DMEM and FBS were purchased from Gibco (Grand Island, United States). Dihydroartemisinin (DHA) was obtained from Must Biotechnology (Chengdu, China). Actinomycin D was purchased from Selleck Biotechnology (Houston, United States). NAC reagent, and ROS or CCK8 detection kits were purchased from Beyotime Biotechnology (Shanghai, China). pcDNA3.1–3×HA-TERT and pCMV-myc-SP1 plasmid were purchased from Hedgehobio (Shanghai, China). Lipofectamine 2000 reagent was provided by Invitrogen (Carlsbad, United States). The antibodies against SP1, Ki-67, and GAPDH were purchased from Cell Signaling Technology (Beverly, United States). The antibody against hTERT was purchased from Abcam (Cambridge, United Kingdom). Goat anti-Rabbit IgG was purchased from BOSTER (Wuhan, China).

Human esophageal squamous cancer cell lines Eca109, KYSE150, and TE1 were obtained from the Translational Medicine Research Center of North Sichuan Medical College and were cultured in DMEM medium containing 10% fetal bovine serum at 37°C in an atmosphere containing 5% CO₂.

Human esophageal cancer cell lines Eca109, KYSE150, and TE1 were seeded into a 96-well plate at 8×10³ cells per well in DMEM containing 10% FBS. After 12 h, the cells were treated with DMSO or different concentrations of DHA for 48 h, or DHA at 100 μM for different time points, respectively. After DHA treatment, 10 μl of CCK8 reagent was added to each well and incubated for 2 h at 37°C in an atmosphere containing 5% CO₂. The absorbance (A) was measured at 450 nm by a microplate reader (Sunrise™ microplate reader, TECAN, NC). All determinations were carried out in triplicate.

Esophageal cancer cells Eca109, KYSE150, and TE1 were seeded into 6-well plates at a density of 2×10³ cells per well, respectively. After 5 days, the medium was replaced with 2 ml of 10% FBS contained DMEM with DMSO or 100 μM DHA. The culture was terminated when clones were visible by naked eyes. Then cells were fixed with methanol for 10 min and stained with crystal violet staining solution for 10 min. Images were obtained using a GT-S650 scanner (Epson).

Esophageal squamous cancer cells in logarithmic growth phase were seeded into 6-well plates at a density of 5 × 10⁵ per well and treated with DMSO or 100 μM DHA after 24 h. Forty-8 h later, cells were resuspended in serum-free medium by digestion with 0.25% trypsin and adjusted to a cell density of 5 × 10⁵/ml. A total of 200 μl cell suspension was added to upper chambers and 500 μl DMEM supplemented with 10% FBS was added to the lower chamber. After incubating for 24 h, the upper chamber was removed, fixed with methanol and stained with crystal violet solution, subsequently photographed under a microscope.

Esophageal cancer cell lines were seeded into 6-well plates at a density of 5 × 10⁵ per well. The cell layer was scratched with a 10 μl pipette tip when the cell confluence was about 90%. Cells was washed with PBS twice and cultured with DMEM containing 1% FBS with or without DHA for 24 h. Scratched wound was monitored, and pictures were taken at the indicated time points.

Esophageal cancer cell lines were seeded into a 6-well plate at a density of 5 × 10⁵ per well and treated with or without DHA or NAC for 24 h. Subsequently, DCFH-DA was diluted by 1: 1,000 with DMEM without FBS and incubated with the cells for 30 min at 37°C. After an hour, fluorescence signals in cells were observed by fluorescence microscopy, and the fluorescence intensity was measured by flow cytometry.

