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. 2022 Dec 7;6(1):100–114. doi: 10.1021/acsptsci.2c00186

Inhibition of Protein Disulfide Isomerase (PDIA1) Leads to Proteasome-Mediated Degradation of Ubiquitin-like PHD and RING Finger Domain-Containing Protein 1 (UHRF1) and Increased Sensitivity of Glioblastoma Cells to Topoisomerase II Inhibitors

Rima Mouawad 1, Nouri Neamati 1,*
PMCID: PMC9841782  PMID: 36654750

Abstract

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Glioblastoma (GBM) is the most aggressive brain tumor, and the prognosis remains poor with current available treatments. PDIA1 is considered a promising therapeutic target in GBM. In this study, we demonstrate that targeting PDIA1 results in increased GBM cell death by topoisomerase II (Top-II) inhibitors resulting in proteasome-mediated degradation of the oncogenic protein UHRF1. Combination of the PDIA1 inhibitor, bepristat-2a, produces strong synergy with doxorubicin, etoposide, and mitoxantrone in GBM and other cancer cell lines. Our bioinformatics analysis of multiple datasets revealed downregulation of UHRF1, upon PDIA1 inhibition. In addition, PDIA1 inhibition results in proteasome-mediated degradation of UHRF1 protein. Interestingly, treatment of GBM cells with bepristat-2a results in increased apoptosis and resistance to ferroptosis. Our findings emphasize the importance of PDIA1 as a therapeutic target in GBM and present a promising new therapeutic approach using Top-II inhibitors for GBM treatment.

Keywords: glioblastoma, protein disulfide isomerase, topoisomerase II inhibitor, UHRF1, ferroptosis

Glioblastoma (GBM), a grade IV glioma, is considered the most common and aggressive malignant primary brain tumor.1 The current therapeutic approach for GBM is surgical resection followed by radiation in addition to chemotherapy with temozolomide (TMZ), a DNA alkylating agent.2 However, only 7.2% of patients survive more than 5 years after diagnosis due to the development of resistance to therapy and rapid tumor recurrence.3 Therefore, there is an urgent need for the identification of new drugs or therapeutic combinations to improve treatment outcomes and increase patient survival.

Select cancers including GBM are characterized by elevated endoplasmic reticulum (ER) stress and activation of the unfolded protein response (UPR) as an adaptation strategy for survival. One of the outcomes of the UPR is upregulation of protein-folding ER chaperones such as the protein disulfide isomerase (PDI) family members.2,4,5 The PDI family of proteins generally reside in the ER and are best known for their oxidoreductase and molecular chaperone functions. A promising approach for cancer therapy is targeting the ER protein-folding machinery, which is frequently upregulated in tumors due to the high rate of protein synthesis.2 PDIA1, encoded by the P4HB gene, is the canonical PDI family member. PDIA1 is upregulated in various human cancers including GBM and is associated with tumor progression and patient survival, and thus considered a promising therapeutic target.2,4

Various PDIA1 inhibitors have been developed and studied in the context of cancer (e.g., PACMA31, BAP2, 35G8, T8, and CCF642),6−11 Huntington’s disease (LOC14),12,13 and thrombosis (bepristat-2a and isoquercitin).14,15 Some PDIA1 inhibitors such as PACMA31 and CCF642 are characterized as irreversible inhibitors through binding covalently to the PDIA1 conserved CGHC active site and are expected to interact with other PDI family members.6,11 PACMA31 was studied in the context of ovarian cancer where it inhibited ovarian tumor growth in vivo,6 and CCF642 exhibited anticancer activity in a mouse model of multiple myeloma.11 In contrast, other PDIA1 inhibitors such as BAP2, LOC14, bepristat-2a, and T8 are categorized as reversible allosteric inhibitors that affect substrate binding to PDIA1 and are presumably more selective for PDIA1 over other PDI family members.8,10,12,14 BAP2 showed antitumor activity on GBM both in vivo and in vitro and impacted the response of tumor cells to TMZ and radiation.7,8 Bepristat-2a was identified in the context of thrombosis where it inhibited platelet aggregation and thrombus formation in a mouse model of thrombosis.14 In addition, bepristat-2a was recently studied in the context of breast cancer where it showed antimetastatic activities.16 Although various PDIA1 inhibitors have been developed, none of them has been approved for clinical use. Therefore, further understanding of PDIA1 structure and cellular functions is important for developing new selective PDIA1 inhibitors that can reach regulatory approval.

ER stress or inhibition of PDIA1 has been shown to impact DNA repair pathways.7,8,17−20 For example, tunicamycin-induced ER stress results in defects in DNA damage repair through the downregulation of RAD51 and increased sensitivity of human lung cancer cells to ionizing radiation and cisplatin.20 Treatment of GBM cells with ER-stress-inducing agents results in the downregulation of various DNA repair proteins and sensitized cells to TMZ.18,19 Treatment of GBM cells with BAP2 or BAP2 analogs results in a decrease in the expression of DNA repair genes including RAD51.7,8 Besides, treatment of GBM cells with PACMA31 or knockdown of PDIA1 gene (P4HB) results in downregulation of DNA repair genes.7 Knockdown of P4HB results in a decrease in RAD51 levels and sensitizes GBM cells to radiation in vivo.17 These studies demonstrate a significant role of PDIA1 in DNA repair processes and response of GBM to chemotherapy and radiotherapy. Targeting DNA repair is considered a promising therapeutic strategy in GBM since multiple pathways are involved in repair of DNA lesions induced by TMZ and ionizing radiation.21,22

In this study, we performed combinations of PDIA1 inhibitor bepristat-2a with various DNA-damaging agents and observed that bepristat-2a is synergistic with Top-II inhibitors for cytotoxicity in GBM and other cancer cell lines. We discovered that PDIA1 inhibition results in proteasome-mediated downregulation of UHRF1, an oncogenic protein that is overexpressed in many tumors and is associated with tumor progression and poor prognosis. We observed that bepristat-2a is protective against ferroptosis and increases apoptosis induced by Top-II inhibitors. Our study provides a promising new combination therapy for GBM and a novel link between PDIA1 and the oncogenic protein UHRF1 that can be utilized as a biomarker when developing future PDIA1 inhibitors.

