key: cord-0913869-zxpmtnrx
authors: Yi, You-Cai; Liang, Rui; Chen, Xiao-Yu; Fan, Hui-Ning; Chen, Ming; Zhang, Jing; Zhu, Jin-Shui
title: Dihydroartemisinin Suppresses the Tumorigenesis and Cycle Progression of Colorectal Cancer by Targeting CDK1/CCNB1/PLK1 Signaling
date: 2021-11-02
journal: Front Oncol
DOI: 10.3389/fonc.2021.768879
sha: 922c3577fc3f7523a77d50b9321646bd6ae88797
doc_id: 913869
cord_uid: zxpmtnrx

Dihydroartemisinin (DHA), a well-known antimalarial drug, has been widely investigated for its antitumor effects in multiple malignancies. However, its effects and regulatory mechanisms in colorectal cancer (CRC) are still unproved. In this study, in vitro experiments including CCK8, EdU, Transwell, and flow cytometry analyses and an in vivo tumorigenesis model were conducted to assess the effects of DHA on the bio-behaviors of CRC cells. Additionally, RNA-seq combined with gene ontology and Kyoto Encyclopedia of Genes and Genomes analyses was used to obtain the targets of DHA, and these were verified by molecular docking, qRT-PCR, and Western blotting. As a result, we found that DHA significantly suppressed the proliferation, DNA synthesis, and invasive capabilities and induced cell apoptosis and cell cycle arrest in HCT116, DLD1, and RKO cells in vitro and in vivo. Further analyses indicated that the targets of DHA were predominantly enriched in cell cycle-associated pathways, including CDK1, CCNB1, and PLK1; and DHA could bind with the CDK1/CCNB1 complex and inhibit the activation of CDK1/CCNB1/PLK1 signaling. Moreover, cucurbitacin E, a specific inhibitor of the CDK1/CCNB1 axis, enhanced the inhibitory effects of DHA on DNA synthesis and colony formation in HCT116 and DLD1 cells. In short, DHA could suppress the tumorigenesis and cycle progression of CRC cells by targeting CDK1/CCNB1/PLK1 signaling.

Colorectal cancer (CRC), one of the most malignant tumors, accounts for approximately 10% of all annually diagnosed cancers and cancer-related deaths (1) . According to the International Agency for Research on Cancer, CRC is the third most commonly diagnosed cancer and the second leading cause of cancer death, with an estimated 1.9 million new cases and 0.9 million deaths in 2020 (2) . Although its overall incidence and mortality continue to decline in older age groups, the incidence of early-onset CRC is increasing at an alarming rate (3, 4) . Because of the absence of the specific and distinct symptoms, most patients are diagnosed at an advanced and unresectable stage, and their prognosis is not optimistic. Consequently, it is necessary to explore new treatment strategies for improving the prognosis of the patients with CRC.

Recently, traditional medicine extract has received great attention because of its wide range of functions. Researchers found that in addition to its well-known functions, many new features have also been discovered, such as anti-inflammatory, antibiotic, antivirus, and anti-oxidation, especially antineoplastic. For examples, rhein suppresses the malignancy of CRC in vivo and in vitro by targeting mTOR pathway (5) . Toosendanin inhibits the proliferation of ovarian cancer cells through caspase-dependent mitochondrial pathway (6) . Sanguinarine combats the activation of EphB4 and HIF-1a pathways induced by hypoxia to hinder the growth of breast cancer (7) . Dihydroartemisinin (DHA), one of the derivatives of traditional Chinese medicine Artemisia annua, has become an essential component of antimalarial treatment for a long time (8) . Increasing evidence shows that DHA possesses multiple pharmacological actions against inflammation, viral infections, and tumor progression (9) . DHA can ameliorate psoriatic skin inflammation by diminishing CD8 + T-cell memory (10) , modulate the mammalian target of rapamycin pathway (11) , and suppress lipopolysaccharide-induced acute kidney injury by inhibiting inflammation and oxidative stress (12) . DHA also inhibits JC polyomavirus replication (13) , prevents malaria among HIV-infected pregnant women (14) , and prevents SARS-CoV-2 replication (14) . In addition, numerous studies reveal that DHA can exhibit antitumor effects in a variety of tumors, such as epithelial ovarian cancer (15) , lung cancer (16) , bladder cancer (17) , breast cancer (18) , and CRC (19) .

