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. Author manuscript; available in PMC: 2025 Aug 25.
Published in final edited form as: Cancer Lett. 2023 Nov 15;580:216493. doi: 10.1016/j.canlet.2023.216493

Elevated 2-oxoglutarate antagonizes DNA damage responses in cholangiocarcinoma chemotherapy through regulating aspartate beta-hydroxylase

Katsuya Nagaoka 1,3, Xuewei Bai 1, Dan Liu 1, Kevin Cao 1, Joud Mulla 1, Chengcheng Ji 1, Hongze Chen 2, Muhammad Azhar Nisar 2, Amalia Bay 1, William Mueller 1, Grace Hildebrand 1, Jin-Song Gao 1, Shaolei Lu 4, Hiroko Setoyama 3, Yasuhito Tanaka 3, Jack R Wands 1, Chiung-Kuei Huang 1,2
PMCID: PMC12372592  NIHMSID: NIHMS2103815  PMID: 37977350

Abstract

Cholangiocarcinoma (CCA) is resistant to systemic chemotherapies that kill malignant cells mainly through DNA damage responses (DDRs). Recent studies suggest that the involvement of 2-oxoglutarate (2-OG) dependent dioxygenases in DDRs may be associated with chemoresistance in malignancy, but how 2-OG impacts DDRs in CCA chemotherapy remains elusive. We examined serum 2-OG levels in CCA patients before receiving chemotherapy. CCA patients are classified as progressive disease (PD), partial response (PR), and stable disease (SD) after receiving chemotherapy. CCA patients classified as PD showed significantly higher serum 2-OG levels than those defined as SD and PR. Treating CCA cells with 2-OG reduced DDRs. Overexpression of full-length aspartate beta-hydroxylase (ASPH) could mimic the effects of 2-OG on DDRs, suggesting the important role of ASPH in chemoresistance. Indeed, the knockdown of ASPH improved chemotherapy in CCA cells. Targeting ASPH with a specific small molecule inhibitor also enhanced the effects of chemotherapy. Mechanistically, ASPH modulates DDRs by affecting ATM and ATR, two of the major regulators finely controlling DDRs. More importantly, targeting ASPH improved the therapeutic potential of chemotherapy in two preclinical CCA models. Our data suggested the impacts of elevated 2-OG and ASPH on chemoresistance through antagonizing DDRs. Targeting ASPH may enhance DDRs, improving chemotherapy in CCA patients.

Introduction

Cholangiocarcinoma (CCA) is a rare type of tumor with little ongoing research efforts, but it has a dismal prognosis with a 7-20% 5-year survival rate if the disease has spread outside the liver(1). CCAs are classified as three subtypes, including intrahepatic, perihilar, and distal CCA, based on their anatomical site of origin. Although recent whole genomic sequencing data has discovered several potential targets in intrahepatic CCA, such as IDH1 and FGFR2 mutations, the 5-year survival rate is not significantly improved in these patients (2, 3). In early studies, several clinical trials have been proposed aiming to target advanced CCA in all subtypes by using combinational therapies of two or three chemotherapeutic agents, including doxorubicin (Dox), cisplatin (Pt), 5-fluorouracil, and gemcitabine (GCB). Although some trials showed improvement in controlling tumor growth, the statistical analysis of overall survival showed no significant difference(4, 5). This raises questions as to why this ineffectiveness occurs in all subtypes of CCA patients.

There are multiple mechanisms associated with chemoresistance in malignant tumors, including inactivation of anticancer drugs, multi-drug resistance due to increasing the release of drugs outside the cell and reducing the absorption of drugs, inhibition of the cell death, changing drug metabolism, changing the chemotherapeutic agents' targets, and enhancing the DNA damage repair(6). All these mechanisms may directly affect the therapeutic effects of chemotherapy by modulating the availability and activity of drugs(6). The research progress of CCA chemoresistance mainly focused on changes in intracellular drug concentrations, which are finely controlled by drug uptake and efflux genes. Among the most common mechanisms accounting for multi-drug resistance is the presence of ATP-binding cassette (ABC) transporters, which include multidrug resistance protein 1 (MDR1) and breast cancer resistance protein (BCRP), encoded by the ABCB1 and ABCG2 genes, respectively(7, 8). These transporters are responsible for drug efflux. Therefore, overexpression of these proteins would lead to reduced drug bioavailability, resulting in drug resistance. However, their roles in CCA chemoresistance may not be conclusive, as controversial findings have been reported(9). Drug resistance of CCA tumors has also been linked to the impaired transporters responsible for drug uptake, such as organic cation transporter 1 encoded by the SLC22A1 (10, 11). However, the GCB is transported mainly via human equilibrative nucleoside transporter 1(12). Understandably, the impaired function of these transporters would result in reduced drug concentration inside tumor cells, dampening the therapeutic effects. Nevertheless, the combination of GCB and Pt has been demonstrated to decrease tumor growth and extend patients' lives for 3.6 months (4), indicating that these drugs are delivered into the cells and executing their functions of killing cancer cells. The newly completed trials also supported this statement. The NUC-1031 (13), which is a phosphoramidate modification of GCB, could overcome the transporter deficiency issue but still fail to provide further benefits when compared with GCB and Pt (14, 15). Under these circumstances, there must be other mechanisms, such as cancer stem cells, increased DNA damage repair, and reduced cell apoptosis, contributing to the chemoresistance developed in patients with advanced CCA. Furthermore, all these mechanisms are suggested to be associated with DNA damage responses (DDRs) identified in GCB and Pt treatments(16), thus indicating the importance of DDRs in CCA chemoresistance.