Results

Bepristat-2a Increases the Sensitivity of Tumor Cells to Topoisomerase II Inhibitors

Previously, we showed that PDIA1 inhibition increases the sensitivity of GBM cells to radiation.8,17 On the basis of our previous results, we hypothesized that inhibition of PDIA1 would increase the sensitivity of tumor cells to other DNA-damaging agents. We used bepristat-2a as an established reversible and selective PDIA1 inhibitor,14 and performed combination studies with various DNA-damaging agents using GBM cell line GB-1. The combination of bepristat-2a with Top-II inhibitors, doxorubicin, etoposide, and mitoxantrone, resulted in reduced clonogenic survival of GB-1 cells as compared to Top-II inhibitors alone (Figure 1A). Pretreatment of GB-1 cells with bepristat-2a resulted in more cell death in comparison to pretreatment of GB-1 cells with Top-II inhibitors (Figure S1A,B), indicating that PDIA1 inhibition sensitizes GBM cells to Top-II poisons. In addition, PDIA1 knockdown using siRNA resulted in increased sensitivity of GB-1 cells to etoposide and doxorubicin (Figure S1C,D). In contrast, bepristat-2a was not synergistic with Topoisomerase I (Top-I) inhibitors, camptothecin and irinotecan, or other DNA-damaging agents such as TMZ and oxaliplatin (Figure 1B). We tested multiple cancer cell lines to determine if the synergy between bepristat-2a and Top-II inhibitors is true in other cancer cell lines in addition to GBM. The addition of sublethal concentration of bepristat-2a (10 μM) significantly increases the sensitivity of ovarian carcinoma cells, OVCAR8, to etoposide with a more than 10-fold decrease in the IC50 value. (Figure 1C). In clonogenic assay, bepristat-2a showed strong synergy with etoposide and doxorubicin, but not camptothecin for inhibition of clonogenic cell survival (Figures 1D and S2A). The same results were observed using lung adenocarcinoma cells, A549, and colon cancer cells, HCT116 (Figures 1E and S2B,C). Interestingly, we did not observe a strong synergy between bepristat-2a and etoposide in normal human foreskin fibroblast cell line HFF-1 (Figure S2D) indicating that PDIA1 inhibition increases sensitivity to Top-II inhibitors selectively in cancer cells. In comparison to OVCAR8 (Figure 1C), bepristat-2a (10 μM) did not strongly affect the sensitivity of HFF-1 cells to etoposide (around 4-fold change in IC50) (Figure S2E). Collectively, these results suggest that PDIA1 inhibition increases the sensitivity of various cancer cells to Top-II inhibitors but not Top-I inhibitors or other DNA-damaging agents.

Figure 1.

Figure 1

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Bepristat-2a is synergistic with Top-II and not Top-I inhibitors in various cancer cell lines. (A) Colony formation assay of bepristat-2a (bep2a) in combination with Top-II inhibitors (doxorubicin, etoposide, and mitoxantrone) in GB-1 cells. The cells were treated for 7 days before staining with crystal violet. The synergy of doxorubicin and bepristat-2a in GB-1 cells is illustrated with Combenefit software using HSA model. The intensity of each well in the colony formation assay was calculated using ImageJ. (B) Colony formation assay of bepristat-2a in combination with Top-I inhibitors (camptothecin and irinotecan) and other DNA-damaging agents (TMZ and oxaliplatin) in GB-1 cells. The cells were treated for 7 days before staining with crystal violet. (C) Dose–response curve for cell viability of OVCAR8 cells treated with etoposide alone or in combination with bepristat-2a for 72 h as measured by the MTT assay. Etoposide was combined with fixed concentration of bepristat-2a (10 μM). The error bars represent standard deviation of three biological replicates (D) Synergy of doxorubicin and bepristat-2a in OVCAR8 and (E) A549 and HCT116 cells is illustrated with Combenefit software using HSA model. The intensity of each well in the colony formation assay was calculated using ImageJ.

PDIA1 Inhibition Results in the Downregulation of UHRF1

Previously, we showed that inhibition of PDIA1 results in the downregulation of genes involved in DNA damage and repair pathways.7,8,17 We performed an in-depth analysis of DNA repair genes that were downregulated after chemical or genetic inhibition of PDIA1 using five published bromouridine sequencing (Bru-seq) datasets7,17 (Table S1). Treatment of U87 glioma cells with BAP2 or PACMA31 or knockdown of PDIA1 using siRNA-P4HB or shRNA-P4HB resulted in the downregulation of DNA repair genes (Figures 2A and S3A). We analyzed the Gene Set Enrichment Analysis (GSEA) gene set, GO_DNA_REPAIR, and focused on the core enrichment genes that were downregulated in all five Bru-seq datasets. We detected 44 common downregulated genes and focused on UHRF1 in this study (Figure 2B, Tables S1 and S2). UHRF1 is a multifunctional protein involved in the regulation of various cellular processes including DNA methylation, cell cycle, and DNA repair, and it is highly overexpressed in many cancers and impacts tumorigenesis.23 In addition, UHRF1 was originally identified as a regulator of Top-II-α (TOP2A) expression.24−26 In agreement with previous reports,23 using the Gene Expression Profiling Interactive Analysis (GEPIA) database, we observed that UHRF1 is upregulated in various tumors including GBM and is associated with poor survival in adenoid cystic carcinoma, mesothelioma, and pancreatic adenocarcinoma (Figures 2C and S3B,C). The expression of UHRF1 is regulated by various transcription factors including FOXM1, E2F1, and E2F8.27 Interestingly, we observed that these transcription factors were downregulated in all five Bru-seq datasets (data not shown), indicating that PDIA1 inhibition may have impacted UHRF1 expression through downregulation of its regulators.

Figure 2.

