Inhibition of AKT Sensitizes Cancer Cells to Antineoplastic Drugs by Downregulating Flap Endonuclease 1 Free

The leading causes of worldwide cancer-related mortality are lung cancer (1, 2). To date, chemotherapy and radiotherapy are considered the most effective intervention in lung cancer treatment. These antineoplastic compounds act through the process of interfering with cell division, initiating apoptosis, resulting in cell death. However, the effect of these treatments is weakened by the cancer cell's enhanced ability to resist cell apoptosis (3–5). Several reports stated the association of elevated DNA repair ability of cancer cells with the resistance to chemotherapy drugs (6–8). The initial discovery of Flap endonuclease 1 (FEN1), as a structure-specific endonuclease in long-patch base excision repair (LP-BER) pathway (9) in cellular DNA metabolism, plays a critical role in antineoplastic drug resistance (10). In DNA repair, FEN1 removes the flap structure created by LP-BER, allowing the reparation and survival of damaged cells. FEN1 E160D mutations result in autoimmunity, chronic inflammation and cancers (9). Our laboratory and other teams identified the FEN1 mutation in colorectal cancer cells and elucidated its role in cancer progression (10, 11).

Because fundamental functions of FEN1 include maintenance of genome stability, complete knockout of FEN1 activity leads to embryonic lethality (12, 13). Nevertheless, other DDR component knockouts such as ATR or PARP1 plus PARP2 targeted by agents tolerate embryonic lethality, in vivo, and clinical studies with appropriate dosing schedules. DDR component knockouts allow tumor-cell killing, especially when exploiting synthetic lethal concepts such as BRCAm or ATM deficiencies to spare the normal tissue (14–16).

FEN1 levels get elevated in rapidly dividing cells, including various types of cancer cells (17–20), in which FEN1 overexpression is associated with elevated malignancy and increased survival rates. FEN1 assists in determining the most effective treatment plan (21). The FEN1 expression levels have hypothesized to impact therapeutic response (22), as higher FEN1 expression often associates with increased malignancy and poor prognosis.FEN1 maintains genomic stability through the DNA replication and repair pathway, which makes FEN1 an ideal target for cancer therapy as it opens the opportunity to target both pathways in cancer cells (23). Increasing the amount of DNA damage leads to an increase in cytotoxicity and subsequent apoptosis. Furthermore, Ward and colleagues (24) showed that cells with certain DNA repair deficiencies remain particularly sensitive to FEN1 knockdown or inhibition (25). It has been reported that DNA repair pathways enable tumor cells to survive after DNA damage induced by antineoplastic drugs. Therefore, inhibitors of specific DNA repair pathways may prove to increase the efficacy of the treatment when used in combination with DNA-damaging chemotherapeutic drugs. A previous study revealed that silencing FEN1 leads to increased sensitivity to cisplatin in glioma and gastric cancer cells (26), which shows the importance of FEN1 in DNA damage repair. Our team reported that the suppression of FEN1 in cancer cells leads to the retardation of DNA replication, accumulation of DNA double-strand breaks (DSB), and subsequent apoptosis (22, 27). SC13, a small-molecular compound inhibits FEN1 activity, resulting in the sensitization of cancer cells to DNA damage-inducing therapeutic agents. Therefore, targetingFEN1 in cancer cells has the potential for effective cancer treatment (23, 28, 29).