It was well characterized that IDH mutations occurred in 15-20 % of CCA patients(17) and that the mutations would produce the oncometabolite, 2-hydroxyglutarate (2-HG)(18). Besides, 2-HG could suppress the functions of 2-oxoglutarate (2-OG)-dependent enzymes(19). Interestingly, a recent study demonstrated the involvement of 2-HG in DNA repair(20), thus indicating the potential impact of 2-OG-dependent enzymes in the failure of chemotherapy in CCA patients. There are many 2-OG-dependent enzymes identified and investigated in cancer progression(21-24). Among them, ASPH has been demonstrated to be highly expressed in CCAs but is barely detectable in bile duct cells and hepatocytes(25). We have previously shown that the knockdown of ASPH could promote DDRs in hepatocellular carcinoma(26), suggesting the potential impact of 2-OG/ASPH signaling cascade in CCA chemoresistance. Thus, the current study aimed to determine the role of ASPH in CCA chemoresistance.

Materials and Methods

Human Subjects

Serum from CCA patients (n=14) were obtained at Kumamoto University Hospital before receiving systemic chemotherapy and were used for the 2-OG measurements within one month. Treatment was classified as complete response (CR), partial response (PR), stable disease (SD), and progressive disease (PD) according to response evaluation criteria in solid tumors (RECIST) based on the first computed tomography evaluation after two courses of chemotherapy. Serum 2-OG levels were measured with the alpha-Ketoglutarate Assay Kit (MAK054, Merck) according to the manufacturer's instructions. Other blood tests were collected and performed at the clinical laboratory of Kumamoto University Hospital. Written informed consent was obtained from all patients. This study was approved by the Institutional Review Board of Kumamoto University Hospital (Approval no. 1094) and performed in accordance with the ethical principles associated with the Declaration of Helsinki.

Cell Lines and Reagents

Cell lines used in this study were either purchased or obtained from another laboratory through a material transfer agreement. The H1 (OCUCh-LM1-H1)(27), HuCCT1, OUMS-29, Rat BDE-Neu-CL#24, and SSP-25 cell lines were provided by Dr. Jack Wands. The HEK-293T (CRL-3216™, ATCC) cell line was purchased from ATCC. The H1, HuCCT1, and SSP-25 cells were cultured in RPMI growth medium containing 10% fetal bovine serum (FBS), 2mM L-glutamine, and 100U/ml penicillin-Streptomycin. The OUMS-29 and HEK-293T cells were cultured in DMEM growth medium with 10% FBS, 2mM L-glutamine, and 100U/ml P/S. The plasmids used in this study were obtained from Addgene (Watertown, MA). The psPAX2 (Addgene plasmid # 12259) and pMD2.G (Addgene plasmid # 12260) were gifts from Didier Trono. pLKO.1-shRNA-Luc was a gift from Dr. Chawnshang Chang (28). The shRNA and gRNA targeting ASPH were purchased from Horizon Discovery (Waterbeach, United Kingdom) (RHS3979-201772964, GSGH11935-247585597). The lentiCRISPR v2 was a gift from Feng Zhang (Addgene plasmid # 52961).

Animal Experiments

Fisher-344 male rats (F344/NHsd, Harlan Laboratories, Indianapolis, IN) were employed for the rat intrahepatic CCA model. The intrahepatic inoculation of BDE-Neu-CL#24 cells (2×106 cells/ rat) and bile duct ligations were performed as previously described (29). Female NCI Ath/nu mice were purchased from Charles River Laboratories (Wilmington, MA) for the human CCA animal model. The human H1 CCA cells (2×106 cells/ mouse) were inoculated into the liver. As for both experiments, an analgesic with buprenorphine was injected on the day of surgery and for 2 days thereafter. Body weight was measured twice per week. All protocols were approved by the Institutional Animal Care and Use Committee (IACUC) at Rhode Island Hospital, and all experiments were conducted in accordance with the guidelines of this IACUC.

MTT assay

The cell growth assay was measured using 5mg/ml of Thiazolyl blue tetrazolium bromide (MTT; #M5655 MillporeSigma (Burlington, MA)). Cells were seeded in a 96-well plate at 3,000 cells/well density and allowed to grow as indicated. The MTT solution was added to the wells. After incubation for 1 hour, the crystals were dissolved with DMSO, and the absorbance was measured in a microplate reader at a wavelength of 595 nm with a reference at 690 nm.