Total RNA was extracted using TRIzol reagent according to the manufacturer’s instructions. Then 1 μg of RNA was reversed transcribed into cDNA by reverse transcription system kit. Gene-specific amplification was performed in Roche Lightcycler 96 real-time PCR system (LightCycler^® 96 System, Roche, United States). The reaction system consisted of 5 μl 2 × TB Green Premix Ex Taq II (Takara), 0.2 μl forward primer (10 nM, Sangon, China), 0.2 μl reverse primer (10 nM, Sangon, China), 0.5 μl cDNA and 4.1 μl ddH₂O. All measurements were performed in triplicate, the thermal cycling for indicated genes was 95°C for 30 s and followed by 35 cycles of 95°C for 5 s, 58°C for 30 s and 72°C for 20 s. The 2^−ΔΔCT method was used to calculate the relative expression of hTERT. Primers were used as following: hTERT forward primer: 5′-CCT​TCC​TCA​GCT​ATG​CCC​GGA​CCT-3′, hTERT reverse primer: 5′-ACA​CTT​CAG​CCG​CAA​GAC​CCC​AA-3′; β-actin forward primer: 5′-CAT​GTA​CGT​TGC​TAT​CCA​GGC-3′, β-actin reverse primer: 5′-CTCCTTAATGTCAC GCACGAT-3′.

Cells were lysed using RIPA lysis buffer containing protease inhibitors. A total of 40 µg of protein sample was used for each lane of an SDS-PAGE gel and transferred to a polyvinyldifluoride (PVDF) membrane. Afterward, the PVDF membranes were blocked with 5% skim milk for 1 h at room temperature and then probed with primary antibodies overnight at 4°C. Subsequently, PVDF membranes were incubated with secondary antibodies for 1 h at room temperature. The bands were detected by an enhanced chemiluminescence system (vilber fusion FX7, France).

Esophageal cancer cell lines were seeded into 6-well plates at a density of 3 × 10⁵ per well and transfected with pcDNA3.1–3×HA-TERT or pCMV-Myc-SP1 plasmids using Lipofectamine 2000 reagent in OPTI-MEM medium after 24 h. Six hours later, medium was replaced with DMEM containing 10% FBS for 48 h.

Male 4-week-old BALB/c nude mice weight around 18–20 g were purchased from Beijing Laboratory Animal Research Center (Beijing, China). Animal care and experiments protocol were approved by the Animal Ethics Committee of North Sichuan Medical College. All animals were kept in a standard environmental condition at the experimental animal center of North Sichuan Medical College (Temperature 25°C; Humidity 55%; Light/Dark cycle, 12 h light and 12 h dark). A total of 1 × 10⁶ Eca109 cells were implanted subcutaneously on the back of the nude mice (100 µl suspension). When the tumor volume of Eca109 xenografts reached about 100 mm³, the nude mice were randomly divided into two groups. The mice were injected intraperitoneally with DMSO or DHA every 2 days. Tumor length and width were measured with a vernier caliper, and tumor volume was calculated by the formula: Volume (mm³) = Length × Width²/2. On day 19th, the mice were sacrificed, and lung, liver, kidney, heart, and tumor tissues were collected. Tissues were fixed in 4% paraformaldehyde and embedded in paraffin for immunohistochemistry or HE stain.

Tissue sections from paraffin-embedded esophageal cancer xenograft tissues were immunostained. Briefly, after deparaffinization and hydration of xenograft tissue sections, endogenous peroxidase was blocked by incubation with 0.3% hydrogen peroxide at 37°C for 30 min. Subsequently, sections were incubated with citrate buffer to repair antigen, and blocked with 5% BSA for 30 min at 37°C. Primary antibody was incubated at 4°C overnight and the secondary antibody was incubated at 37°C for 30 min. Finally, DAB was used as the chromogenic reagent, and counterstaining was carried out using hematoxylin. Tissue sections were dehydrated and sealed and observed with a microscope and photographed.

After deparaffinization and hydration of paraffin-embedded xenograft tissue sections, sections were stained with hematoxylin for 5 min. Then, the sections were differentiated with 1% hydrochloric acid alcohol solution, stained with eosin for 3 min. Finally, sections were dehydrated with an ethanol and xylene gradient, and were observed under microscope after being sealed with a neutral resin.