Because FEN1 plays avital role in the DNA metabolism, it is under a complex and stringent regulation. Previous studies focused on the post-translational modifications (PTM) of FEN1 and the regulation of its protein stability and enzyme activities. Although the PTMs of FEN1 have been carefully studied, including methylation, phosphorylation, acetylation, and several other biochemical modifications (30, 31), the upstream signaling of FEN1 is not well understood. In our research, we identified AKT as an upstream regulator of FEN1, which proved that the inhibition of AKT activity can sensitize lung cancer cells to the effects of DNA damage-inducing drugs. Signal pathways involved in both tumorigenesis and suppression are the key to the development and improvement of pharmacological treatment strategies for cancer (32). AKT plays the central role in PI3K activation and has prompted a focus on targeting the PI3K pathway in cancer cells as a cancer treatment (33, 34). Reports on AKT, also known as PKB, remain constitutively active in non–small cell lung cancer, prostate cancer, and breast cancer (35, 36). Activated AKT can phosphorylate nuclear factor-κB (NF-κB), promoting NF-κB entering into the nucleus and regulating its target genes. NF-κB, a transcription factor regulating many genes, contains RelA(p65), NF-κB1(p50), NF-κB2(p52), RelB and c-rel. As early as a hundred years ago, a scientist named Rudolf Virchow—the father of modern pathology—suggested that inflammation might promote the development of cancer (37). Studies show that chronic inflammation caused 15% of malignant tumors (38), but the detailed mechanism is not known. Numerous studies show that chronic inflammation can activate factors released by NF-κB to promote tumor development (39). In 2009, Wang and colleagues (38) found that NF-κB could promote the inflammatory cells and intestinal epithelial cells transforming into colorectal cancer, but the mechanism is not known. In addition to colorectal cancer, NF-κB gets activated in lung cancer, bladder adenocarcinoma, multiple myeloma, liver cancer, esophageal adenocarcinoma, neoplasia, breast cancer, and leukemia (39). A new p53-dependent mechanism for blocking NF-kB survival pathways has been identified in cancer cells (40). Hence, NF-kB activation is closely related to tumor development.

To summarize, our study revealed two primary components, AKT inhibitors sensitizing A549 lung cancer cells to conventional chemotherapy, and identity of FEN1 as an AKT-regulated downstream target gene. AKT can phosphorylate IKB and the activated NF-κB can enter the nucleus, upregulate the transcriptional level of FEN1, increase the protein expression of FEN1, enhance the ability of DNA damage repair and improve the drug resistance of tumor cells. Our data proved that NF-κB/p65 directly and specifically bind to FEN1 promoter. We hypothesize that the combination therapy of AKT and cisplatin can significantly increase DNA damage in FEN1 high expressed cancer cells, indicating this therapy as a potential strategy for cancer treatment.

All cell lines used in this study were from the ATCC and were cultured under conditions as directed by the product instructions.

PI3K inhibitor LY294002 (Cat. No. S1105) and NFKB inhibitor BAY 11-7082 (Cat. No. S2913) were ordered from Selleck Chemicals. The chemical structure of LY294002 and BAY 11-7082 were shown in Supplementary Fig. S1A and S1D, respectively.

The sensitivity to DNA damage-inducing reagents was determined by cell growth inhibition assays. First, cells were seeded (1,500/well). A549 lung cancer cells were then incubated (overnight, 37°C), and treated (1 hour, 37°C) with multiple dilutions of LY294002 and cisplatin. Then, the cells were incubated (72 hours) under normal growth conditions (37°C, 5% CO₂). The number of viable cells was determined by the CellTiter 96 AQueous one-solution cell proliferation assay (Promega). At least four replications were performed for each group to ensure accuracy. The mean sensitivity level of each group was obtained and expressed as a percentage of growth with respect to the untreated control group (10, 41).

Tissues fixed in 10% formalin. Paraffin-embedded sections from tissue specimens were de-paraffinized, then heated at 97°C in 10 mmol/L citrate buffer (pH 6.0) for 20 minutes for antigen retrieval. Primary antibodies used in immunocytochemistry analyses were specific to the proteins being identified. Immunoreactivity was analyzed by estimating the percentage of cells showing characteristic staining and the intensity of staining present.