Figure 2

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Chemical or genetic inhibition of PDIA1 leads to the downregulation of UHRF1 in glioma cells. (A) GSEA plots showing that treatment of U87 cells with PACMA31 or knockdown of P4HB (PDIA1) in U87 cells negatively correlates with enrichment of DNA repair gene set. NES: normalized enrichment score. (B) Treatment with BAP2 or PACMA31 or knockdown of P4HB using siRNA (in U87) or doxycycline-inducible shRNA (in U87 or D54) negatively correlates with enrichment of GO_DNA_REPAIR including 44 common downregulated core enrichment genes among which is UHRF1. The Bru-seq datasets used in this analysis are listed in Table S1. (C) Box plot representing UHRF1 expression in GBM samples (red) versus matched normal data (black) (TCGA + GTEx) using GEPIA online tool. Asterisk (*) indicates p < 0.05. TPM: transcripts per million. (D) Bar graph (top) and representative Western blot (bottom) showing that treatment of U87 cells with BAP2 for 48 h at indicated concentrations results in downregulation of UHRF1. The band intensity of each sample was normalized to GAPDH or β-tubulin, which were used as loading controls, and each bar represents the log10 fold change (logFC) of UHRF1 relative to the control sample. The error bars represent standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. (E) Bar graph (top) and representative Western blot (bottom) showing that knockdown of P4HB in doxycycline-inducible shRNA-P4HB U87 cells and in GB-1 cells using two different siRNAs results in downregulation of UHRF1. Doxycycline (2 μg/mL) was added for 72 h on U87-shRNA cells. P4HB-siRNA1, P4HB-siRNA2, and scrambled RNA (scRNA) were added to GB-1 cells for 72 h. PDIA1 antibody was used to assess the knockdown of P4HB. The band intensity of each sample was normalized to GAPDH, which is used as a loading control, and each bar represents the logFC of UHRF1 relative to the control sample. The error bars represent the standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. (F) Bar graph (left) and representative Western blot (right) showing that treatment of DBTRG cells with bepristat-2a (bep2a) for 16 h at indicated concentrations results in dose-dependent downregulation of UHRF1. The band intensity of each sample was normalized to β-tubulin, which is used as a loading control, and each bar represents the logFC of UHRF1 relative to the control sample. The error bars represent the standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. (D–F) Uncut Western blots for three biological replicates are shown in the Supporting Information.

To validate the Bru-seq data, we treated U87 glioma cells with BAP2 for 48 h, and this treatment resulted in significant downregulation of UHRF1 protein level in a dose-dependent manner (Figure 2D). To confirm that PDIA1 inhibition results in the downregulation of UHRF1, we performed knockdown of PDIA1 using doxycycline-inducible shRNA-P4HB in U87 cells and siRNA-P4HB in GB-1 cells. Our results show that the knockdown of PDIA1 resulted in significant downregulation of UHRF1 protein in both cell lines (Figure 2E). We then asked whether PDIA1 inhibition through bepristat-2a also results in the downregulation of UHRF1. We treated another GBM cell line, DBTRG, with increasing concentrations of bepristat-2a which resulted in the downregulation of UHRF1 in a dose-dependent manner (Figure 2F). We also performed a time-course Western blot analysis of UHRF1 following bepristat-2a treatment in DBTRG cells. UHRF1 protein levels start to decrease as early as 6 h after bepristat-2a treatment and the most downregulation appears after 24 h (Figure S3D). RAD51, a DNA repair protein previously reported to be downregulated by PDIA1 inhibition,17 was also downregulated after bepristat-2a treatment (Figure S3D). These results suggest that chemical or genetic inhibition of PDIA1 in glioma cell lines leads to the downregulation of UHRF1 at both transcript and protein levels.

PDIA1 Inhibition Results in Proteasome-Mediated Degradation of UHRF1

To determine whether the downregulation of UHRF1 is proteosome-dependent, we treated a panel of glioma cell lines, GB-1, DBTRG, and U87, with BAP2 and bepristat-2a in the presence or absence of proteasome inhibitor, MG132. In the three cell lines, treatment with bepristat-2a resulted in the downregulation of UHRF1, which was rescued by the addition of MG132 (Figure 3A). Treatment with BAP2 for 24 h resulted in significant downregulation of UHRF1 only in DBTRG cells, and in GB-1 cells, the effect of bepristat-2a on UHRF1 level is minimal in comparison to the other cell lines. One possible explanation is the different mutation profiles of these glioma cell lines, which may impact the effect of PDIA1 inhibition on the degree of UHRF1 downregulation. In agreement with previous studies,17 RAD51 was also downregulated after treatment with the PDIA1 inhibitors in a proteasome-dependent manner. Bepristat-2a was more effective than BAP2 in downregulating UHRF1 in all three cell lines. To validate that PDIA1 inhibition results in the downregulation of UHRF1 in a proteosome-dependent manner, we performed knockdown of PDIA1 using doxycycline-inducible shRNA-P4HB in U87 cells and siRNA-P4HB in GB-1 cells in the presence or absence of MG132. Knockdown of PDIA1 in both cell lines resulted in the downregulation of UHRF1 protein levels which was partially rescued by MG132 treatment (Figure S4A,B). Collectively, these results suggest that in addition to downregulating UHRF1 gene expression, PDIA1 inhibition results in proteasome-mediated degradation of UHRF1 protein.

Figure 3.

Figure 3

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PDIA1 inhibition leads to proteasome-mediated downregulation of UHRF1 and RAD51 levels. (A) Bar graph (top) and representative Western blot (bottom) indicating downregulation of UHRF1 by BAP2 and bepristat-2a (bep2a) which is rescued by MG132 treatment. RAD51 was also blotted but not displayed in the bar graph. GB-1, DBTRG, and U87 cells were treated with BAP2 (10 μM), bepristat-2a (30 μM), and MG132 (5 μM) for 24 h. The band intensity of each sample was normalized to β-tubulin, which is used as a loading control, and each bar represents the logFC of UHRF1 relative to the control sample. The error bars represent standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. Uncut Western blots for three biological replicates are shown in the Supporting Information. (B) Bar graph showing the log2 fold change of UHRF1 in cells treated with the indicated ER-stress-inducing agents in comparison to control (p-value < 0.05 for all). The cell line corresponding to each dataset is shown on top of the bar. The six datasets used for this analysis are listed in Table S3. (C) Western blot analysis of GB-1 and DBTRG cells treated with tunicamycin (5 μg/mL) with or without MG132 (5 μM) for 24 h. UHRF1 and RAD51 levels are repressed by tunicamycin and rescued by MG132 treatment. The band intensity of each sample was normalized to β-tubulin, and the numbers above each band represent the fold change of UHRF1 and RAD51 relative to the control sample.

