluated using the 2^−ΔΔCq method (18). The primers were obtained from Genscript Corporation (Nanjing, China) and the sequences were as follows: Bcl-2, forward 5′-TTGCCCTCAAACAGAACAGC-3′ and reverse 5′-TGCAGCTCCTCTTGGCTAAA-3′; Bax, forward 5′-GGCCGGGTTGTCGCCCTTTT-3′ and reverse 5′-CCGCTCCCGGAGGAAGTCCA-3′; and GAPDH, forward 5′-GTTCCCACTGTCGATGTCTCA-3′ and reverse 5′-CCCTTCATCTTGCCCTCAGA-3′.

Cells were lysed with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) Sample Loading buffer (Beyotime Institute of Biotechnology, Haimen, China) supplemented with proteinase inhibitors (Roche Diagnostics, Basel, Switzerland), and the concentration of total protein was measured using a Bradford Protein assay kit (Beyotime Institute of Biotechnology). Proteins (5 µg/µl) were separated on 8 or 12% SDS-polyacrylamide gels by electrophoresis and were transferred to polyvinylidene fluoride (PVDF) membranes (EMD Millipore, Billerica, MA, USA). The PVDF membranes were blocked with 5% skimmed milk protein and 0.1% Tween 20 for 1 h. Primary antibodies were added and incubated overnight at 4°C, prior to the addition of goat anti-rabbit secondary antibody for 2 h at room temperature. The primary antibodies used in the present study, from Cell Signalling Technology Inc. (Danvers, MA, USA) and used at a dilution of 1:1,000, were: Rabbit anti-E-cadherin (cat. no. 3195), rabbit anti-vimentin (cat. no. 5741), rabbit anti-zinc finger protein SNAI1 (Snail; cat. no. 3879), rabbit anti-Bcl-2 (cat. no. 2870), rabbit anti-Bax (cat. no. 2772). Rabbit anti-GAPDH was supplied by Sigma-Aldrich (cat. no. G9545) and used at a dilution of 1:5,000. The secondary antibody used was polyclonal horseradish peroxidase-conjugated goat anti-rabbit IgG (dilution, 1:1,000; cat. no. BA1054; Boster Biological Technology, Pleasanton, CA, USA). Background autofluorescence was removed and blots were imaged using a ChemiDoc XRS+ imaging system and blots were analyzed with Image Lab software (Bio-Rad Laboratories, Inc., Hercules, CA, USA), with GAPDH acting as a reference protein. Relative expression of the relevant protein was calculated using the formula: Relative expression = (density value of the experimental group − value of GAPDH) / (density value of the control group − value of GAPDH).

SW480 and SW620 cells (1×10⁶) were treated with 10 µM JZL184 for 48 h and plated onto transwell cell culture inserts (8 µm microporous filters; BD Biosciences), which provided an artificial basement membrane, for 12 h. The upper chamber of the transwell system contained DMEM (in the case of SW480 cells) or RPMI-1640 (for SW620 cells); the lower chamber contained corresponding medium supplemented with 20% FBS. The migrating cells were stained with 0.1% crystal violet and five randomly selected areas were used to count the number of cells using an Olympus 1X71 inverted microscope (Olympus Corporation, Tokyo, Japan) (16).

Each experiment was performed at least three times independently. The data are presented as the mean ± standard deviation. SPSS v. 18.0 software (SPSS, Inc., Chicago, IL, USA) was used for all statistical analyses. Student's t-test was used to statistically analyze the data. P<0.05 was considered to indicate a statistically significant difference.

JZL184 was dissolved in DMSO and administered to the SW480, LoVo and HCT116 colon cancer cell lines. DMSO was used as a negative control. After a 48 h incubation, the inhibition ratio was measured by MTT assay. The inhibition ratio was non-significantly increased (P>0.05) in all three cell lines following treatment with 10 µM JZL184 (Fig. 1A). In addition, JZL184 (10 µM) was combined with various concentrations of 5-fluorouracil. In the colorectal cancer cells treated with JZL184, cell growth was decreased (P>0.05). The inhibition ratio was increased in the cells treated with both JZL184 and 5-fluorouracil, as compared with in the cells treated with 5-fluorouracil alone (Fig. 1B; P>0.05).

The present study demonstrated that JZL184 had marked effects on the proliferation of colorectal cancer cell lines. Proliferation is closely associated with apoptosis in cancer; therefore, apoptosis of the colorectal cancer cells treated with JZL184 was investigated by flow cytometry. Compared with the control group, the number of cells in early (Q2) and late (Q4) apoptosis was increased in the treatment group (Fig. 2A). The rate of apoptosis was increased in the three colorectal cancer cell lines (Fig. 2B). These results indicate that JZL184 may induce apoptosis via decreasing proliferation in colorectal cancer cell lines. In addition, the expression levels of MAGL were significantly decreased following treatment with JZL184 (Fig. 2C and D).

JZL184 promoted the apoptosis of colorectal cancer cells. Bcl-2 and Bax are closely associated cell apoptosis proteins (19); therefore, the effects of JZL184 on their mRNA expression levels were detected by RT-qPCR. The mRNA expression levels of Bcl-2 were significantly lower in the JZL184 intervention groups, as compared with the control group; however, the expression levels of Bax were significantly higher following JZL184 administration, as compared with in the control group in the three colorectal cancer cell lines (Fig. 3A). The protein expression levels of Bcl-2 and Bax were also detected following JZL184 administration. Consistent with the mRNA expression levels, following JZL184 intervention Bcl-2 protein expression levels were significantly decreased, as compared with the control group, whereas Bax protein expression levels were increased (Fig. 3B and C). These results indicate that JZL184 is able to regulate apoptosis via altering the mRNA and protein expression of Bcl-2 and Bax.

Migration is required for metastasis in colorectal cancer. To investigate whether JZL184 affected the migratory ability of colorectal cancer cells, SW480 and SW620 cells were selected. SW480 cells were initially derived from colon cancer tissue from a patient with Duke's type B colon cancer, whereas SW620 cells were initially derived from the metastatic lymph nodes of a patient with Duke's type C colon cancer, as listed by the American Type Culture Collection (www.lgcstandards-atcc.org/). The metastatic potential of these cells from two stages of colorectal cancer differs (20). Following administration of JZL184, the migration of SW480 and SW620 cells was significantly decreased. Compared with the SW480 cells, the suppression of SW620 cell migration was markedly increased (Fig. 4A).

EMT in epithelial tumor cells is closely associated with the process of tumor cell metastasis. In the present study, migration was observed to be inhibited by JZL184 in colorectal cancer cell lines; therefore, the protein expression levels of EMT markers were detected by western blotting. Following JZL184 administration, the SW480 and SW620 colorectal cancer cell lines exhibited significantly increased E-cadherin protein expression and decreased vimentin and Snail protein expression (Fig. 4B), thus indicating that MAGL may regulate migration via EMT.