Antibodies used in this series of experiments: Anti-p53 antibody (SC-126, Santa Cruz Biotechnology), anti–caspase-3 (SC-7148, Santa Cruz Biotechnology), anti-vinculin antibody (MAB3574, Millipore), anti-FEN1 (70185, Gentex), anti-Tubulin (AM103a, Bioworld), anti–γ-H2AX (ab2893, Abcam), anti–cleaved Caspase-3 antibody (#9661, Cell Signaling Technology), anti-53BP1 (SC-22760, Santa Cruz Biotechnology), anti–P-AKT-S473 (CST#4060P), anti–AKT-pan (CST#4691P), anti-p65 (CST#8242S), anti–P-p65 (CST#3303T), anti–P-p53(S15, CST#9286P), anti–Ki67 (H-300, SC-15402), anti-ubiquitin (FC-76, SC-9133), anti–His-tag (BS-0287R), Alexa Fluor 488 goat-anti–rabbit A-11008 (Life Technologies), Alexa Fluor 594 donkey-anti–rabbit (R37119, Life Technologies).

Cells were cultured in 6-well plates containing acid-treated cover slides and incubated overnight. The cover slides were then washed with PBS, fixed with 4% formaldehyde in PBS for 30 minutes and washed with PBS again. Triton X-100 (0.05%) was added for 10 minutes to permeate the cells. Slides were blocked with 3% BSA then incubated with the primary antibody overnight at 4°C. Next, the slides were washed and incubated with the secondary antibody conjugated FITC. The slides were washed again with PBS, then stained with DAPI for 10 minutes. The mounted slides were viewed with a Nikon 80I 10-1500X microscope and images were captured with a camera.

All animal experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Female BALB/c nude mice between the age of 5 and 6 weeks were used in this study. A549 cells (2 × 10⁶) suspended in 100 μL serum-free medium were inoculated subcutaneously. Approximately 2 weeks later, when the average tumor volume reached 70 mm³, the mice were randomly divided into groups. PI3K inhibitor (50 mg/kg mice body weight) and cisplatin (25 mg/kg mice body weight) were administered intraperitoneally daily for 5 consecutive days. Tumor size was measured with slide calipers and volumes were calculated as (L × W²)/2, where L and W are the major and minor diameters, respectively. All mice were humanely euthanized when the cancer volumes of the control mice reached 1,000 mm³. All mice were housed and maintained under standard NIH protocol.

Cells were cultured in 6-well plates containing acid-treated cover slides and incubated overnight. The cover slides were then washed with PBS, fixed with 4% formaldehyde in PBS for 30 minutes and then washed with PBS again. Triton-X-100 (1%) was added for 5 minutes to permeate the cells. Three percent H₂O₂ was then added for 10 minutes. Subsequently, cover slides were washed twice with ice-cold PBS. Cells were incubated with TdT marker solution at 37°C for 1 hour and then gently washed with PBS 3 times. Cells were incubated in the dark with 100 μL dyeing buffer solution for 30 minutes, washed with PBS and stained with DAPI.

The chromosome breakage assay was performed as described previously (42–44). Cells were collected and treated with colchicine to arrest cells in metaphase. Cells were incubated for 30 minutes at room temperature with hypotonic solution (75 mmol/LKCl), then placed in a 37°C water bath for 5 minutes and fixed with Carnoy's solution. The fixation process was repeated three times and a dropper was used to place cells onto a clean slide. The cell spread was incubated at 55°C overnight, then stained with Giemsa solution and checked for aberrant chromosomes under the microscope.

Lentivirus particles expressing FEN1 shRNA were generated by transfecting 293T cells with shFEN1 expressing plasmids onto packaging plasmids. The virus-containing medium was collected every 24 hours for 3 days. The A549 cells were incubated with the lentivirus-containing medium plus 4 μg/mL polybrene for 24 hours and were then selected for 72 hours in 1.0 μg/mL puromycin. All lentivirus particles were prepared by Guangzhou Fitgene Biotechnology CO, LTD.

Tumor cells cultivated in chamber slides were treated with PBS or a different adenoviruses. After 72 hours, cells were incubated with Hoechst 33258 (Molecular Probes, Eugene) for 15 minutes, then washed with PBS twice and lastly observed under a fluorescence microscope.