To determine whether ER stress affects UHRF1 protein level, we analyzed published gene expression datasets of various cells treated with ER-stress-inducing agents such as tunicamycin, thapsigargin, and dithiothreitol (DTT) (Table S3). We observed that the expression of UHRF1 is significantly repressed in all of the six datasets that we have analyzed (Figure 3B). To determine if ER stress impacts UHRF1 protein level, we treated GB-1 or DBTRG cells with tunicamycin for 24 h. Treatment with tunicamycin leads to the downregulation of UHRF1 and RAD51 protein levels which is rescued with the addition of MG132 (Figure 3C). These results suggest that PDIA1 inhibition or ER stress induction results in proteasome-mediated degradation of UHRF1.

Previous studies showed that UHRF1 is essential for RAD51 loading to DNA damage sites and is important for DNA repair;28,29 however, it is not known whether UHRF1 impacts RAD51 expression. To answer this question, we knocked down UHRF1 in GB-1 and DBTRG cells using two different sets of siRNAs and measured RAD51 protein levels. Strikingly, the knockdown of UHRF1 for 72 h results in the downregulation of RAD51 protein in both cell lines (Figure S5A,B). Similar results were obtained using OVCAR8 cells (Figure S5B). In contrast, the knockdown of RAD51 in DBTRG cells did not significantly impact the level of UHRF1 (Figure S5C) indicating that UHRF1 is an upstream regulator of RAD51, but RAD51 levels do not significantly impact UHRF1 expression. In addition, we found a significant positive correlation between UHRF1 and RAD51 gene expression levels in GBM and other tumors (Figure S5D,E). These results suggest that PDIA1 inhibition causes downregulation of UHRF1 which in turn impacts RAD51 protein level.

Bepristat-2a Increases the Level of SLC7A11 and Is Protective against Ferroptosis

Previously, it was shown that PACMA31, an irreversible PDI inhibitor, induces ferroptosis and is synergistic with cystine-glutamate antiporter xCT (SLC7A11) inhibitors such as erastin and sorafenib.30,31 To determine if bepristat-2a induces cell death in a similar mechanism, we performed combination studies with ferroptosis inhibitors, ferrostatin-1 and liproxtatin-1, and ferroptosis activators including (1S,3R)-RSL3, erastin, FINO2, and FIN56. For these combination studies, we pretreated cells with bepristat-2a or PACMA31 for 24 h before adding the ferroptosis compounds. Strikingly, unlike PACMA31, bepristat-2a showed significant antagonism with all of the tested ferroptosis activators in the clonogenic cell survival assay (Figures 4A,B and S6A). Bepristat-2a did not show antagonism with ferrostatin-1 or liproxtatin-1 in contrast to PACMA31 which shows strong antagonism with ferrostatin-1 (Figure S6A–C). To determine the mechanism of ferroptosis inhibition, we treated GB-1 cells with bepristat-2a and measured the expression of proteins involved in ferroptosis regulation such as SLC7A11 (system xCT), activating transcription factor 4 (ATF4) and glutathione peroxidase 4 (GPX4). SLC7A11 and GPX4 inhibit ferroptosis by preventing the accumulation of lipid hydroperoxides,32 while ATF4 prevents ferroptosis through upregulating SLC7A11.33 Indeed, we found a significant increase in the levels of SLC7A11 and ATF4 after treatment of GBM cells with 1× or 2× IC50 cytotoxic concentration of bepristat-2a (as measured via MTT assay), but no significant impact on GPX4 level (Figure 4C,D). Interestingly, treatment of DBTRG cells with higher cytotoxic concentrations of PACMA31 did not induce a similar level of SLC7A11 (Figure S6D). In addition, we treated GB-1 cells with 2× IC50 cytotoxic concentration of bepristat-2a, PACMA31, or BAP2 (IC50 was measured via MTT assay) and measured the levels of ATF4 protein. Strikingly, only bepristat-2a significantly increased the level of ATF4 at this cytotoxic concentration indicating that bepristat-2a is a stronger inducer of ER stress and the UPR (Figure S6E). System xCT is important for cellular glutathione biosynthesis. Interestingly, we observed that bepristat-2a was antagonistic with buthionine sulphoximine (BSO), an inhibitor of glutathione synthesis, in both GB-1 and DBTRG cells as measured by clonogenic cell survival assay (Figures 4E and S6F). Collectively, these results suggest that treatment with bepristat-2a protects cells against ferroptosis through upregulating ATF4 and system xCT.

Figure 4.

Figure 4

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Bepristat-2a increases the level of SLC7A11 and is protective against ferroptosis. (A) Colony formation assay of bepristat-2a (bep2a) and ferroptosis inducers ((1S,3R)-RSL3 and erastin) in combination in GB-1 cells. GB-1 cells were pretreated with bepristat-2a for 24 h. The cells were treated for 7 days before staining with crystal violet. The antagonism between bepristat-2a and (1S,3R)-RSL3 and erastin is illustrated with Combenefit software using HSA model. The intensity of each well in the colony formation assay was calculated using ImageJ. (B) Colony formation assay of bepristat-2a and ferroptosis inducers (FINO2 and FIN56) in combination in GB-1 cells. GB-1 cells were pretreated with bepristat-2a for 24 h. The cells were treated for 7 days before staining with crystal violet. (C) Bar graph (left) and representative Western blot (right) showing induction of SLC7A11 and ATF4 after treatment of GB-1 cells with bepristat-2a (25 and 50 μM) for 6 h. The band intensity of each sample was normalized to β-tubulin or GAPDH, and each bar represents the logFC of SLC7A11, ATF4, and GPX4 relative to the control sample. The error bars represent the standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. Uncut Western blot images for three biological replicates are shown in the Supporting Information. (D) Western blot analysis showing induction of SLC7A11 after treatment of GB-1 and DBTRG cells with bepristat-2a for 24 h. The band intensity of each sample was normalized to β-tubulin, and the numbers above each band represent the fold change of SLC7A11 relative to the control sample. (E) Colony formation assay of bepristat-2a and buthionine sulfoximine (BSO) in combination in GB-1. The cells were pretreated with bepristat-2a for 24 h. The cells were treated for 7 days before staining with crystal violet.