In the present study, we also found that DHA could inhibit cell proliferation, DHA synthesis, and cell invasion and induce cell apoptosis and cycle arrest in CRC cells in vitro and in vivo. Mechanical investigations showed that the targets of DHA were predominantly enriched in the cell-cycle pathway including CDK1, CCNB1, and PLK1 and that DHA could bind with CDK1 and inhibit the tumorigenesis and cycle progression by targeting CDK1/CCNB1/PLK1 signaling, contributing to the suppression of CRC tumorigenesis.

Human CRC cells lines (HCT116, DLD1, and RKO) were obtained from the Fudan University Shanghai Cancer Center and our Laboratory of Gastroenterology and cultured in Roswell Park Memorial Institute (RPMI) 1640 medium that contained 1% penicillin-streptomycin solution and 10% fetal bovine serum. The cell culture environment was sterile, humidified, and thermostatic at 37°C with 5% CO 2 .

To determine the 50% inhibitory concentration (IC 50 ) of DHA, HCT116 cells were seeded in a 96-well plate and treated with various DHA concentrations for 48 h. After that, the OD 450 was measured by CCK8 (Dojindo, Tokyo, Japan) following the manufacturer's instructions. Additionally, the dose-response curves were plotted in GraphPad Prism 7.0, and IC 50 was calculated by non-linear regression analysis. CCK8 was also used to measure the cell proliferation of HCT116, DLD1, and RKO cells when treated with 10 and 20 mM of DHA for 0, 24, 48, 72, and 96 h.

HCT116, DLD1, and RKO cells were seeded into 6-well plates (2 × 10 3 cells per well), treated with different DHA concentrations after the cells were attached to the wall, and cultured for 7 days. Cells were fixed with 4% paraformaldehyde and stained with 0.1% crystal violet for 30 min. Colony formation assay was examined by counting the number of stained colonies.

The EdU assay was conducted to evaluate DNA synthesis of CRC cells. The HCT116, DLD1, and RKO cells were incubated with 25 mM of EdU for 2 h, followed by staining according to manufacturer's protocol using a Cell-Light EdU Apollo567 in vitro kit (RiboBio, China). Images were obtained by using a fluorescence microscope (Olympus IX70, Olympus Corporation, Japan) and analyzed by ImageJ software (NIH, USA).

The cell mobility assay was carried out in a wound healing system. The six-well plates seeded with 4 × 10 5 cells per well were cultured until the cells covered the entire orifice. Then, the confluent monolayer cells were scratched with a 200-ml pipette tip. The wells were washed with phosphate-buffered saline (PBS) two or three times, treated with various DHA concentrations, and then cultured in the incubator. Wound healing photographs were taken at the same area as the scratching position at 0, 12, and 24 h after scratching.

Migration and invasion assays were assayed with uncoated (for migration) and matrigel-coated (for invasion) Transwell chambers with polycarbonate membrane of 8 mM (Corning, NY, USA) and 12 mM (Jet Biofil, China). Then 5 × 10 5 CRC cells suspended in 200 ml of serum-free medium were added into the upper chamber, and 700 ml of 20% fetal bovine serum (FBS) medium was added into the lower chamber. Then, the cells were incubated for 48 h. Cells were fixed with 4% paraformaldehyde and stained with 1% crystal violet. Subsequently, the migrated and invaded cells were identified by microscopy, and the results were analyzed by counting at least three random fields at ×20 magnification. For the cell cycle analysis, HCT116, DLD1, and RKO cells were treated with different DHA concentrations for 48 h and detached with trypsin after washing with cold PBS. Then, 70% ice-cold ethanol was used for cell fixation at 4°C overnight. After that, the cells were used for cell cycle analysis by a DNA Content Quantification Assay (Cell Cycle, Solarbio, China). Briefly, the cells were incubated with 100 ml of RNase at 37°C for 30 min, stained with 400 ml of propidium iodide (PI) for 30 min at 4°C in the dark, and then tested by flow cytometry. The data were analyzed by FlowJo 10.4 software (Becton, Dickinson & Company, NJ, USA).