Cells were harvested from the plates and re-suspended in lysis inducing buffer. Protein concentrations were determined by the Bio-Rad protein assay system. Aliquots of cell extracts containing 30 μg proteins were separated by SDS-PAGE gel electrophoresis on an 8% to 12% gel and transferred to PVDF membrane. The membrane was blocked for 1 hour with blocking buffer (5% non-fat milk powder in TBST: 10 mmol/L Tris-HCl pH 8.0, 150 mmol/L NaCl and 0.05% Tween20). The membrane was incubated with primary antibodies, then secondary antibodies, which were detected by chemiluminescence, using ECL western blot detection reagents (Pierce Biotechnology).

The electrophoretic mobility shift assay (EMSA) was performed as described previously by using purified recombinant NF-κB/p65 protein and the FAM-labeled double-strand DNA in a buffer containing 25 mmol/L Tris/HCl (pH 8.0), 1 mmol/L DTT, 5% glycerol, 0.25 mg/mL BSA, 50 mmol/L NaCl and 0.2 mmol/L EDTA. The mixtures was assembled on ice and pre-incubated for 10 minutes. Then incubated for 10 minutes at 37°C. At last, the mixtures were separated by 6% native polyacrylamide gel and analyzed by Odyssey FC.

Chromatin immunoprecipitation (ChIP) assay was according to the ChIP kit protocol (Beyotime). The rabbit anti-NF-κB/p65 antibody was purchased from Cell Signaling Technology. Rabbit IgG conjugated with magnetic beads were purchased from Cell Signaling Technology as the negative control. ChIP primers for the FEN1 promoter in Supplementary Data.

Luciferase activity was assessed according to the Dual-Luciferase Reporter Assay protocol (Promega) using a Veritas 96-well Microplate Luminometer (Promega) with substrate dispenser (Promega). HEK293 cells were cotransfected with 500 ng pGL3-Promoter FEN1, 500 ng NF-κB/P65 or vector and 50 ng pRLTK in a 24-well plate. After 48 hours of transfection, the cells were harvested for firefly and Renilla luciferase activity assay. The Renilla luciferase activities were used to normalize the transfection efficiency.

Total mRNA was isolated using TRizol reagent (Life Technologies). Reverse transcription reaction was performed according to the manufacturer's instructions. qRT-PCR was performed by using a reaction mixture with SYBR qPCR SuperMix-UDG (Invitrogen). All the PCR amplification was performed in triplicate and repeated in three independent experiments.

The IP assay was performed as described previously (34). Cell extracts were diluted with IP buffer (50 mmol/L Tris–HCl pH 8.0, 100 mmol/L NaCl, 5 mmol/L MgCl₂, 1% NP-40). Antibodies were incubated with protein A/G agarose (SC2003) in advance, then they were added to the diluted cell extract. After an overnight incubation, the beads were washed with IP buffer and the immunoprecipitated proteins were analyzed by western blot.

Cells were plated in 6-cm dishes and incubated for approximately 15 days at 37°C. The cells were then washed with PBS and stained with 0.05% crystal violet. Stained plates were washed and dried before scoring the colonies.

The statistical significance of the difference between various groups in the same experiments was determined by ANOVA and Newman–Keuls test for multiple comparisons. In vivo survival curves were estimated with the Kaplan–Meier method by the log-rank test for pair-wise survival analysis. Statistical significance was assumed when P < 0.05. All experiments were repeated at least twice to confirm reproducibility. All data are displayed as mean ±SD.