Bepristat-2a Increases Apoptosis Induced by Doxorubicin

To determine the cell death mechanism of bepristat-2a, we treated GB-1 cells with a combination of bepristat-2a and apoptosis inhibitors, NS3694 and Z-VAD-FMK, autophagy inhibitor (chloroquine) and necroptosis inhibitor (necrostatin-1). The combination of bepristat-2a and apoptosis inhibitors resulted in increased clonogenic survival of GB-1 cells indicating that apoptosis inhibitors partially rescued cell death induced by bepristat-2a (Figure 5A). In contrast, chloroquine and necrostatin-1 did not rescue cell death by bepristat-2a (Figure S7A–D). Doxorubicin induces tumor cell death through apoptosis;34,35 therefore, we hypothesized that the combination of bepristat-2a increases apoptosis induced by doxorubicin. We treated GB-1 cells with bepristat-2a, doxorubicin, or a combination of both and performed a Western blot analysis of apoptosis markers. We observed that treatment of GB-1 and DBTRG cells with the combination of bepristat-2a and doxorubicin leads to more prominent repression of UHRF1 and RAD51 and more prominent induction of apoptosis markers, cleaved caspase 3 and 9, in comparison to each treatment alone (Figures 5B,C and S7E). To validate that PDIA1 inhibition increases apoptosis induced by doxorubicin, we treated GB-1 cells with doxorubicin in combination with siRNA-P4HB and analyzed apoptosis markers. Indeed, we observed increased levels of apoptosis markers when cells were treated with both doxorubicin and siRNA-P4HB in comparison to each treatment alone (Figure S7F). To determine whether the combination of bepristat-2a and doxorubicin induces apoptosis through increasing DNA damage, we measured the level of γH2AX, which is a marker of double-strand breaks (DSBs). The combination of subtoxic concentration of bepristat-2a with doxorubicin increases the level of γH2AX in comparison to doxorubicin alone (Figure 5D). These results suggest that bepristat-2a sensitizes tumor cells to doxorubicin through the downregulation of DNA repair proteins and accumulation of DNA damage resulting in cell death through apoptosis.

Figure 5.

Figure 5

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Combination of bepristat-2a and doxorubicin decreases UHRF1 level and increases apoptosis. (A) Colony formation assay of bepristat-2a (bep2a) and apoptosis inhibitors (NS3694 and Z-VAD-FMK). Antagonism between bepristat-2a and apoptosis inhibitors in GB-1 cells is illustrated with Combenefit software using HSA model. GB-1 cells were pretreated with apoptosis inhibitors for 30 min. The cells were treated for 7 days before staining with crystal violet. The intensity of each well in the colony formation assay was calculated using ImageJ. (B) Bar graph (left) and representative Western blot (right) of UHRF1, RAD51, and apoptosis markers in GB-1 cells treated with bepristat-2a (20 μM), doxorubicin (2.5 μM), or combination of both for 24 h. The band intensity of each sample was normalized to β-tubulin or GAPDH. Only β-tubulin is shown in the Western blot (right) for simplicity (refer to the uncut blots in the Supporting Information). Each bar represents the logFC of UHRF1, RAD51, cleaved-caspase-9, and cleaved-caspase-3 relative to the control sample. C-PARP is only blotted once (n = 1) so it is not represented in the bar graph. The error bars represent the standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. (C) Bar graph representing Western blot analysis of UHRF1, RAD51, and cleaved-caspase-3 in DBTRG cells treated with bepristat-2a (20 μM), doxorubicin (2.5 μM), or combination of both for 24 h. The band intensity of each sample was normalized to β-tubulin, and each bar represents the logFC of UHRF1, RAD51, and cleaved-caspase-3 relative to the control sample. The error bars represent the standard deviation of three biological replicates. The asterisk (*) indicates p < 0.05. A representative blot is shown in Figure S7E, and uncut Western blots for three biological replicates are shown in the Supporting Information. (D) Western blot analysis of γH2AX in DBTRG and GB-1 cells treated with bepristat-2a (20 μM), doxorubicin (2.5 μM), or a combination of both for 24 h. The band intensity of each sample was normalized to β-tubulin, and the number above each band represents the fold change of γH2AX relative to the control sample.

Since treatment with PDIA1 inhibitors results in the downregulation of DNA repair genes, we hypothesized that bepristat-2a would be synergistic with DNA repair inhibitors. We performed combination studies in GB-1 and DBTRG cells using bepristat-2a and various DNA damage repair inhibitors. We observed that the combination of bepristat-2a with DNA-PK inhibitor, NU7441, resulted in reduced clonogenic survival of GB-1 and DBTRG cells compared to each treatment alone (Figure S8A,B). DNA-PK is an essential enzyme involved in the nonhomologous end joining (NHEJ), the major DSB repair pathway.36−38 Interestingly, bepristat-2a showed weak or no synergy with niraparib and olaparib, inhibitors of PARP1/2 which are involved in base excision repair (BER) and nucleotide excision repair (NER) pathways39 (Figure S8A,B). These results suggest that PDIA1 inhibition by bepristat-2a primarily impacts the NHEJ DNA repair pathway and most probably sensitizes tumor cells to Top-II inhibitors through this mechanism.

Discussion

The PDI family of proteins generally reside in the ER and are best known for their oxidoreductase and molecular chaperone functions.2 The role of the PDI family members in DNA repair mechanism has been realized only recently. Various PDI family members such as PDIA1, PDIA3, and PDIA9 have been shown to impact DNA repair pathways.7,8,17,40,41 PDIA1, the canonical PDI family member, is upregulated in various tumors including GBM and is correlated with tumor progression and patient survival; therefore, PDIA1 is considered a promising target for cancer treatment.2,4,42 In addition to its role in the UPR, our previous studies indicated that PDIA1 impacts DNA repair processes and affects the sensitivity of GBM cells to radiation and chemotherapy.7,8,17 In this study, we performed an in-depth analysis of our previously published Bru-seq datasets to determine DNA repair genes that are altered by inhibition of PDIA1. Strikingly, this analysis allowed us to discover, for the first time, that PDIA1 inhibition leads to the transcriptional repression as well as proteasome-mediated degradation of the oncogenic protein UHRF1 in GBM and other cancer cell lines.

