The present study was approved by the Research Ethics Committee of Hangzhou First People's Hospital (Hangzhou, China). A total of 94 CRC and paired non-tumorous tissues were collected from patients following surgery at the Department of Gastrointestinal Surgery, Hangzhou First People's Hospital. Written informed consent was obtained from all patients. Tumor and paired non-tumorous tissues were stored at −80°C.

HCT8, HCT116, SW480 and SW620 human CRC cell lines were obtained from the American Type Culture Collection (Manassas, VA, USA). The HEK293T human embryogenic kidney cell line was obtained from The Cell Bank of Type Culture Collection of Chinese Academy of Sciences (Shanghai, China). All cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) supplemented with 10% fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc.), 100 U/ml penicillin and 100 mg/ml streptomycin.

For ectopic expression of NAC1, NAC1 cDNA was cloned into the vector pLKO.1 (catalog no. 10878; Addgene, Inc., Cambridge, MA, USA). Lentivirals were constructed in HEK293T cells and cell lines were infected to establish NAC1 stable overexpression cells as described previously (25). The siRNAs targeting NAC1 (UGA UGU ACA CGU UGG UGC CUG UCA CCA and GAG GAA GAA CUC GGU GCC CUU CUC CAU) and HOXA9 (ACU ACU ACG UGG ACU CGU UC and AAU CAA CAA AGA CCG AGC AAA) were purchased from Genepharm Biotech (Taipei, Taiwan). A scrambled sequence was used as a negative control. Cell transfection was performed using Lipofectamine^® 3000 according to the manufacturer's protocol (Invitrogen; Thermo Fisher Scientific, Inc.).

Total RNA from tissue samples and cells was extracted using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.). First-strand cDNA was synthesized using the PrimeScript cDNA Synthesis kit (Takara Biotechnology, Co., Ltd., Dalian, China), according to the manufacturer's protocol. qPCR was performed using the SYBR^® -Green Master mix kit (Takara Biotechnology Co., Ltd., Dalian, China) and the Applied Biosystems 7500 Real-Time PCR system (Applied Biosystems; Thermo Fisher Scientific, Inc.). Cycling conditions were as follows: An initial predenaturation step at 95°C for 30 sec, followed by 40 cycles of denaturation at 95°C for 30 sec and annealing at 60°C for 30 sec. All reactions were performed in triplicate. The primers used for detecting NAC1 were as follows: Forward, 5′-AAGCTGAGGATCTGCTGGAA-3′ and reverse, 5′-CCAGACACTGCAGATGGAGA-3′. The primers used for GAPDH were as follows: Forward, 5′-ACCACAGTCCATGCCATCA-3′ and reverse, 5′-TCCACCACCCTGTTGCTGTA-3′. The expression levels of NAC1 were normalized to that of GAPDH in each sample using the 2^−∆∆Cq method (26).

Total proteins from tissues and cells were extracted using radioimmunoprecipitation assay buffer containing protease and phosphatase inhibitors (Beyotime Institute of Biotechnology, Haimen, China). For western blot analysis, 60 µg of total protein lysates underwent 10% SDS-PAGE and were subsequently transferred onto polyvinylidene difluoridemembranes (Bio-Rad Laboratories, Inc., Hercules, CA, USA) as described previously (27). The protein was detected using an Enhanced Chemiluminescence substrate (Beijing Transgen Co., Ltd., Beijing, China). Rabbit anti-NAC1 (catalog no. ab29047) and rabbit anti-HOXA9 (catalog no. ab83480) primary antibodies were purchased from Abcam (Cambridge, UK). A mouse anti-β-actin (sc-47778) primary antibody was obtained from Santa Cruz Biotechnology, Inc. (Dallas, TX, USA) and served as an internal control. Membranes were probed with primary antibodies diluted 1:1,000 at 4°C overnight, followed by a horse radish peroxidase-conjugated goat-anti-rabbit or anti-mouse secondary antibody (catalog no. ab6789; Abcam, Cambridge, UK) diluted 1:5,000 for 1 h at room temperature.

CRC and paired non-tumorous tissues were fixed in formalin, embedded in paraffin and sectioned into 4-µm thick slices. Following deparaffinization and rehydration, antigen retrieval was performed using boiling 0.01 M citrate buffer (Boster Systems, Inc., Pleasanton, CA, USA) for 30 min and endogenous peroxidases were blocked with 3% H₂O₂. Sections were washed three times with PBS and goat serum (Boster Systems, Inc.) was used to block nonspecific binding for 30 min. Tissues were subsequently incubated with an anti-NAC1 antibody (1:200 dilution; catalog no. ab29047; Abcam) at 4°C overnight followed by incubation with a peroxidase-labeled goat anti-rabbit secondary antibody (catalog no. BA1055; Boster Systems, Inc.) for 1 h at 37°C. 3′3-Diaminobenzidine (Boster Systems, Inc.) was used to visualize tissue antigens. Following this, the sections were counterstained with hematoxylin and dehydrated and observed under a microscope.