To confirm the activity of AKT in lung cancer cells, we assessed the AKT activity by immunoblotting with specific antibodies to phosphorylate AKT-S473. Figure 1A and B shows the elevated phosphorylated S473 in A549 cells comparing with normal human lung embryonic fibroblast (HELF) cells. To investigate whether AKT inhibitor LY294002 (Supplementary Fig. S1A) could decrease the drug resistance of cancer cells, we exposed cancer cells to three treatments. The first treatment included the AKT inhibitor LY294002, the second with the chemotherapeutic agent (cisplatin or 5-FU) and the third with a combination of AKT inhibitor and chemotherapeutic agent. The combination of LY294002 and cisplatin resulted in enhanced cytotoxic effect within A549 lung cancer cells, indicating a dose- (Fig. 1C) and time-dependent effect (Fig. 1D). Also, A549-cisplatin IC₅₀ value decreased from 15 μmol/L (without LY294002) to 7.5 μmol/L (with LY294002), as shown in Fig. 1C. We measured the combinative cytotoxic effect of LY294002 and different concentrations of 5-FU on A549 cells (Fig. 1E), indicating a time-dependent manner of response. To confirm that the above results are not due to cell specificity, we repeated the experiment on MCF7, a breast cancer cell line with relatively higher AKT activity. As shown in Fig. 1F, the results indicate the significant cytotoxic effect of cisplatin enhancement, suggesting a phenomenon that is not specific to A549 lung cancer cells. In further experiments, LY294002 cannot enhance the cytotoxic effect of cisplatin in HELF cells with reduced AKT activity. Figure 1G indicates AKT activity indispensable to the enhanced cytotoxic effect of LY294002 and cisplatin. Figure 1H shows the typical cell morphology of A549 cells treated with both LY294002 and cisplatin. To verify the above data, we performed the colony formation assay. As shown in Fig. 1I and J, the combination of LY294002 and cisplatin significantly reduced colony formation efficiency in A549 cells.

Past works proved that DNA damage induced by cisplatin lead to the accumulation of unrepaired DNA intermediates and DNA DSBs. On the basis of this information, we wanted to investigate whether the LY294002 treatment would enhance the accumulation of DSBs induced by cisplatin. To test this hypothesis, we examined the foci accumulation of γH2AX, an established marker of DNA DSBs in cells carrying gene damage. As expected, the results showed an enhanced accumulation of γH2AX (Fig. 2A and B) in A549 cells treated with LY294002 and cisplatin. In subsequent analyses, the cell immunofluorescence staining of 53BP1, another established gene damage marker, confirmed the accumulated DNA damage (Fig. 2C and D). Early research reported that DNA damage induced by cisplatin would lead to the accumulation of chromosomal breaks. Thus, to test the impacts of LY294002 on DNA stability, we analyzed metaphase nuclei for chromosomal aberrations induced by cisplatin. Figure 2E and F represents cells treated with LY294002 and cisplatin exhibiting significantly elevated levels of chromosomal fragments and breaks. The above data prove that the combination of LY294002 and cisplatin can lead to accumulation of increased unrepaired DSBs and chromosome breakage in A549 cancer cells.

A549 cells harbor a mutant EGFR and elevated AKT activity but the p53 gene remained intact (45–47). From cisplatin stress, p53 gets activated and acts as a transcription factor, activating the apoptotic genes. Therefore, we speculated that LY294002 treatment could enhance the apoptotic response in A549 cells after cisplatin exposure. As expected, LY294002 treatment increased the protein expression level of phosphorylated P53 (Fig. 3A and B). In the downstream event of P53 activation, the cleaved caspase-3 was upregulated in A549 cells after cisplatin exposure (Fig. 3C and D). Subsequently, we performed TUNEL assay to verify the apoptosis (Fig. 3E and F). We observed a significantly higher percentage of apoptotic cells after treatment with LY294002 and cisplatin than that of single treatment. These above data suggested that the combination of LY294002 and cisplatin efficiently activated the P53-mediated intrinsic pathway of apoptosis in A549 cells.

To further investigate the antitumor effect of the combination of LY294002 and cisplatin in vivo, we established a tumor xenograft model using nude mice. When the average tumor volume reached 70 mm³, we injected the mice with intraperitoneal injections of LY294002, cisplatin or the combination for 5 consecutive days. As shown in Fig. 4A, the average tumor volume increased much more drastically in the single treatment group than in the combinative group. Figure 4B shows the representative pictures of the tumor xenografts in all four groups. Figure 4C documented the average weights of the tumors depicted in Fig. 4B. Together, the combinative injection of LY294002 and cisplatin suppressed the growth of the transplanted A549 cells more than either of LY294002 or cisplatin alone.