UHRF1 is a multifunctional protein involved in various cellular processes including DNA methylation, cell cycle, transcriptional repression, and DNA repair.43 UHRF1 is considered a promising target for cancer therapy. Overexpression of UHRF1 promotes tumorigenesis in many cancers including lung, breast, liver, prostate, and others through altering DNA methylation and repressing tumor suppressor genes.23,44 In GBM, UHRF1 is significantly overexpressed in tumor tissues in comparison to normal tissues, and it was shown to be involved in silencing of the tumor suppressor gene p16INK4A affecting GBM cell proliferation.45,46 Therefore, inhibition of UHRF1 could be a potential new therapeutic approach in GBM.

In addition to its role in DNA methylation, UHRF1 has been shown to impact DNA repair through various mechanisms. UHRF1 plays a role in the recognition and repair of DNA interstrand crosslinks caused by various genotoxic agents.47,48 Additionally, it was reported that UHRF1 plays a role in the BER pathway.49 Importantly, UHRF1 impacts DSB repair pathways and affects the sensitivity of various tumor cells to irradiation and chemotherapeutic agents.50−57 Therefore, inhibition of UHRF1 in combination with available DNA-damaging antitumor agents is a promising approach for cancer therapy. It was shown that UHRF1 is important for RAD51 loading to DNA damage sites which is an important step in DSB repair.28,29 In this study, we observed that UHRF1 gene expression significantly correlates with RAD51 gene expression and showed that the knockdown of UHRF1 leads to the downregulation of RAD51 protein. In our previous studies, we indicated that PDIA1 inhibition leads to the downregulation of RAD51 protein in glioma cell lines.7,17 Thus, we hypothesize that PDIA1 inhibition impacts RAD51 levels through downregulation of UHRF1. Our findings emphasize the significance of PDI as a promising target for GBM cancer therapy not only through induction of ER stress but also through impacting the level of the oncogenic protein UHRF1.

Previously, our studies indicated that PDIA1 inhibition increases the sensitivity of GBM cells to radiation.8,17 In this study, we screened various chemotherapeutic agents in combination with a selective PDIA1 inhibitor, bepristat-2a, and identified a significant synergy between bepristat-2a and Top-II poisons in various cancer cell lines. We observed that PDIA1 inhibition increases the sensitivity of GBM and other cancer cell lines to Top-II inhibitors, doxorubicin, etoposide, and mitoxantrone. Strikingly, PDIA1 inhibition with bepristat-2a did not impact the sensitivity of tumor cells to Top-I poisons, TMZ or oxaliplatin. These results indicate that PDIA1 inhibition impacts DNA damage response pathways that are required to repair lesions induced predominantly by Top-II poisons such as DSBs. Similar to radiation, Top-II poisons induce replication-independent DSBs that are repaired by two pathways, homologous recombination (HR) or NHEJ, where NHEJ is the predominant repair pathway.38,58 In contrast, Top-I poisons result primarily in single-strand breaks (SSBs) that are converted to DSBs after DNA replication, so Top-I-induced DNA damage is primarily repaired through SSB repair pathways in addition to DSB repair pathways if DSBs were generated.38,59 On the other hand, alkylating and crosslinking agents such as TMZ and oxaliplatin are mainly repaired through SSB repair pathways such as direct repair, BER, mismatch repair (MMR), and NER.60,61 Interestingly, bepristat-2a was synergistic with NU7441, an inhibitor of DNA-PK which plays an essential role in the NHEJ pathway.36−38 Previous reports showed that DNA-PK inhibition increases sensitivity of tumor cells to Top-II inhibitors.62−64 In contrast, bepristat-2a was not synergistic with inhibitors of PARP enzymes which are best known for their function in SSB repair pathways such as BER and NER.38,39 Overall, our data support the hypothesis that PDIA1 inhibition impacts DSB repair pathways mainly the NHEJ arm leading to increased sensitivity to Top-II inhibitors.

Various PDIA1 inhibitors have been reported and studied in the context of cancer therapy.6−11,14,16,65 Some PDIA1 inhibitors bind in the PDIA1 active site such as PACMA31,6 35G89 and CCF642,11 whereas others bind in the substrate binding domain such as T8,10 BAP2,7 and bepristat-2a.14 PACMA31 and 35G8 induce cell death through ferroptosis,9,30,31 and it was reported that PACMA31 induces ferroptosis through direct binding and inhibition of GPX4.31 In stark contrast, we showed that bepristat-2a is protective against ferroptosis. In agreement with our data using bepristat-2a, a recent report showed that induction of ER stress in glioma cells is protective against ferroptosis through upregulation of PERK-ATF4-HSPA5 pathway.66 Another report showed that overexpression of ATF4 protects glioma cells from ferroptosis through upregulation of SLC7A11 (system xCT).33 In fact, inhibition of SLC7A11 results in diminished cystine uptake resulting in glutathione depletion and eventually ferroptosis.67,68 Consistent with these studies, we observed that treatment of GBM cells with bepristat-2a results in significant upregulation of ATF4 and SLC7A11 and inhibition of ferroptosis. At a comparable cytotoxic dose, bepristat-2a resulted in a stronger ER stress response in comparison to PACMA31 and BAP2 as measured by the amount ATF4 and SLC7A11 proteins suggesting that bepristat-2a may be protective against ferroptosis through a strong induction of UPR. In addition, a more recent study indicated that PDIA1 knockdown in breast cancer cells inhibited erastin-induced ferroptosis69 which emphasizes our findings that PDIA1 inhibition does not induce ferroptosis but in fact is protective against ferroptosis. On the other hand, we observed that bepristat-2a sensitizes tumor cells to Top-II inhibitors through increasing apoptosis. In agreement with our results, a previously reported PDIA1 inhibitor (T8) sensitized tumor cells to etoposide through increasing apoptosis.10 Overall our data favor the hypothesis that PDIA1 inhibition by bepristat-2a results in ER stress and activation of the PERK-ATF4 pathway which is protective against ferroptosis on the one hand, but induces apoptosis on the other hand.70