All in vitro experiments were performed three times, and data were presented as the mean ± SD. Statistical differences were evaluated by Student’s t-test or one-way analysis of variance (ANOVA) using SPSS 22.0 software. Differences with p < 0.05 were considered statistically significant.

Previous studies revealed that mutations in the promoter region of hTERT contributed to tumorigenesis, whether dysregulation of hTERT contribute to esophageal cancer is limited. We investigated the expression of hTERT mRNA in esophageal cancer from TCGA database and found that hTERT expression was higher in esophageal cancer than that in adjacent tissues (Figure 1A). In our cohort (n = 32), we found that the expression of hTERT in esophageal cancer tissues was evidently promoted, which was consistent with TCGA database (Figure 1B). Moreover, the expression of hTERT was positively correlated with worse prognosis in esophageal squamous cancer patients from TCGA database (Figure 1C). Futhermore, we attempted to uncover the association of hTERT with clinicopathological characteristics including gender, age, tumor size, tumor location, stage and lymph node metastasis, but non of these abovementioned characteristics was associated with hTERT (p > 0.05) (Supplementary Table S1). In order to elucidate the role of hTERT dysregulation in esophageal squamous cancer, hTERT was overexpressed in esophageal cancer cell lines Eca109 and TE1 (Figure 1D), and found that force expression of hTERT could significantly promote esophageal cancer cell proliferation and migration in vitro (Figures 1E–H, Supplementary Figures S1A,B). Altogether, these results demonstrated that hTERT plays an oncogenic role in esophageal squamous cancer.

Our previous study showed that DHA induced cell cycle arrest in Eca109 cells in vitro and in vivo (Ma et al., 2020). Whether the inhibition effect is universal or if other mechanisms involve in DHA treated esophageal cancer cell lines is unknown. Hence, we investigated the effect of DHA on the proliferation and metastasis of esophageal cancer cells Eca109, KYSE150, and TE1. As shown in Figure 2A, DHA significantly inhibited the viability of esophageal cancer cell lines (Eca109, KYSE150, and TE1) in a dose- and time-dependent manner. Plate colony formation assay showed that 100 μM DHA evidently inhibited the proliferation of esophageal cancer cells (Figure 2B). Moreover, transwell assay and scratch experiment revealed that DHA significantly inhibited esophageal cancer cells migration compared with cells treated with DMSO (Figures 2C,D, Supplementary Figures S1C,D). Taken together, the abovementioned results showed that DHA significantly inhibit the proliferation and migration of esophageal cancer cells.

Cause hTERT plays an oncogenic role in esophageal squamous cancer, we wonder whether DHA regulates the expression of hTERT in esophageal cancer cell lines. We measured the expression of telomerase reverse transcriptase (hTERT) on mRNA and protein levels in Eca109, KYSE150, and TE1 cells treated with indicated concentrations of DHA. The results showed that DHA markedly downregulated the expression of hTERT on mRNA and protein levels in esophageal cancer cells in a dose-dependent manner (Figure 3A). Moreover, 100 μM DHA also inhibited the expression of hTERT in a time-dependent manner (Figure 3B). Furthermore, we elucidated whether overexpression of hTERT could attenuate the anticancer activity of DHA. We found that overexpression of hTERT in esophageal cancer cells could alleviate the inhibition of DHA on the proliferation and migration of Eca109 cells (Figures 3C–F, Supplementary Figure S1E). These results demonstrated that hTERT was an oncogene in esophageal squamous cancer and could be used as a therapeutic target of DHA.