To further study the mechanisms of apoptosis induced by LY294002 and cisplatin in vivo, the tumor xenografts were cut into slides. As shown in Fig. 4D, H–E staining recorded representative results of the tumor xenografts. The immunohistochemistry (IHC) assay using Ki-67 antibody showed that the combination of LY294002 and cisplatin efficiently suppressed tumor progression. Terminal dUTP nick-end labeling (TUNEL) of fragmented DNA can provide information of DNA damage (48, 49). The TUNEL-positive cells staining confirmed the apoptotic response in A549 xenograft treated with the combination of LY294002 and cisplatin. The enhanced positive staining of 53BP1 and γH2AX also substantiated the increased accumulation of DNA damage in A549 xenograft treated with the combination of LY294002 and cisplatin (Fig. 4D). These above results suggested that the inhibition of AKT could augment the efficacy of cisplatin on the lung cancer xenograft mouse model.

Based upon the published data, base excision repair (BER) is one of the major repair pathways to remove DNA damage caused by endogenous or exogenous agents. LP-BER is the major pathway to repair damaged bases in nuclei and mitochondria. The defects in LP-BER can lead to DSBs and genomic instability. To investigate the specific mechanism of LY294002 enhancing the antitumor effect of cisplatin, we analyzed the key protein expression level of the BER pathway. As shown in Fig. 5A, in LY294002-treated A549 cells, FEN1 protein level decreased significantly. In contrast, DNA polymerase β (Pol β), APE1 and other enzymatic members of the LP-BER pathway showed no obvious change. The Q-PCR result of Fig. 5B showed that AKT activity inhibition also decreasedFEN1 mRNA level, implying that AKT transcriptionally regulated FEN1 expression. To confirm the above findings, we performed immune cell staining (Fig. 5C and D), the results indicating that FEN1 protein expression level decreased significantly in LY294002-treated A549 cells. Further experiments indicated this regulation of FEN1 by AKT could also be demonstrated in an animal model. As shown in Fig. 5E and F, LY294002 show antitumor effects by downregulating AKT and consequently decreasing FEN1 protein expression in vivo.

The next hypothesis, we wanted to test was whether FEN1/AKT interaction is the key to understand the antineoplastic drug resistant mechanism of the AKT signal. Prior research has established that cisplatin induced DNA damage can lead to the accumulation of unrepaired DNA intermediates and consequently, DNA DSBs (50). Therefore, we anticipated cells with FEN1 downregulation to show higher levels of DNA DSBs and less responsive to LY294002 treatment. To test this hypothesis, we established a FEN1 knockdown stable cell line using the lentiviral technique (Fig. 6A). As shown in Fig. 6B and C, A549 FEN1 knockdown cells are more sensitized to cisplatin. However, when administration of LY294002, or in combination with cisplatin, sensitivity level did not differ from cisplatin alone. In successive analyses of the foci of γH2AX, as shown in Fig. 6D and F, LY294002 treatment did not increase γH2AX accumulation induced by cisplatin in A549 FEN1 knockdown cells. The above data indicate that FEN1 plays a critical role in the antineoplastic drug resistance regulated by the AKT pathway. To confirm this finding, we examined the TUNEL assay. As shown in Fig. 6E and G, FEN1 knockdown cells showed a bigger percentage of apoptotic cells. In addition, LY294002 treatment did not exhibit a significant effect on enhancing the chemotherapy drug sensitivity of A549 FEN1 knockdown cells to chemotherapeutic agent cisplatin. These data suggested that LY294002 sensitize the cells with high FEN1 expression but not those with low FEN1 expression to cisplatin.