Despite advances in GBM research, overall patient survival remains poor, thus identifying new therapeutic approaches is essential to improve prognosis. Our study highlights the significance of PDIA1 as a promising therapeutic target in GBM. PDIA1 inhibition protects cells from ferroptosis, but on the other hand, leads to downregulation of UHRF1 which in turn negatively impacts DNA repair pathways leading to increased sensitivity of tumor cells to Top-II inhibitors and eventually cell death through apoptosis (Figure 6). Although Top-II inhibitors are not currently approved for GBM treatment, there are a number of preclinical and early clinical studies showing the potential of using Top-II inhibitors for the treatment of GBM through novel delivery methods.71−75 Therefore, combination of a PDIA1 inhibitor with Top-II inhibitors is a promising therapeutic approach for GBM treatment. However, developing an optimal CNS permeable small-molecule PDIA1 inhibitor that can be used clinically is urgently needed.

Figure 6.

Figure 6

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PDIA1 inhibition results in increased sensitivity of tumor cells to Top-II inhibitors through degradation of UHRF1. Treatment of tumor cells with PDIA1 inhibitors results in ER stress leading to transcriptional repression of UHRF1 gene and proteasome-mediated degradation of UHRF1 protein. PDIA1 inhibition results in resistance to ferroptosis, and downregulation of UHRF1 impacts DNA repair pathways resulting in the accumulation of DNA damage induced by Top-II inhibitors ultimately leading to apoptosis.

Experimental Section

Bioinformatics Analysis

Published Bru-seq datasets were analyzed for DNA repair pathways using GSEA of pre-ranked gene lists. The Bru-seq datasets included three datasets for U87 cells treated with BAP2, PACMA31, siRNA-P4HB,7 and two datasets for doxycycline-inducible shRNA-P4HB stable cell lines (U87 and D54).17 Core enrichment genes within DNA repair pathway (GO_DNA_REPAIR) were compared between the Bru-seq datasets. Published or available RNA-seq and microarray datasets for cells treated with ER-stress-inducing agents were analyzed for UHRF1.76−78 A list of these datasets is shown in Table S3. The datasets were either downloaded from the study or from gene expression omnibus (GEO, NCBI). GEPIA2 online tool (http://gepia2.cancer-pku.cn/#index) was used to determine UHRF1 expression in tumor samples and paired normal tissues. GEPIA2 was used to generate Kaplan–Meier survival plots for tumor samples with high expression versus low expression of UHRF1. Correlation analysis of RAD51 and UHRF1 was done using cBioPortal (https://www.cbioportal.org/).

Cell Lines and Cell Culture

Glioma cancer cell lines (GB-1, DBTRG, U87) were purchased from ATCC. GB-1 and DBTRG cell lines were maintained in DMEM 11965-092 (Gibco) supplemented with 10% FBS (Gibco), 1% Pen/strep (Gibco), and 1% l-glutamine (Gibco). U87 cells were maintained in DMEM 11995-065 (Gibco) supplemented with 10% FBS. Ovarian cancer cell line (OVCAR8) and lung adenocarcinoma cell line (A549) were purchased from the NCI Developmental Therapeutics Program and were maintained in RPMI 11875-093 (Gibco) supplemented with 10% FBS (Gibco). Colon carcinoma cell line (HCT116) was maintained in McCoy’s 5A 16600-082 (Gibco) supplemented with 10% FBS (Gibco). HFF-1 cells were maintained in DMEM 11995-065 (Gibco) supplemented with 10% FBS. All cells were in culture under 30 passages and maintained at 37 °C in a humidified atmosphere of 5% CO2. All cells were routinely tested for Mycoplasma contamination using the PlasmoTest (InvivoGen).

Western Blot

Cells were seeded in six-well plates for 24 h at 37 °C and then treated with various compounds. After treatment, the cells were washed with DPBS (Gibco) and harvested with lysis buffer (25 mM tris(hydroxymethyl)aminomethane, 150 mM NaCl, 17 mM Triton X-100, 3.5 mM SDS, pH 7.4) containing 1× protease/phosphatase inhibitor (Thermo Scientific). Cell lysates were sonicated, and the supernatant was collected after centrifugation at 12,000 rpm for 15 min at 4 °C. The protein concentration was determined with the BCA assay (Thermo Scientific). Protein samples were prepared (35 μg) and loaded onto 10 or 15% acrylamide (Bio-Rad) gels and electro-transferred into PVDF membranes (Trans-Blot Turbo RTA Mini PVDF Transfer Kit, Bio-Rad). Membranes were blocked in blocking buffer (StartingBlock, Thermo Scientific) for 1 h at room temperature, and incubated with primary antibodies overnight at 4 °C. Primary antibodies: UHRF1 (Rabbit pAb, ABclonal), PDIA1 (Rabbit mAb, Cell signaling), RAD51 (Rabbit mAb, Cell signaling), GAPDH (Rabbit mAb, Cell signaling), β-tubulin (Rabbit mAb, Cell signaling), cleaved caspase-3 (Rabbit mAb, Cell signaling), cleaved PARP (Rabbit mAB, Cell signaling), cleaved caspase-9 (Rabbit mAB, Cell signaling), SLC7A11 (Rabbit mAb, Cell signaling), ATF4 (Rabbit mAb, Cell signaling), GPX4 (Rabbit mAb, Abcam), γH2AX (Rabbit mAb, Cell signaling). All of the primary antibodies were diluted 1:1000. The membranes were then washed and probed with anti-rabbit secondary antibody (Cell Signaling, 1:7500) for 1 h at room temperature and membranes were imaged using Odyssey imaging system (LI-COR Biosciences). Band intensities were quantified using Image Studio Lite v5.2.