Cell viability assay

For the viability assay, cells were seeded into 24-well plates and treated with or without Dox for 3 days. After fixation with 10% neutral buffered formalin for 20 minutes, 0.1% crystal violet solution was added to the well and incubated for 30 min. The stained cells were washed with distilled H2O, and 500μL of DMSO was added to the well and incubated for 1 hour. The absorbance was measured using a microplate reader at a wavelength of 570 nm with a reference at 690 nm.

Colony formation assay

A colony formation assay was performed as previously described(30, 31). In brief, CCA cells (2.5x103) were mixed with 0.8% agar to form a 0.4% agar mixture and placed on a 0.8% bottom agar. The cells were cultured for 3 weeks and treated with or without Dox. After fixation, formed colonies were stained with 0.5% crystal violet solution and then counted using Image J software (U. S. National Institutes of Health, Bethesda, Maryland, USA, https://imagej.nih.gov/ij/).

Comet assay

The comet assay was performed using the Comet SCGE assay kit following the manufacturer’s instruction (ADI-900-166, Enzo Life Science, Farmingdale, NY). In brief, cells at 1 x 105/ mL were combined with molten LMAgarose (at 37 °C) at a ratio of 1:10 (v/v), and 75 μL of the mixture was immediately pipetted onto the Comet Slide. Then, the Comet Slide was resolved using DNA electrophoresis for 20-40 minutes. The Comet Slide was imaged using a fluorescence microscope with the FITC filter. The Comet tail length was recorded in 75 cells and averaged per sample.

Immunoprecipitation assay

Immunoprecipitation experiments were performed using 1mg protein samples as described in the following study(32). Briefly, protein samples were incubated overnight with specific antibodies (1 in 100 μl solution) or nonrelevant IgG as a negative control at 4 °C. Then, 30μl protein agarose A/G (SC-2003, Santa Cruz Biotechnology, Dallas, TX) was used to incubate at 4°C for 1 hour to pull down the protein complexes. The precipitated protein complex pellets were washed with RIPA buffer 3 times. Then, 50μl protein sample buffer was added to the complexes, boiled for 5 minutes, and placed on the ice for 5 minutes before performing the immunoblotting assay.

Immunofluorescence assay

The HEK-293T cells were seeded on 6-well plates with cover glass (CGN-1763-N25, FisherScientific, Hampton, NH) placed in each well. The cells were fixed with 10% formalin (FisherScientific, Hampton, NH) for 30 minutes and were permeabilized with 0.5% Triton X-100 for 20 minutes. The cells were subsequently treated with 10% H2O2 for 20 minutes and blocked with 1% BSA for 30 minutes. Next, cells on the cover glass were incubated with ASPH (a gift from Dr. Jack Wands) and ATM antibodies (MAT3-4G10/8, monoclonal, MilliporeSigma, Burlington, MA). The cells were covered with parafilm, incubated 4°C overnight, and washed with PBS for 3 minutes. The cells were incubated with the secondary antibodies for 1 hour (VECTOR, Newark, CA). The nuclear staining dye, 4'6-diamino-2-phenylindole (DAPI, VECTOR), was added to each well. The cover glass containing the cells was placed on an object slide for analysis and imaged by the laser scanning immunofluorescence microscope (Olympus).

Immunoblotting assay

Protein samples were extracted by lysing cells and tumor tissue samples with RIPA buffer (AAJ63324AK, Thermo Scientific, Waltham, MA). Protein concentrations were quantified using the Pierce™ BCA Protein Assay Kit (#23225, Thermo Scientific, Waltham, MA). A 50μg protein sample was used for all immunoblotting experiments. For ASPH, α-tubulin, cleaved PARP, GAPDH, HA tag, Histone H3, and Myc tag, 10% SDS-PAGE gels were used for the assay. For γH2aX examination, 12% SDS-PAGE gels were used. For all other targets, 6% SDS-PAGE gels were used for determining expression levels. The secondary antibodies were anti-rabbit-HRP and anti-mouse-HRP prepared in 2.5% non-fat milk in PBS with 0.5% tween 20. Images were taken using the ChemiDoc MP imaging system (Bio-Rad, Hercules, CA).

Lentivirus production

To produce lentivirus for ASPH knockdown and knockout, HEK-293T cells were plated into a 10cm dish 24 hours before transfection. To perform transfection experiments, the combination of PsAX2, pMD2G, and shRNA-ASPH (ratio=1:2:5) were transfected into HEK-293T cells using TransIT®-LT1 (MIR 2300, Miru, Marietta, GA). Following 48- and 72-hours post-transfection, lentivirus supernatant was collected and filtered using a syringe with a 0.45 μm syringe filter. The subsequent lentivirus supernatant was diluted with cell culture medium (ration 1:2) and used to infect target cells in the presence of 8 μg/ml polybrene (Hexadimethrine bromide, H9268, MilliporeSigma, Burlington, MA). Cells were used for the experiments for 72 hours post-infection.

Statistical analysis

The student t-test was used for statistical analysis between the two groups. The one-way analysis of variance was used for statistical analysis among three or more groups.

Results

Elevated 2-OG is correlated with poor chemotherapeutic responses.