The indicated cells were treated with 5-FU (50 ng/ml) oroxaliplatin (10 µM) (Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) and cultured for 12 h at 37°C. To measure capase-3/7 activity of cells following treatment with a Caspase-Glo^® 3/7 assay kit (Promega Corporation, Madison, WI, USA) was used according to the manufacturer's protocol. Cell viability was evaluated using the Cell Counting kit-8 (Dojindo Molecular Technologies, Inc., Kumamoto, Japan). The absorbance was measured at a wavelength of 450 nm using a microplate reader. Experiments were repeated at least three times.

HCT116 and SW620 cells were transfected with NAC1 or control siRNAs. The cells were subsequently transfected with a pGL3 control vector or the pGL3-HOXA9 promoter vector and Renilla. A luciferase activity assay was performed using the Dual-Luciferase^® Reporter assay system (Promega Corporation) according to the manufacturer's protocol. Firefly luciferase activity was normalized to Renilla activity.

SPSS software version 21.0 (IBM SPSS, Armonk, NY, USA) was used for statistical analysis. Data are presented as the mean ± standard error of at least three experiments. Data were analyzed by an unpaired Student's t-test for comparison between two groups, or a one-way analysis of variance followed by Student-Newman-Keuls post hoc test for comparison between multiple groups. P<0.05 was considered to indicate a statistically significant difference.

To investigate the involvement of NAC1 in the progression of CRC, the mRNA expression levels of NAC1 in 30 CRC and adjacent non-tumorous tissues were analyzed by RT-qPCR. The results indicated that compared with non-tumorous tissue, NAC1 expression levels were significantly upregulated in CRC tissue (Fig. 1A; P=0.0008). Additionally, a GSE6988 dataset generated from the Gene Expression Omnibus database consisting of 28 healthy and 49 CRC tissues was investigated, and the mRNA expression levels of NAC1 were significantly increased in CRC tissues (Fig. 1B; P=0.004). Furthermore, western blot analysis and immunohistochemistry revealed that the protein expression levels of NAC1 were elevated in CRC tissue (Fig. 1C and D). The results indicated that the expression levels of NAC1 were increased in tumor compared with non-tumor tissues, implicating an oncogenic function for NAC1 in CRC.

Chemoresistance is a major challenge for CRC treatment; therefore, the present study investigated the potential function of NAC1 in CRC cells following chemotherapy. NAC1 was stably expressed in HCT8 and SW480 cell lines and western blot analysis was used to confirm the overexpression of NAC1 (Fig. 2A). The cells were subsequently treated with 5-FU and oxaliplatin at a range of doses. The concentrations of 5-FU were as follows: 1, 4, 16, 64 and 256 ng/ml, and the concentrations of oxaliplatin were 1, 2, 8, 32 and 100 µM. The results indicated that overexpression of NAC1 in HCT8 and SW480 cells significantly increased the resistance of cells to 5-FU and oxaliplatin-induced cell death (Fig. 2B). In addition, caspase-3/7 activity was significantly decreased following overexpression of NAC1. This suggested a low level of apoptosis, and was consistent with the cell viability assay (Fig. 2C). Taken together, these data suggested that NAC1 increased the resistance of CRC cells to cytotoxic drugs.

To further characterize the role of NAC1 in the regulation of CRC cell death, the present study transfected target-specific siRNA against NAC1 into HCT116 and SW620 cells. NAC1 siRNA led to a significant decrease of protein expression levels in the two cell lines (Fig. 3A). These NAC1-knockdown cells became sensitive to apoptosis induced by 5-FU and oxaliplatin (Fig. 3B), suggesting an anti-apoptotic role for NAC1 in CRC cells. In addition, the cytotoxic agent-induced activation of caspase-3/7 significantly increased following NAC1 knockdown (Fig. 3C). Collectively, these data indicated that knockdown of NAC1 expression resulted in restored sensitivity to 5-FU and oxaliplatin in CRC cells.

The present study investigated the potential mechanism underlying NAC1-mediated chemoresistance in CRC cells. NAC1 may transcriptionally regulate the gene expression of a variety of pluripotency factors and modulate stem cell properties. The HOX gene family was selected for investigation due to its involvement in mediating cell stemness. It was revealed that HOXA9 promoter activity was significantly decreased following knockdown of NAC1 (Fig. 4A). These findings indicated that HOXA9 may be a downstream regulator of NAC1. Following this, western blot analysis revealed that knockdown of NAC1 suppressed HOXA9 expression (Fig. 4B). Furthermore, overexpression of NAC1 was demonstrated to induce HOXA9 expression, which was subsequently silenced by siRNA (Fig. 4C). The caspase-3/7 activity of CRC cells overexpressing NAC1 was determined, following silencing of HOXA9 expression (Fig. 4D). The results revealed that the NAC1-induced reduction in cell apoptosis was reversed by silencing HOXA9 expression, suggesting that HOXA9 is a downstream regulator of NAC1-mediated CRC cell resistance to anticancer drugs.