Total RNA was isolated from the HCT116 control group and DHA-treated group (20 mM) using TRIzol reagent (Invitrogen), according to the instructions. RNA integrity was assessed by Bioanalyzer 2100 (Agilent, CA, USA) with RIN number >7.0 and confirmed by electrophoresis with denaturing agarose gel. Poly(A) RNA was purified from 1 mg of total RNA using Dynabeads Oligo (dT)25-61005 (Thermo Fisher, CA, USA). Following purification, the mRNA was fragmented into small pieces using the Magnesium RNA Fragmentation Module (NEB, cat.e6150, USA) at 94°C for 5-7 min. Then the cleaved RNA fragments were reverse transcribed to create the final cDNA library in accordance with the protocol for the TruSeq RNA Library Prep Kit v2 (Illumina). After that, we performed the 2 × 150 bp paired-end sequencing (PE150) on an Illumina Novaseq ™ 6000 (LC-Bio Technology Co., Ltd., Hangzhou, China).

To identify the enriched functions of the differentially expressed genes (DEGs) in the DHA treatment group, Gene Ontology (GO) analysis was performed by using R software (version 3.6.3) in the following three independent categories: biological process (BP), cellular component (CC), and molecular function (MF). Moreover, Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis was performed using the R package cluserProfiler pathview to map the enriched pathways. The top 100 DEGs were listed in a heatmap by the heatmap package.

Total RNA was extracted from HCT116, DLD1, and RKO cells using TRIzol reagent (Invitrogen, Thermo Fisher Scientific, USA). The cDNA was synthesized from RNA by using the PrimeScript ™ Reverse Transcription Kit (TaKaRa, Bio Inc., Japan). SYBR ® Premix Ex Taq ™ II (TaKaRa, Bio Inc., Japan) and the QuantStudio ™ 7 Flex Real-Time PCR System (Applied Biosystems, USA) were used for qPCR analysis. The reaction conditions for amplification were set according to the instructions, and the results were assessed by the 2 −DDCt method. The primers used are listed in Table 1 .

Total proteins were extracted from the CRC cells using radioimmunoprecipitation assay (RIPA) lysis buffer with a protease inhibitor cocktail (Beyotime Biotechnology, China) and boiled with loading buffer. Then, the protein samples were separated on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels (Epizyme, Shanghai, China) and transferred onto polyvinylidene difluoride (PVDF) membranes (Millipore PVDF 0.45 mm, USA). The membrane was blocked with 5% non-fat milk. After being washed with TBST twice, the membranes were incubated with diluted primary antibodies (1:1,500), including anti-CDK1 (9116, CST, USA), anti-CCNB1 (12231, CST, USA), anti-PLK1 (4513, CST, USA), and anti-GAPDH (5173, CST, USA) at 4°C overnight. Then, the corresponding secondary antibodies were diluted 1:3,000 and incubated with membranes at room temperature for 1.5 h. Finally, the membranes were washed twice with TBST and further visualized with the enhanced chemiluminescence (ECL) Western blotting substrate solution (NcmECL Ultra, Biotech Co., Ltd) and analyzed with Image-Pro Plus 6.0 software.

The animal experiments were approved by the Ethics Committee of Shanghai Jiao Tong University Affiliated Sixth People's Hospital. For the construction of a subcutaneous xenograft tumor model, 2 × 10 6 HCT116 cells were trypsinized and suspended in 0.2 ml of PBS and injected into BALB/c nude mice (n = 6 mice per group). After being fed in specificpathogen-free animal houses for 2 weeks, each mouse received an intraperitoneal injection of saline, low DHA (25 mg/kg), or high DHA (50 mg/kg) each day. Their body weight and tumor volume were recorded every 3 days.

Tumor tissues from sacrificed mice were fixed in formalin and t h e n e m b e d d e d b y p a r a ffi n . H & E s t a i n i n g a n d immunohistochemistry (IHC) analysis of Ki-67 (ab16667, Abcam, USA), CDK1, CCNB1, and PLK1 index were carried out according to our previous study methods (20) .

