Inhibition of Autophagy Reveals ATR Protein Kinase as a Key Mediator of Cisplatin Sensitivity in Osteosarcoma
Janice S. Pereira, Gabriel Rosa, Aine Pears, Beata Burczynska, Britta Stordal, Scott J. Roberts, Helen C. Roberts
TL;DR
This study shows that blocking autophagy and ATR protein improves cisplatin effectiveness in osteosarcoma, potentially leading to better treatment outcomes.
Contribution
The study identifies ATR as a novel upstream regulator linking DNA damage, autophagy, and chemoresistance in osteosarcoma.
Findings
Higher autophagy levels correlate with worse outcomes in osteosarcoma tumors.
ATG7 knockout enhances cisplatin sensitivity in HOS-143B cells by reducing p53 phosphorylation.
ATR inhibition simultaneously disrupts DDR signaling and autophagy, increasing cell death in cisplatin-treated cells.
Abstract
Osteosarcoma (OS) is the most common primary malignant bone tumor. Although the introduction of chemotherapy has improved the survival rate of OS patients, chemoresistance remains a major clinical problem underlying poor survival outcome. This study investigated the role of autophagy in OS chemoresistance and identified ATR as a novel upstream regulator linking DNA damage signaling, autophagy, and chemoresistance. Elevated levels of autophagy were found in advanced grade and stage OS tumors, and higher autophagy levels were shown to be associated with poorer OS disease outcome. Chemotherapy significantly increased autophagy levels in HOS‐143B cells, while autophagy inhibition by autophagy‐related gene 7 knockout (ATG7 −/− ) significantly enhanced cisplatin (CIS) sensitivity in HOS‐143B cells. A kinase screen revealed a reduction in the phosphorylation of p53 (S15) in ATG7 −/−…
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FIGURE 7| Inhibitor | Fold change |
|
|---|---|---|
| VE‐821/ATRi A | −1.44 | 0.050 |
| BAY1895344/ATRi B | −4.89 | 0.008 |
| AZD6738/ATRi C | −3.72 | 0.003 |
| AZ20/ATRi D | −4.58 | 0.004 |
- —Bone Cancer Research Trust10.13039/100011719
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Taxonomy
TopicsAutophagy in Disease and Therapy · Cell death mechanisms and regulation · Microtubule and mitosis dynamics
Introduction
1
Osteosaroma (OS) is a primary malignant tumor of the bone with a worldwide incidence of 3.4 cases per million per year [1, 2]. It can affect any bone but shows a predilection for bones with rapid growth rate such as the distal femur (43%), the proximal tibia (23%), and the proximal humerus (10%) [3]. The age distribution of OS is bimodal, with the first peak at 10–20 years of age and a second peak at 60–70 years of age. The former peak suggests a relationship with the expanding growth plates during puberty and the latter peak represents a consequence of Paget's Disease of Bone, dedifferentiated chondrosarcomas and/or prior radiation therapy [4]. Before 1970, surgical resection was the primary treatment for OS. However, the advent of dose‐intensive chemotherapy led to a remarkable improvement in prognosis for patients with localized OS (80% of cases) [2]. Today, multi‐agent dose‐intensive chemotherapy, mainly consisting of cisplatin (CIS), doxorubicin hydrochloride (DOX) and methotrexate is the gold standard treatment for OS [5]. Neoadjuvant chemotherapy is initiated 10 weeks before surgery (limb‐salvage surgery or amputation) to shrink large tumors. Adjuvant chemotherapy is administered for one year following radical resection to destroy the remaining tumor cells. This multimodal treatment approach has evolved to be the most efficacious way to increase the survival rate of OS patients and has significantly improved the 5‐year survival rate of OS patients without metastasis from 20% to almost 70% [2, 6]. However, some patients can resist chemotherapy leading to tumor relapse and metastasis [7].
OS exhibits both intrinsic as well as acquired resistance to chemotherapy. Due to intrinsic resistance, OS is often treated with a higher dose of chemotherapy than other cancers. Despite this, it quickly develops acquired resistance, making OS one of the most resistant cancers to chemotherapy [8, 9]. Accumulated evidence suggests that a cell survival process called autophagy is one of the main mechanisms underlying chemoresistance in several cancers, including OS [10, 11, 12, 13]. Studies have reported that autophagy is activated in response to DNA damage‐inducing chemotherapy to restore cellular homeostasis and evade cellular death [14]. In this process, damaged cellular organelles, cytoplasmic components as well as misfolded proteins are sequestered by autophagosomes which later fuse with lysosomes for lysosomal degradation. Subsequently, degradation products such as nucleotides, amino acids, fatty acids, sugars and ATP are transported to the cytosol where they are reused by the cell to improve cellular viability, giving rise to the chemoresistant phenotype [9, 15].