We further explored the molecular mechanism by which DHA downregulates hTERT expression in esophageal cancer cells. It is well-known that regulation of gene expression is mainly at the transcriptional and post-transcriptional levels. However, it is unclear whether DHA regulates hTERT expression at the transcriptional or post-transcriptional level. We firstly measured the mRNA stability of hTERT in DHA treated esophageal cancer cells by using actinomycin D experiments, and found that the stability of hTERT mRNA was not significantly difference between cells treated with DHA and vehicle (Figure 4A). Therefore, we speculated that downregulation of hTERT by DHA might trough the post-transcriptional level. It has been reported that the core promoter region (−181 bp) of hTERT contains two c-Myc and SP1 binding sites, and c-Myc and SP1 have been verified to upregulate telomerase activity through promoting the transcription of hTERT (Kyo et al., 2000). Since c-Myc and SP1 are directly involved in the expression of hTERT, we further determined whether DHA regulates c-Myc and SP1 protein expression in esophageal cancer cells. As shown in Figure 4B, DHA could downregulate the expression of SP1, but not c-Myc (data not shown), in Eca109, KYSE150, and TE1 cells. Moreover, overexpression of SP1 in esophageal cancer cells significantly upregulated the expression of hTERT (Figure 4C). In addition, we found that overexpression of SP1 enhanced the proliferation (Figures 4D,E) and migration (Figures 4F,G, Supplementary Figures S1F,G) ability of Eca109 and TE1 cells. Meanwhile, overexpression SP1 could attenuate the inhibition effect of DHA on the proliferation and migration ability in Eca109 cells (Figures 4H–K, Supplementary Figures S1H,I). Altogether, these data indicated that DHA downregulated hTERT expression through inhibiting transcriptional factor SP1 in esophageal cancer cells, subsequently inhibiting the esophageal cancer cells proliferation and migration.

Some studies have reported that DHA exerted antitumor effects mainly through the induction of ROS production (Xu et al., 2016). Camptothecin (CPT) can promote hTERT expression and telomerase activity by inducing ROS production, leading to SP1 activation (Dilshara et al., 2019). In RD and Rh30 rhabdomyosarcoma (RMS) cells, Methyl 2-trifluoromethyl-3,11-dioxo-18β-olean-1,12-dien-3-oate (CF3DODA-Me)can downregulate the expression of SP1 by inducing ROS (Kasiappan et al., 2019). Our previous study also revealed that DHA induced ROS production in Eca109 cells (Ma et al., 2020), whether DHA can induce ROS in other esophageal cancer cells is unknown. Therefore, we determined ROS level in DHA treated esophageal cancer cell lines Eca109, KYSE150, and TE1 by fluorescence microscopy and flow cytometry. As shown in Figures 5A,B, DHA significantly induced intracellular ROS production in esophageal cancer cells, and the antioxidant NAC (N-acetyl-l-cysteine) could alleviate DHA induced ROS production. In addition, Western Blot results indicated that NAC could attenuate the downregulation of SP1 and hTERT protein expression caused by DHA (Figure 5C). These results verified that DHA inhibits hTERT transcription by inducing ROS production and down-regulating transcriptional factor SP1.

To investigate the anti-esophageal cancer effect of DHA in vivo, we established Eca109 tumor-bearing nude mouse models. As shown in Figure 6A,B, the tumor sizes were significantly decreased in the DHA-treated group compared to the DMSO-treated group. Moreover, the tumor weight was lower (Figure 6C) in DHA treated Eca109 xenograft mouse models. Furthermore, DHA evidently downregulated the expression of SP1 and hTERT on mRNA and protein level in tumor tissues (Figures 6D,E). The IHC results also elucidated that the expression of hTERT and SP1 significantly downregulated in DHA treated xenograft esophageal cancer tissues, as well as the proliferation marker Ki-67 (Figure 6F). Intriguingly, side effect of DHA on heart, liver, lung, and kidney tissues of xenograft mouse models was evaluated by HE staining, and the results revealed that DHA exhibit no significant toxicity on these key organs of the mouse model (Figure 6G). Taken together, these data suggested that DHA can inhibit the growth of esophageal cancer in vivo without significant toxicity.