To determine how 2-OG levels impact the effects of chemotherapy, we investigated the serum 2-OG levels of intrahepatic cholangiocarcinoma patients prior to anticancer drug therapy and their association with post-treatment efficacy. The patient serum samples collected before the start of chemotherapy were used for the measurements within one month. Based on the results of post-treatment imaging evaluation, treatment responses were classified as Partial response (PR), Stable disease (SD), and Progressive disease (PD). Patient information (Table S1), clinical characteristics of the PD and PR/ SD groups (Table S2), and the correspondence between 2-OG levels and specific clinical data for all patients (Table S3) are shown. Serum aspartate aminotransferase (AST) was significantly higher in the PD group than the PR/SD one (Fig. 1A). There was no significant difference in alanine aminotransferase (ALT) and total bilirubin (T-Bil) between the PD and PR/SD groups (Fig. 1B and C). Although AST was elevated in the PD group, the 2-OG levels were not significantly correlated with the AST and ALT levels (Fig. 1D and E). To investigate the direct link between 2-OG and the action of anticancer drugs, we measured 2-OG levels in the serum samples of PD and PR/SD groups, which we collected before chemotherapy. The 2-OG levels of patients classified as PD were relatively and significantly higher than those classified as the PR and SD (Fig. 1F). In line with the previous findings that inhibiting 2-OG dependent enzymes promoted chemotherapeutic responses(33-35), our results suggest that high 2-OG levels likely inhibit chemotherapy in CCA patients.

Figure 1. Elevated serum 2-oxoglutarate (2-OG) levels in CCA patients with progressive disease and liver function abnormalities.

Figure 1.

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CCA patients classified as progressive disease (PD) after receiving systemic chemotherapy (Gemcitabine plus cisplatin) demonstrating significantly higher serum 2-OG levels than those defined as partial response (PR) and stable disease (SD). Response to treatment was classified based on the initial assessment of the Computed Tomographic studies. (A) Serum AST levels, (B) Serum ALT levels, and (C) Serum T-Bil levels were determined in these patients. (D) Scatter plots evaluating the correlation between serum 2-OG and AST levels. (E) Scatter plots between serum 2-OG and ALT levels. The Pearson correlation coefficient (r), the coefficient of determination (R2), and the p-value are indicated. (F) Serum 2-OG levels of SD/PR and PD patients. AST, aspartate aminotransferase. ALT, alanine aminotransferase. T-Bil, total bilirubin. SD/PR, n=8; PD, n=6. Student t-test was performed and indicated when the p-value ≤ 0.05.

ASPH is associated with the negative impacts of 2-OG on CCA chemotherapy.

To determine whether elevated 2-OG can affect CCA chemotherapy, we adopted the Dox treatment, which has been used in the CCA clinical trial without significant improvement in CCA patients(5). We found that 2-OG treatment significantly antagonized the therapeutic effects of Dox since 2-OG promoted CCA cell growth in the presence of Dox (Fig. 2A). We then determined if 2-OG may affect DDRs in these cells by measuring the expression of γH2aX, the DNA damage marker(36). Interestingly, 2-OG treatment inhibited the Dox-induced γH2aX expression (Fig. 2B). We further evaluated the impacts of 2-OG on chemotherapy-induced DDRs in a non-malignant cell line that has low ASPH levels. Several concentrations were adopted, including two pathophysiological conditions-6 and 17μM(37), a condition possibly mimicking tumor microenvironment-100μM, and a super physiological dose- 1000μM of 2-OG. Consistently, 2-OG treatment suppressed γH2aX expression (Fig. 2C). To identify the potential molecule involved in this regulation, we focus on ASPH based on the rationale that we have previously suggested the involvement of ASPH in DDRs(26). Surprisingly, overexpression of ASPH could mimic the impacts of 2-OG on chemotherapy-induced DDRs (Fig. 2C), indicating the impact of ASPH enzymatic activity in this regulation. Indeed, overexpression of ASPH antagonized the DDRs induced by GCB plus Pt or Dox alone but not ASPH variant 3, which has no enzymatic domain (Fig. 2D). Together, our data demonstrated the involvement of 2-OG/ASPH in CCA chemoresistance.

Figure 2. 2-OG inhibited DNA damage responses produced by chemotherapy.

Figure 2.

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(A) Relative cell survival rate was determined in H1 cells treated with 0, 25, or 50 μM 2-OG in the presence of 1.25μM Dox for 2 days. **, p<0.01 by ANOVA analysis. (B) The protein expression levels of γ-H2aX and α-tub were examined in H1 CCA cells treated with 2-OG overnight and challenged with 1μM Dox for 2 hours. (C) ASPH, γ-H2aX, and α-tub were determined in 0, 6, 17, 100, 1000 μM 2-OG pre-treated HEK-293T cells expressing empty vector or ASPH for 24 hours before receiving DMSO control or 20μM CPT challenge for 2 hours. (D) γ-H2aX, GAPDH, and myc-ASPH expression was determined in HEK-293T cells expressing either ASPH (full-length) or ASPH variant form 3 (ASPHv3), which has no catalytic domain or enzymatic activity, in the presence of 4μM GCB plus 10μM Pt or 1μM Dox for 2 hours.