To confirm the above data, we established a FEN1-overexpressing stable cell line based on the lentiviral technique. As shown in Fig. 7A, we identified the His-tag–labeled FEN1 overexpression with antibodies specific to FEN1and His-tag, respectively. Next, we decided to address whether FEN1 overexpression was a sufficient alteration that would re-establish the drug resistance to the pharmaceutical cisplatin in A549 cells treated with LY294002. As shown in Fig. 7B and C, the exogenous FEN1 expression enhanced the drug resistance to cisplatin in A549 cells. To confirm the above data, we examined the expression level of γH2AX in A549 cells treated with LY294002 and cisplatin. As shown in Fig. 7D, DNA damage and p53 expression increased in A549 cells.

Reports state that the accumulation of DNA damage induced by cisplatin will consequently lead to chromosomal aberrations or breaks (7). To test the effect of FEN1 expression on chromosomal breaks induced by cisplatin, the metaphase nuclei were analyzed for chromosomal aberrations. As shown in Fig. 7E and G, FEN1-overexpressing cells exhibited significantly decreased levels of chromosomal fragmentation and breaks. To confirm these findings, we next examined the apoptotic changes by the TUNEL assay. As shown in Fig. 7Fand H, FEN1-overexpressing cells became more resistant to cisplatin treatment and showed less apoptosis under the combinative treatment of LY294002 and cisplatin. The accumulated data established that the FEN1 overexpression can partially reduce the apoptotic phenotype in A549 cells induced by the combinative treatment of LY294002 and cisplatin.

Next, we wanted to know the possible mechanism of the regulation of FEN1 expression by AKT signal. The immunoprecipitation results of Supplementary Fig. S1B showed that there is no direct interaction between AKT and FEN1 protein. Considering that the decreased expression level of FEN1 mRNA after AKT inhibitor treatment, we speculated that the regulation of FEN1 by AKT signal might take place on the transcriptional level, but not on post-translation level such as phosphorylation. Thus, there may exist some transcription factors, which can be phosphorylated by the kinase activity of AKT signal, to regulate the transcription level of FEN1.AKT activates NF-κB by phosphorylation of IkB kinase (IKK), and active NF-κB enters into the nuclear to regulate the transcription of target gene. As shown in Supplementary Fig. S1C, AKT activity inhibition reduced the phosphorylation of NF-κB/p65. Furthermore, NF-κB inhibitor BAY117082(Supplementary Fig. S1D) treatment significantly decreased the FEN1 expression in both protein (Supplementary Fig. S1E) and mRNA (Supplementary Fig. S1F) levels. In A549 cells treated with TNF-α, a cytokine that can activate NF-κB signaling pathway, FEN1 expression significantly increased in both protein and mRNA level, as showed in Supplementary Fig. S1G and S1H, respectively. To verify the above data, we activated NF-κB/p65 with TNF-α in A549 cells after treatment with LY294002. The results showed a higher expression level of FEN1 than that of the single treatment with LY294002, as shown in Supplementary Fig. S1I and S1J. The above data showed that activated NF-κB/p65 can upregulate FEN1 expression efficiently. The expression of APE1 and Polβ are not affected by NF-κB/p65, as shown in Supplementary Fig. S2, indicating that the activation is specifically. To further confirm the above results, we ordered AKT activator (SC79), and the results proved that AKT activation also induces FEN1 mRNA and protein expression specifically, as shown in Supplementary Fig. S3.

Our bioinformatics studies based on software prediction (LASAGNA-Search 2.0 and UCSC) predicted that there might exist a potential NF-κB–binding site on FEN1 promoter region, as shown in Supplementary Fig. S4A. To confirm this, we designed a primer of FEN1 promoter with a FAM that NF-κB/p65 may bind to, and purified NF-κB/p65 protein with a GST label. As shown in Supplementary Fig. S4B, we found that the DNA-binding ability increased gradually with the concentration of the NF-κB/p65 protein increased. Chip assay showed that inhibition of AKT activity could weaken the binding ability of NF-κB/p65 with FEN1 promoter significantly, as shown in Supplementary Fig. S4C. Consistent with the above results, the dual-luciferase reporter assay showed that the transcriptional activity of NF-κB/p65 could be specifically improved by p65 or TNF-a, as shown in Supplementary Fig. S4D–S4F. These above data strongly suggest that activated NF-κB/p65 can bind to FEN1 promoter to upregulate transcription of FEN1, which is regulated by AKT activity. All these above data revealed a novel drug-resistant mechanism mediated by the AKT–NFKB–FEN1 pathway, as summarized in Supplementary Fig. S5.