TIMER2 (tumor immune estimation resource, version 2, http:// timer.cistrome.org/) was used for analyzing the expression 

Forward primer Reverse primer

profiling of CDK1, CCNB1, and PLK1 between the tumor tissues and corresponding normal tissues (21) . Additionally, with the complement of the Genotype-Tissue Expression (GTEx) database, we used the Gene Expression Profiling Interactive Analysis, version 2 (GEPIA2) tool (http://gepia2.cancer-pku.cn/#analysis) (22) to obtain box plots of the expression difference between tumor and adjacent normal tissues in colon adenocarcinoma (COAD) and rectum adenocarcinoma (READ), with the settings of log2FC (fold change) cutoff = 1, p-value cutoff = 0.01, and "Match TCGA normal and GTEx data." We used the UALCAN tool (http://ualcan.path.uab.edu/ analysis-prot.html), an interactive web resource for analyzing cancer Omics data, to conduct protein expression analysis of the Clinical Proteomic Tumor Analysis Consortium (CPTAC) dataset (23) . We explored the total protein or phosphoprotein expression of CDK1, CCNB1, and PLK1 between primary tumor and normal tissues in COAD.

The canonical SMILES and the 3D structure of DHA were obtained according to our previous study (24) . The crystal structure of the CDK1/CCNB1 complex (PDB ID:4y72) was downloaded from the RCSB PDB database (http://www.rcsb.org/). Then, we used AutoDock Tools (version 1.5.6) to convert the pdb of DHA, CDK1, and CCNB1 to the pdbqt format; and the virtual molecular docking was performed by AutoDock Vina (25) . Furthermore, the best docking pose was selected according to the score of binding energy. The docking image was acquired using PyMOL (version 1.7.2.1).

The results were statistically analyzed with GraphPad version 8.0 Prism (GraphPad Software, La Jolla, CA, USA) and SPSS 22.0 (IBM, SPSS, Chicago, IL, USA). Each experiment was repeated at least three times, and the continuous variables were presented as the mean ± standard deviations. Student's t-test was used for two groups of normally distributed data, and one-way ANOVA was used for comparisons among the three groups. Additionally, we used the chi-square test for comparison of counting data. A pvalue <0.05 was considered statistically significant.

The raw data files for RNA sequencing analysis have been deposited in the National Center for Biotechnology Information (NCBI) Gene Expression Omnibus (GEO) under accession number GEO: GSE185141. Figure 1A) . Based on its IC 50 , the cell viability of HCT116, DLD1, and RKO cells was assessed by CCK8 when treated with 10 and 20 mM of DHA at different time points. We found that DHA significantly inhibited the proliferation viability of HCT116, DLD1, and RKO cells in a time-and dose-dependent manner as compared with the control group ( Figures 1B-D) . Likewise, colony formation and EdU assays showed that DHA remarkably suppressed cell colony formation (Figures 1E, F) and DNA synthesis ( Figures 1G, H) in HCT116, DLD1, and RKO cells.

To verify the potential effects of DHA on the mobility capacity of CRC cells, a wound healing assay was conducted, which indicated that DHA led to a marked decline of migration ability at 12 and 24 h in a dose-dependent manner in DLD1 and RKO cells (Figures 2A-D) . The Transwell assay consistently demonstrated that DHA significantly reduced the migration and invasion abilities of HCT116, DLD1, and RKO cells ( Figures 2E, F) .

In order to explore the effects of DHA on cycle distribution and cell apoptosis in CRC cells, we conducted flow cytometry analysis and found that DHA treatment caused G 2 arrest with the percentage of HCT116, DLD1, and RKO cells increased in the G 2 phase ( Figure 3A ). In addition, the proportion of apoptotic HCT116, DLD1, and RKO cells was dramatically increased when treated with DHA as compared with the control group ( Figure 3B) . Collectively, we discovered that DHA treatment induced cell cycle arrest and cell apoptosis in CRC cells.

To delineate the genome-wide landscape of transcription changes in DHA-treated HCT116 cells, we applied genomewide RNA-seq analysis of DEGs between DHA treatment and control groups and identified a large number of altered genes in the DHA treatment group (p < 0.05, fold change >1), including 441 upregulated genes and 2,979 downregulated genes ( Figure 4A) . Then, both KEGG and GO analyses revealed that these DEGs induced by DHA treatment were predominantly enriched in the cell cycle pathway, including CDK1, CCNB1, and PLK1 ( Figures 4B, C) , which has been shown to be associated with the tumor progression and therapeutic results (26, 27) . It is known that CDK1 interacts with CCNB1 to form a complex, controlling the G 2 /M phase transition, and PLK1 is the target gene. In addition, qRT-PCR and Western blotting analysis showed that treatment with 10 and 20 mM of DHA decreased the expression of CDK1, CCNB1, and PLK1 in HCT116, DLD1, and RKO cells (Figures 4D, E) .