ASPH expression correlated with DDRs and cell death.

To further determine the impacts of ASPH on DDRs, we either overexpressed ASPH in non-malignant cell lines which have low ASPH expression or knocked down ASPH in CCA cell lines which have very high ASPH expression (30). As shown (Fig. 3A and 3B), overexpression of ASPH repressed DDRs in non-malignant cell lines, whereas knockdown of ASPH promoted DDRs in CCA cell lines upon challenging with DNA damage agents, Dox and camptothecin (CPT). We then directly measured the DNA damage breaks by utilizing the comet assay, which has a limit of sensitivity about 50 strand breaks per diploid mammalian cell (38). As expected, GCB plus Pt or Dox treatment could induce DNA damage breaks, as evidenced by the increased comet tail length. Intriguingly, the knockdown of ASPH substantially increased DNA damage breaks (roughly 400%~600%) compared with the control group (Fig. 3C and 3D). Subsequently, the increased DNA damage breaks led to increased cell death in ASPH knockdown CCA cells, whereas overexpression of ASPH inhibited cell death in non-malignant cells (Supplemental Fig. 1). These data further indicate that the expression of ASPH antagonized chemotherapy-induced DNA damage breaks which should subsequently result in chemoresistance.

Figure 3. ASPH expression negatively regulated DDR and inhibited cell death.

Figure 3.

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(A) γ-H2aX, α-tub, and ASPH were determined in SSP25 and HuCCT1 treated with shLuc or shASPH in the presence of 2.5μM Dox or 20μM CPT for 2 hours. (B) Immunoblotting results of ASPH, γ-H2aX, and α-tub in OUMS29 and HEK-293T cells expressing empty vector or ASPH and treated with as indicated in the presence of 20μM CPT for 2 hours. (C) The relative comet tail lengths of H1-shLuc, H1-shASPH, (D) SSP25-shLuc, and SSP25-shASPH CCA cells treated with control (Veh), 2.5μM Dox, 10μM Pt plus 1μM GCB for 2 hours were shown. *, p<0.05; **, p<0.01, when compared to the relevant controls.

Depleting ASPH enhanced the chemotherapeutic effects on CCA cells.

To determine if ASPH plays a critical role in the chemoresistance of CCA, we knocked down ASPH in two CCA cell lines and treated these cells with Dox. In line with our previous findings, knockdown of ASPH inhibited CCA cell survival (Fig. 4A and 4B). Interestingly, knockdown of ASPH could further promote the effects of Dox on CCA cells (Fig. 4B). The MTT assay is mainly used to determine mitochondria activity which can indirectly reflect cell survival rate. To directly determine cell survival by measuring cell numbers, we used crystal violet staining, which binds to protein and DNA. Our results showed that ASPH knockdown and Dox suppressed cell survival as indicated by the reduced absorbance of crystal violet staining. More importantly, the combination of ASPH knockdown and Dox has the highest suppression in cell survival (Fig. 4C), suggesting that inhibiting ASPH likely antagonized the chemoresistance of CCA cells. We then employed the soft-agar colony formation assay to evaluate the impacts of ASPH on chemotherapy. Similarly, the combination treatment has the most potent suppressive effects on anchorage-independent cancer cell survival in both cell lines (Fig. 4D). Taken together, these results suggest that ASPH depletion enhances the therapeutic effects of chemotherapy on CCA cells via promoting DDRs.

Figure 4. Knockdown of ASPH enhanced the chemotherapeutic effects in CCA cells.

Figure 4.

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(A) Immunoblotting results of ASPH and GAPDH in H1 and HuCCT1 cells stably expressing the control (shLuc) or knockdown of ASPH (shASPH). (B) MTT assays showed relative absorbance at 0, 1, 2, and 3 days in shLuc- and shASPH-H1 and HuCCT1 cells treated with or without 0.63μM Dox. (C) Cell viability (crystal violet staining) assays were performed in shLuc- and shASPH-H1 and HuCCT1 cells treated with DMSO or 0.5 μM Dox for 3 days. Representative photographs of crystal violet staining (left panel, 200x magnification, Scale bar 100μm, n=3) and quantification (right panel) by measuring the absorbance of crystal violet staining (See methods for more details). (D) Colony formation assays were conducted in shLuc- and shASPH-H1 and HuCCT1 cells treated with DMSO or 0.5 μM Dox for 3 weeks. Representative photographs of colony formation (left panel, 40x magnification, Scale bar 500μm). and quantification results (right panel). The colonies were counted using Image J software. (See methods for more information). The quantified results are presented as mean ± s.d. from 3 samples (A) or triplicate samples (B and C) using a student t-test. *p<0.05, when compared with the relevant control.