siRNA and shRNA Knockdown

Predesigned UHRF1-siRNA-1 and siRNA-2 were purchased from Integrated DNA Technologies (IDT) (hs.Ri.UHRF1.13.1 and hs.Ri.UHRF1.13.3). The cells (3 × 105) were seeded in a six-well plate overnight. Transfection of cells was performed using Lipofectamine RNAiMAX (Invitrogen) according to manufacturer’s instructions. The siRNA–lipofectamine complex was formed by combining 30 pmol of UHRF1-siRNA or scrambled RNA in 150 μL of Opti-MEM medium (gibco) with 9 μL of lipofectamine RNAiMAX reagent diluted with 150 μL of Opti-MEM medium. The siRNA–lipofectamine mixture was incubated for 5 min at room temperature and 250 μL was added to cells for a final siRNA amount of 25 pmol/well. The cells were incubated with the siRNA for 72 h and then were harvested for Western blot analysis. The same protocol was used for RAD51 siRNA (RAD51 siRNAs purchased from Thermo fisher s531928 and s531930), and PDIA1 siRNAs (purchased from IDT: hs.Ri.P4HB.13.1 and hs.Ri.P4HB.13.2). For doxycycline-inducible P4HB-shRNA-U87 cells, doxycycline (2 μg/mL; Sigma-Aldrich) was added to cells to induce P4HB knockdown for 72 h, and then the cells were harvested for Western blot analysis.

MTT Assay

Cell viability was assessed by the 3-(4,5-dimethylthiazol-2-yl)-2,5- diphenyltetrazolium bromide (MTT) colorimetric assay. OVCAR8 cells were seeded in 96-well plates at 2000 cells/well. After overnight incubation, the cells were treated with etoposide (Cayman) or etoposide + bepristat-2a for 3 days. MTT (VWR) solution (0.3 mg/mL) was added to each well for 3 h at 37 °C. Afterward, the media was aspirated and dimethyl sulfoxide (DMSO, 100 μL) was added to each well and the plates were shaken for 10–15 min at room temperature. The absorbance of formazan crystals was measured at 570 nm using a microplate reader (Molecular Devices). Cell viability rate was calculated as [(At – Ab)/(Ac – Ab)] × 100, where At, Ac, and Ab represent the absorbance values from the cells treated with the compound, the cells not treated with the compound, and blank, respectively. Dose–response curve was generated using GraphPad Prism. The same protocol was used for the MTT assay in GB-1 cells after the combination of doxorubicin or etoposide with siRNA-P4HB. The knockdown was performed using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer’s instructions. The same protocol was used for the MTT assay in HFF-1 cells. HFF-1 cells were seeded in 96-well plates at 5000 cells/well.

Colony Formation Assay

Cells were seeded in 96-well culture plates at 150–300 cells/well and allowed to attach overnight. The cells were treated with compounds in combination with bepristat-2a for 7 days. Compounds used for combination studies are doxorubicin (Cayman), etoposide (Cayman), mitoxantrone (Cayman), camptothecin (Cayman), irinotecan (Cayman), TMZ (Sigma-Aldrich), oxaliplatin (TSZ chem), ferrostatin-1 (Cayman), liproxtatin-1 (MedChemExpress), erastin (MedChemExpress), (1S,3R)-RSL3 (Cayman), FINO2 (Cayman), FIN56 (Cayman), NS3694 (Cayman), Z-VAD-FMK (Cayman), NU7441 (Cayman), chloroquine (LKT labs), necrostatin-1 (Selleckchem), niraparib (Cayman), and olaparib (MedChemExpress). After removing the medium, the cells were stained with 0.05% crystal violet solution for 20 min and then washed with ddH2O to remove excess stain. Plates were left overnight to dry and were imaged using iBrightFL1000 (Invitrogen). The intensity of each well was calculated using ImageJ, and synergy or antagonism was illustrated using Combenefit software using HSA model.

Acknowledgments

This work was funded by a grant from the NIH (CA193690).

Glossary

Abbreviations

GBM

glioblastoma

Top-II

topoisomerase II

TMZ

temozolomide

UPR

unfolded protein response

ER

endoplasmic reticulum

PDI

protein disulfide isomerase

Bep2a

bepristat-2a

Top-I

topoisomerase-I

Bru-seq

bromouridine sequencing

GSEA

Gene Set Enrichment Analysis

GEPIA

Gene Expression Profiling Interactive Analysis

DTT

dithiothreitol

ATF4

activating transcription factor 4

GPX4

glutathione peroxidase 4

BSO

buthionine sulfoximine

DSB

double-strand break

NHEJ

nonhomologous end joining

BER

base excision repair

NER

nucleotide excision repair

HR

homologous recombination

SSB

single-strand break

MMR

mismatch repair

MTT

3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide

TCGA

The Cancer Genome Atlas

GTEx

Genotype-Tissue Expression

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsptsci.2c00186.

  • PDIA1 inhibition and PDIA1 knockdown increases the sensitivity of GBM cells to topoisomerase II inhibitors; bepristat-2a is synergistic with topoisomerase II and not topoisomerase I inhibitors; UHRF1 is overexpressed in tumors and is correlated with lower survival; downregulation of UHRF1 after knockdown of PDIA1 is partially rescued by MG132 treatment; PDIA1 inhibition leads to proteasome-mediated downregulation of UHRF1, which is correlated with RAD51 levels; bepristat-2a and PACMA31 have the opposite effect on ferroptosis; bepristat-2a is not antagonistic with autophagy and necroptosis inhibitors; bepristat-2a is synergistic with DNA-PK inhibitor and not PARP inhibitors; UHRF1 gene expression change in five Bru-seq datasets; common downregulated DNA repair genes in five Bru-seq datasets; six gene expression datasets of cells treated with ER-stress-inducing agents; and Uncut Western blots (PDF)

Author Contributions

Concept, development, and methodology: R.M and N.N.; data acquisition and analysis: R.M.; writing manuscript: R.M.; revision of the manuscript: N.N.; study supervision and funding: N.N.

The authors declare no competing financial interest.

Supplementary Material

pt2c00186_si_001.pdf (2.9MB, pdf)

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