DNA flap endonuclease 1 (FEN1) has been reported to be involved in various DNA repair pathways, including base excision repair (BER), HR (51, 52) and mismatch repair (53). Moreover, FEN1 has also been shown to be involved in nucleotide excision repair (NER) by association with Ligase I in the final step of NER (54). Given the roles of FEN1 in various DNA repair pathways, it is reasonable to speculate that manipulatingFEN1 could be a potent strategy to alter cisplatin response to cancer cells.

The PI3K/Protein Kinase B (PKB/AKT) pathway presents an appealing target for cancer therapy (33, 34). Though AKT promotes cell survival, chemotherapy resistance, and radiotherapy resistance, the detailed mechanism remains unclear. The function of AKT is to phosphorylate downstream numerous target genes, including BAD, FKHRL1 and many more (55, 56). Reports stated that the sequential combination of 5-FU and LY294002 induce an enhanced cytotoxicity to overcome intrinsic and acquired resistance to the chemotherapeutic drug 5-FU (57). In addition, the increased expression of P-p53 and the diminished expression of bcl-2 was observed (58).

A study conducted at the Birmingham Veterans Affairs Medical Center examining the metastasis of colorectal cancer reported that higher frequency of Smad4 inactivation or loss of expression contributes to chemotherapy drug resistance. Loss of Smad4 in patients with colorectal cancer induced drug resistance to 5-FU–based therapy through activation of the AKT pathway. However, in the study, AKT inhibitors were able to sensitize these patients to 5-FU (59). TWIST1, a master regulator of EMT, has also been reportedly linked to the development of AKT activation and chemotherapy drug resistance in epithelial ovarian cancer (60). Studies involving patients with cisplatin resistant ovarian cancer demonstrated AKT activation leads to inhibition of apoptosis (58). Another study in lung cancer cells indicated that Period 2, a pivotal mammalian circadian clock protein, participates in AKT-mediated drug resistance in A549/DDP lung adenocarcinoma cells (61).

In summary, AKT phosphorylates IKK, and the activated NF-κB enters into the nucleus to upregulate the transcriptional level of FEN1 and increases the protein expression of FEN1 to enhance the ability of DNA damage repair, which finally improve the drug resistance of tumor cells. The accumulated data suggest the PI3K/AKT pathway as a valid therapeutic target and recommends that PI3K/AKT inhibitors, can be used in conjunction with conventional chemotherapy agents as potential strategy in tumor therapy. Our study also elucidates the mechanism of FEN1 regulation, an important component in the DNA base excision repair pathway. Thus, we suggest that targeting the PI3K/AKT pathway could improve the therapeutic efficacy in high FEN1 cancer phenotypes.

No potential conflicts of interest were disclosed.

Conception and design: H. Zhu, L. He, Z. Guo

Development of methodology: H. Zhu, Z. Guo

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): H. Zhu, T. Wu, S. Zhou

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): H. Zhu, T. Wu, W. He, L. Li, A.M. Edick, F.-Y. Pan, L. He, Z. Guo

Writing, review, and/or revision of the manuscript: H. Zhu, C. Wu, W. Xia, A.M. Edick, A. Zhang, Z. Hu, L. He, Z. Guo

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): C. Wu, S. Ci, Y. Zhang, J. Zhang, Z. Guo

Study supervision: Z. Hu

This work was supported by the National Natural Science Foundation of China (81872284), Changzhou Sci and Tech Program (CE20175035), Jiangsu Key Research and Development Program (BE2018714; to Z. Guo), National Nature Science Foundation (31701179), and China Postdoctoral Science Foundation (2016M591877; to J. Zhang).

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