The ASPH-mediated suppressive effects on DDRs are highly associated with ATM and ATR kinases

Understanding the molecular mechanisms of action is essential in developing new target therapy. Although transcriptomic analysis will illustrate ASPH indirect targets in DDR regulations, we examined whether ASPH may affect DDRs through directly regulating ATM and ATR kinases since they are two of the major kinases highly involved in Dox, Pt, and GCB associated DDRs(39). The ATM protein kinase activity is mainly controlled by the MRN complex, composed of Mre11, Rad50, and Nbs1(40). We investigated the protein expression of ATM and MRN complex in non-malignant cells manipulated with empty vector (EV) and ASPH. We also checked their expression levels in CCA cells transduced with shLuc and shASPH via a lentiviral system. Interestingly, neither ATM nor MRN complex protein expression levels were significantly influenced by ASPH expression (Fig. 5A-C). We further examined the protein expression levels of ATR, ATRIP, phosphor-ATRIP (p-ATRIP), and TopBP1, which are the major contributors to ATR kinase activation(41). The expression level of ATR kinase was not affected by ASPH, either. Surprisingly, TopBP1, p-ATRIP, and ATRIP were positively correlated with ASPH expression (Fig. 5A-C). Thus, it is very likely that expression of ASPH leads to upregulation of TopBP1, ATRIP, and pATRIP, which results in ATR kinase activation, in turn suppressing DDRs. However, our findings still could not exclude the involvement of ATM here. Thus, we treated ASPH overexpressing cells or ASPH knockdown cells with ATM (KU55933) and ATR inhibitors (VE-821) overnight before applying chemotherapy. Intriguingly, ATM and ATR inhibitors both reversed the suppressive effects of ASPH on DDRs in ASPH overexpressing cells (Fig. 5D), suggesting that ASPH could inhibit DNA damage via promoting ATM and ATR kinase activities. Under these circumstances, ATM and ATR inhibitors should either have no impact on or slightly enhance DDRs caused by ASPH downregulation. Indeed, ATM and ATR inhibitors did not reverse the effects of ASPH knockdown on chemotherapy-induced DDRs in CCA cells (Fig. 5E).

Figure 5. ASPH inhibited the chemotherapeutic effects by targeting the ATM and ATR pathways.

Figure 5.

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ASPH, ATM, Mre11, Rad50, Nbs1, ATR, TopBP1, p-ATRIP, ATRIP, and α-tub were determined in (A) HEK293T-EV, HEK-293T-ASPH, (B) HuCCT1, and (C) H1 CCA cells transduced with shLuc or shASPH in the presence or absence of vehicle control, 10μM Pt plus 4μM GCB and 1μM Dox for 2 hours. ASPH, γ-H2aX, and GAPDH were determined in (D) EV- and ASPH-HEK-293T as well as in (E) shLuc-SSP25 and shASPH-SSP25 cells pre-treated with 1μM VE-821 or 1μM KU-55933 for 24 hours and challenged with 1μM Dox or 10μM Pt plus 4μM GCB for 2 hours before harvest.

ASPH bound to ATM and the MRN complex to promote ATM activation.

We have previously demonstrated that ASPH localizes in the cell nucleus and cytoplasm(31). Through bioinformatics analysis (https://myahits.sib.swiss/cgi-bin/motif_scan) (Fig. 6A), we identified a bipartite nuclear localization signal (NLS) located between amino acids 315 and 332 of ASPH protein. We separated cytoplasm and nuclear fractions from non-malignant cells and determined the protein expression of ASPH, ATM, Mre11, Rad50, Nbs1, α-tubulin, and H3. α-tubulin served as cytoplasm fraction control, and H3 was a control for nuclear fraction. As suggested in previous studies(42), ATM and the MRN complex are mainly located in the cell nucleus. We also found similar phenotypes. Besides, we discovered that ASPH localizes in the nucleus (30%) and cytoplasm (70%) (Fig. 6B). Therefore, we investigated whether ASPH interacts with ATM and the MRN complex to affect ATM activation. Interestingly, we found that ASPH is bound to ATM and the MRN complex, which would very likely enhance ATM kinase activation (Fig. 6C and D). We also found that ASPH and ATM are co-localized in the cell nucleus (Fig. 6E), further supporting the involvement of ATM in ASPH-mediated DDRs in CCA tumors. Indeed, we found that ASPH knockout substantially inhibited ATM and ATR kinase activities upon Dox challenge, as evidenced by the reduced expression levels of the ATM/ATR substrates (Fig. 6F). Therefore, based on these data, it is very likely that ASPH modulates DDRs via affecting ATM and ATR kinase activities.

Figure 6. ASPH affected ATM activation likely through binding to ATM and the MRN complex.

Figure 6.