It has been reported that cucurbitacin E (CE) is a specific inhibitor of the CDK1/CCNB1 complex that can induce CRC cells accumulate in the G 2 /M phase (28) . Having proved that DHA induced cell cycle arrest in the G 2 /M phase and inhibited the activity of CDK1/CCNB1/PLK1 signaling, we further explored the synergistic effects of DHA and CE on cell cycle distribution, colony formation, DNA synthesis, and CDK1/ CCNB1/PLK1 signaling activity in HCT116 and DLD1 cells and found that single CE or DHA treatment induced cell cycle arrest in the G 2 /M phase, but DHA in combination with CE showed no difference in the effect on cell cycle distribution as compared with CE treatment alone ( Figure 5A ). DHA synergy with CE could result in more inhibitory effects on cell colony formation ( Figure 5B ) and DNA synthesis ( Figure 5C ) as compared with single CE or DHA treatment in HCT116 and DLD1 cells. Then, qRT-PCR and Western blotting demonstrated that DHA is synergistic with CE or that a single CE treatment could repress the activities of CDK1, CCNB1, and PLK1 in HCT116, DLD1, and RKO cells (Figures 5D, E) . 

The antitumor effect of DHA on CRC growth was further validated in vivo. After the subcutaneous xenograft tumor model in nude mice was established by using the HCT116 cells for 2 weeks, the mice were treated with saline solution, 25 mg/kg of DHA, or 50 mg/kg of DHA for 16 days. We found that the tumor volumes and weights in the DHA treatment groups were obviously lower than those in the control group ( Figures 6A-C) . Furthermore, IHC analysis demonstrated that the protein levels of Ki-67, CDK1, CCNB1, and PLK1 in DHA treatment groups were markedly decreased as compared with those in the control group ( Figures 6D, E) . H&E staining also showed the number of the tumor cells in DHA treatment groups was smaller than in the control group ( Figure 6D ).

We further applied the TIMER2 approach to analyze the expression of CDK1, CCNB1, and PLK1 across various cancer types in The Cancer Genome Atlas (TCGA) and found that CDK1, CCNB1, and PLK1 were remarkably upregulated in multiple tumor tissues as compared with the adjacent normal tissues ( Figures S1A-C) , indicating that the activity of CDK1/ CCNB1/PLK1 signaling was frequent even in human malignant tumors. Furthermore, we validated that the expression of CDK1, CCNB1, and PLK1 was increased in CRC tissue samples relative to the normal tissues in COAD and READ cohorts ( Figure S2A) . Similar results were shown for their protein levels in the CPTAC dataset in COAD cohorts ( Figure S2B ). According to TCGA dataset, we performed a gene set enrichment analysis (GSEA), which indicated that these three genes were fundamentally enriched in the cell cycle, DNA replication, and p53 signaling pathways ( Figure S2C ).

Molecular docking has been widely used to predict the binding interfaces between a target protein (enzyme) and drug molecules (ligands) through molecular techniques (29) . We used autodock VINA to perform molecular docking of DHA into the crystal structure of the CDK1/CCNB1 complex. As shown in the Figure 7A , the docking center was set at 30.317 Å × −71.665 Å × 184.638 Å. The affinity of the docking pose was −9.7 kcal/mol, which indicated that there was strong binding between DHA and the CDK1/CCNB1 complex.

Accumulating data indicate that DHA has multiple biological functions, including antimalarial, anti-inflammation, antifibrosis, antiviral, and antitumor activities (12, (30) (31) (32) . A previous study showed that DHA can play a protective role in initiation and development of colitis-associated CRC (19) . Moreover, endoscopic topical drugs Ankaferd hemostat also shows the anti-CRC effect, which suppresses the proliferation of CRC cells by regulating its cell metabolism and cell cycle (33) .