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(A) A nuclear localization signal was identified in the ASPH protein. (B) ASPH, ATM, Mre11, Rad50, NBS1, a-tubulin, and H3 were determined in the cytoplasm (Cyto) and nucleoplasm (Nu) fractions of HEK-293T cells treated with or without Dox. (C) HEK-293T expressing HA-tagged ASPH were treated with or without CPT for one hour and protein lysates were collected for immunoprecipitation (IP) with HA antibody. IgG served as a negative control. ATM and the MRN complex proteins were examined in the ASPH-protein complex. (D) ATM and the MRN complex were determined in HEK-293T cells treated with or without Dox for 2 hours. The protein lysates were subjected to IP with ATM antibody and the protein complex was analyzed using immunoblot (IB) as indicated. (E) ATM and ASPH were examined in HEK-293T cells transfected with human ATM and ASPH plasmids. DAPI was used for nuclei staining. The merged image indicates co-localization of ATM and ASPH in the cell nucleus. (F) ASPH, ATM/ATR substrates, and GAPDH were determined in HuCCT1 treated with CRISPR/Cas9-CTRL or CRISPR/Cas9-ASPH in the presence or absence of 1.25μM Dox for 0, 0.5, 1, 2, 3, 4, 5, and 6 hours.

Targeting ASPH improved chemotherapy in vitro and in vivo.

To evaluate the possibility of applying our findings in a clinical scenario, we used a small molecule inhibitor (SMI) of ASPH that targets its enzymatic activity. In the previous study, we found that the newly synthesized ASPH-specific inhibitor-MO-I-1151 had improved therapeutic effects compared with other ASPH-specific inhibitors(30). Thus, we evaluated the anti-tumor effect of the combination of MO-I-1151 and Dox using human CCA cell lines. The combination cooperatively suppressed cell survival (Fig 7A) and colony formation in soft agar (Fig 7B) in both CCA cell lines. As the in vitro data revealed that the functional inhibition of ASPH led to improved chemotherapy, we wanted to determine whether the findings are reproducible in preclinical CCA models. We evaluated the therapeutic effect of the combination of ASPH knockdown and Dox in a rat synergic CCA model. BDE-Neu CL24 cells with stably transduced shLuc or shASPH (Fig. 8A) were orthotopically transplanted into the liver following common bile duct ligation in Fisher 344 rats to generate CCA tumors as previously described(29). After establishing rat CCA tumors, Dox was intraperitoneally administrated at 1 mg/kg 3 times per a week for 18 days and the rats were sacrificed at 3 weeks. We found that shASPH plus Dox significantly reduced tumor weight when compared with the shLuc control or Dox alone group (Fig. 8B and C). In line with the in vitro data, ASPH knockdown enhanced DDRs in rat CCA tumors (Fig. 8D). To further validate the clinical application of this targeting strategy, we prepared a human CCA xenograft mouse model generated by surgical inoculation of H1 cells into the liver of nude mice. The compound MO-I-1151 and Dox were administered via intraperitoneal injection to the experimental mice. The treatment of MO-I-1151 plus Dox therapy produced the best suppression of mean tumor weight among all groups (Fig. 8E). These results suggest that targeting ASPH improved chemotherapeutic effects in CCA tumors.

Figure 7. Targeting ASPH enzymatic activity by using the small molecule inhibitor enhanced chemotherapeutic effects in vitro.

Figure 7.

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(A) Relative MTT absorbance and (B) soft-agar colony formation assays were examined in H1 and HuCCt1 CCA cells treated with DMSO, 5μM MO-I-1151, 0.5μM Dox, 5μM MO-I-1151 plus 0.5μM Dox, (n=6). Representative photographs of colony formation (left panel, 40x magnification, Scale bar 500μm, and quantification results (right panel). Values are means ±s.d. *, p<0.05, when compared with the relevant control, n=4 in soft-agar colony assay.

Figure 8. Targeting ASPH improved the effects of chemotherapy in vivo in relevant preclinical CCA models.

Figure 8.

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(A) ASPH expression was validated in rat BDE-Neu#CL#24 cells treated with shLuc and shASH. shLuc- and shASPH-treated rat BDE-Neu cells were inoculated into the rat livers. 4 days later, the experimental rats were given 1mg/kg Dox treatment 3 times a week for 18 days. (B) Representative tumor images of treated animals. (C) Tumor weights of rat CCA tumors treated as indicated were shown. shLuc+veh, n=7; shASPH+veh, n=6; shLuc+Dox, n=8; shASPH+Dox, n=7. (D) The expression of γ-H2aX was examined in CCA tumor samples derived from a rat CCA model treated as indicated. (E) Tumor weights of the human orthotopic xenograft CCA tumors treated with vehicle (n=7), MO-I-1151 (25mg/kg, every other day, n=7), Dox (1mg/kg, 3 times a week, n=7), Dox (1mg/kg, 3 times a week)+MO-I-1151 (25mg/kg, every other day) (n=6) for 20 days. *, p<0.05 and **, p<0.01; when compared to the indicated group.