In addition, previous studies demonstrated that other antimalarial drugs also suppress the malignancy of CRC. Artesunate induces apoptosis and autophagy in CRC cells (34) , and amodiaquine remarkably restrains the proliferation of CRC cells (35) . Mefloquine inhibits the nuclear factor kappa B (NF-kB) signaling pathway and induces growth and apoptosis of CRC cells (36) . Chloroquine inhibits the autophagy to strengthen the therapeutic effect of sinoporphyrin sodium-mediated photodynamic therapy in CRC (37) . To obtain a comprehensive understanding of the effects of DHA on the BP in CRC cells, we performed a series of in vitro and in vivo experiments and found that DHA could suppress CRC cell proliferation, colony formation, DNA synthesis, and invasion abilities in a time-or dose-dependent manner in vitro and in vivo, indicating that DHA might be an effective treatment for CRC. Furthermore, we identified the targets of DHA by RNA-seq and GO and KEGG analyses, which showed that the DEGs induced by DHA were enriched in the cell cycle in CRC cells. Numerous studies have shown that DHA exerts its anticancer effects by inducing cell cycle arrest. For example, DHA causes a cell cycle arrest in the G 2 /M phase in head and neck carcinoma and epithelial ovarian cancer via decreasing the expression of FOXM1 (38, 39) , and it inhibits growth of esophageal cancer cells by blocking cycle progression at the G 2 /M phase (40) . Here, we also found that DHA could inhibit CRC tumorigenesis and promote the cycle arrest in the G 2 /M phase, suggesting that DHA might exhibit the anti-CRC effects by suppressing the cycle progression.

We then found that three genes, CDK1, CCNB1, and PLK1, were included in the DHA-regulated cell cycle pathway. CDK1 (also known as CDC2), the first member of cyclin-dependent kinase family, mediates multiple pathways to regulate cell division, cell autophagy, and mitochondrial respiration (41) (42) (43) . It has been proven that CDK1 is connected to poor prognosis and cancer progression. It can initiate tumorigenesis of melanoma with the cooperation of Sox2 (44) , and the stabilization of CDK1 is required for breast cancer cell proliferation (45) . CDK1 also regulates the proliferation of gastric cancer via binding with CCNB1 and CCNB2 (46) . It is well known that CDKs exert its function in cell cycle by binding with cyclins, including cyclin A, cyclin B, cyclin D, and cyclin E; and these cyclins regulate kinase activity in a timely manner (47) . CCNB1, mainly expressed in the G 2 phase, interacts with CDK1 to form a complex and regulates the transition from the G 2 phase of the cell cycle to mitosis (47) . The CDK1/CCNB1 axis promotes cancer progression in multiple tumors, including hepatocellular carcinoma, bladder cancer, and lung cancer (48) (49) (50) . Analogously, our results demonstrated that DHA treatment induced CRC cell cycle arrest in the G 2 /M phase and inhibited the activity of the CDK1/CCNB1 complex. PLK1 is one of the most investigated PLK family members, acting at the G2/M checkpoint. It can be activated by the CDK1/CCNB1 complex, forming a positive feedback loop (51) . Accordingly, we found that DHA suppressed CDK1/CCNB1/PLK1 signaling activation and blocked cell cycle progression in CRC cells. CE is a natural compound derived from the climbing stem of Cucumis melo L. It has been reported that CE specifically suppresses the activity of the CDK1/CCNB1 complex (28) , causing the cell cycle arrest in the G 2/ M phase in CRC. In the present study, we also found that a single CE treatment could induce the cell cycle arrest in the G 2 /M phase and reduce the activity of CDK1/CCNB1/PLK1 signaling. DHA used in synergy with CE treatment showed more inhibitory effects on CRC cell colony formation and DNA synthesis.

Our study demonstrated that DHA significantly inhibited the malignancy of CRC both in vivo and in vitro. Although DHA is primarily used as an anti-malarial drug, its other functions, such as anti-inflammatory, antivirus, and antitumor, are gradually being discovered and might be a new drug for these diseases. In all, we found that DHA suppressed the tumorigenesis and cycle progression of CRC cells by targeting the CDK1/CCNB1/PLK1 feedback loop ( Figure 7B ). Our findings might provide a novel strategy for the treatment of CRC.

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found in NCBI [accession: GSE185141].

The animal study was reviewed and approved by Academic Committee on the Ethics of Animal Experiments of Shanghai Sixth People's Hospital. The total protein expression of CDK1, CCNB1, and PLK1 were analyzed in COAD through the CPTAC dataset. (C) GSEA analysis of the enriched signaling pathways by these three genes. *P < 0.05, **P < 0.01, ***P < 0.001.

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The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.