Discussion

The current study demonstrated that elevated 2-OG levels were negatively associated with therapeutic responses in CCA patients receiving chemotherapy. The treatment of 2-OG antagonized the therapeutic effects of chemotherapy in CCA cells in vitro, possibly through targeting DDRs. Interestingly, overexpression of ASPH could mimic the 2-OG-mediated DDRs but not the ASPH variant 3, which has no enzymatic domain. Consistently, knockdown of ASPH promoted DDRs in CCA cells treated with chemotherapy, whereas overexpression of ASPH suppressed these regulations. Besides, knockdown of ASPH improved its suppressive effects of chemotherapy on CCA cell growth. Mechanistically, it was found that expression of ASPH positively correlates with TopBP1, ATRIP, and pATRIP, which can promote ATR activation. ASPH may have direct or indirect binding with ATM and the MRN complex, thus likely affecting ATM activation through protein-protein interaction. Indeed, diminished expression levels of ATR/ATM substrates were observed in ASPH knockout CCA cells. Also, ATM and ATR inhibitors could reverse the inhibition effects of ASPH on chemotherapy-induced DDRs. The small molecule inhibitor study further demonstrated that targeting ASPH enzymatic activity improved chemotherapy in CCA cells. More importantly, targeting ASPH could improve chemotherapy in vivo in two preclinical CCA rodent models.

The cancer initiation role of IDH1 mutation has been linked to the oncometabolite, 2-HG. It was found that 2-HG could inhibit several 2-OG-dependent enzymes (tumor suppressors), disrupting epigenetic modification, thus resulting in malignant transformation and initiation of cancer(19, 43). Identification of IDH1 mutations in CCA tumors has suggested the potential of treating CCA patients with IDH1 mutations using specific inhibitors. The results of the ClarIDHy Trial indicated that the IDH1 inhibitor Ivosidenib improved the median overall survival in CCA patients from 7.5 months (Placebo group) to 10.3 months (Ivosideib group). However, the overall 5-year survival was not significantly improved in the ClarIDHy Trial (3). Intriguingly, recent studies have uncovered the crucial functions (oncogenic) of 2-OG-dependent enzymes in cancer progression (22, 24, 44, 45). It was also found that the IDH1 mutation sensitizes malignant cells to alkylating chemotherapy agents via inhibiting the 2-OG dependent-dioxygenase(35). IDH1 mutation also alters DNA repair and sensitivity to DNA damage through epigenetically affecting the expression of protein kinase ATM in hematological malignancies (46). The 2-HG produced by mutant IDH1/2 could induce DDRs by targeting DNA repair pathways via repressing 2-OG-dependent enzymes(33). Consistently, the IDH mutation, through suppressing 2-OG-dependent enzymes, enhanced DDRs in the presence of other chemotherapeutic agents (34, 47). Together, inhibiting 2-OG-dependent enzymes by IDH1 mutation results in elevated DDRs, which are associated with the effectiveness of general chemotherapies. Although IDH1 mutation would promote the chemotherapeutic effects through enhancing DDRs, it must be noted that IDH1 mutations were linked to CCA malignant tumor initiation and the IDH1 inhibitor was demonstrated to improve progression-free survival in CCA patients having IDH1 mutation(48). In this case, we must further determine the roles of those 2-OG-dependent enzymes that have oncogenic functions in CCA chemotherapy since we want to improve chemotherapy in CCA patients by blocking the oncogenic functions of 2-OG-dependent enzymes rather than enhancing IDH1 mutation functions, which may result in tumorigenesis and malignancy of adjacent non-malignant cells. Interestingly, the findings of our study suggest that 2-OG elevation antagonizes chemotherapy in CCA cells, likely through activating ASPH enzymatic function. Targeting ASPH also improved the effects of chemotherapy in preclinical CCA models, thus demonstrating the proof-of-concept that targeting those oncogenes belonging to the 2-OG dependent enzymes may overcome chemoresistance in CCA patients.

There are several limitations to this study. The sample size of our clinical study is relatively small. Large clinical trials may be required to ensure these findings. Nevertheless, we found that 2-OG levels are significantly and negatively correlated with the responsiveness of chemotherapy in CCA patients. We did not use the combination of targeting ASPH together with the current chemotherapy, GCB and Pt, in preclinical models. However, our study demonstrated the proof of concept that inhibiting one of the 2-OG dependent enzymes with oncogenic functions improved the effects of chemotherapy. We did not systemically determine the involvements of all 2-OG dependent enzymes in chemoresistance in CCA tumors. Studying the roles of other 2-OG dependent enzymes in the failure of chemotherapy in advanced CCA patients may further illustrate the involvement of 2-OG in this regard.

In conclusion, we found that elevated 2-OG could suppress the therapeutic effects of chemotherapy in CCA tumors through antagonizing DDRs. The 2-OG-mediated DDRs are partly modulated by the ASPH’s enzymatic activity. Targeting ASPH improved chemotherapeutic responses in preclinical models, thus providing a novel combination of targeting ASPH together with chemotherapy in CCA patients

Supplementary Material

Supplementary material

Acknowledgement

This work was supported by the JSPS KAKENHI Grant Number 22K15971 and 20KK0185 to K.N., AASLD Pinnacle Research Career Development Award and NIAAA R21AA030335 to C-K. H., and institutional funds to JR.W. The study was partly supported by the NIH R01CA270795 to J.R.W. and R. T. Funding sources provide financial support to the current study but have not been involved in the writing of the manuscript and the decision to submit it for publication.

Footnotes

Conflict-of-interest statement:

The authors have declared that no conflict of interest exists

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