The autophagy pathway is regulated by various autophagy‐related proteins and studies have been focusing on targeting these protein levels to investigate the prospects for improving survival outcome of cancer patients. A study by Desai et al. [16] showed that the transcriptional expression of ATG7 is controlled by heat shock factor 1 (HSF1) and has an inverse correlation with the chemotherapeutic prognosis of breast cancer patients. Furthermore, knockdown of HSF1 reduced autophagy induction via ATG7 inhibition in breast cancer cells treated with chemotherapy and enhanced the breast cancer cells' sensitivity to carboplatin treatment. In esophageal squamous cell carcinoma, ATG7 interference was shown to increase sensitivity to cisplatin treatment by enhancing apoptotic cell death. These and several other studies suggest that autophagy inhibition via suppression of ATG7 levels alongside standard of care treatment could enhance the anti‐cancer effects and thereby improve patient survival outcome [17, 18].
Emerging studies in other tumor types have implicated upstream kinases and stress response pathways such as the Ataxia Telangiectasia and Rad3‐related protein (ATR) in modulating autophagy through phosphorylation of autophagy‐related proteins such as BECLIN1, LC3, ATG31, ATG14L and ATG7 [19, 20, 21]. ATR is primarily known for its role in the DNA damage response (DDR) in response to replication stress, which is often induced by chemotherapeutic agents. However, the role of ATR in regulating autophagy in OS and, more specifically, in influencing ATG7‐dependent autophagy and chemoresistance has not yet been elucidated.
The primary aim of this study was to further investigate the role of autophagy and ATR signaling in OS chemoresistance by knocking out ATG7 in highly aggressive/metastatic HOS‐143B cells. In doing so, the study aimed to uncover novel interactions between the DDR pathway and autophagy, potentially identifying ATR as a key regulator of chemoresistance in OS.
Our study reveals novel therapeutic approaches and proposes combinatory strategies involving inhibition of autophagy and targeting of the ATR pathway to overcome resistance and improve outcomes for OS patients.
Materials and Methods
2
LC3 Immunohistochemistry of OS Tissue Microarray and MAP1LC3B Gene Profile Analysis
2.1
A commercially available human OS tissue microarray (AMSBIO, OS804c) was probed for LC3 (A/B subtypes) expression to assess autophagy levels in OS biopsies. Sections were deparaffinized with xylene and rehydrated through graded ethanol. Antigens were retrieved with sodium citrate buffer (95°C–98°C) for 10 min and quenched with 3% hydrogen peroxide before blocking in 5% Normal Goat Serum. LC3 A/B (D11) XP Rabbit mAb (Cell Signaling Technology, 3868) primary antibody was incubated overnight at 4°C. The next day, after washing the slide, the secondary antibody (SignalStain Boost Detection Reagent, Cell Signaling Technology, 8114) was applied for 30 min at RT and stained with SignalStain DAB Substrate Kit (Cell Signaling Technology, 8059P). The nuclear counterstain was carried out using hematoxylin. Images were obtained using Eclipse 50i (Nikon, Tokyo, Japan) and stain intensity was quantified using ImageJ software. For this study, an area fraction method was used to calculate LC3 expression in the OS histological samples. This calculates the area percentage of DAB chromogen signal normalized to the overall area of the biopsy tissue (quantification was performed at 40× magnification). MAP1LC3B gene profile analysis from an integrative study of 88 mixed OS tumors (mesenchymal) (GSE42352) was obtained from R2 Genomics Analysis and Visualization platform (https://hgserver1.amc.nl/cgi‐bin/r2/main.cgi).
Cell Lines and Culture
2.2
HOS‐143B (ATCC, CRL‐8303, authenticated) cells were cultured in 1× DMEM + GlutaMAX (ThermoFisher Scientific) supplemented with 10% FBS (ThermoFisher Scientific), 1% non‐essential amino acids (ThermoFisher Scientific), 1% sodium pyruvate (ThermoFisher Scientific), and 1% penicillin–streptomycin (ThermoFisher Scientific). In addition, 0.2% 5‐bromo‐2′‐deoxyuridine was added for cell expansion (Sigma). The cells were grown in a humidified chamber with 5% CO_2_ at 37°C and were routinely checked for mycoplasma and were mycoplasma‐free.
Dose Response Assay
2.3
1 × 10^4^ cells/well in 100 μL of cell culture media/well were seeded in a 96 well plate and left to attach overnight. Cells were treated in triplicates with a two‐fold serial dilution of chemotherapy (Doxorubicin or Cisplatin, Sigma‐Aldrich) to reach a final volume of 200 μL/well. After 48 h of incubation, the cells were washed with PBS and incubated with 100 μL of fresh acid phosphatase substrate (Sigma‐Aldrich) in sodium acetate buffer (0.00263 g/mL) for 1.5 h. Subsequently, 50 μL of 1 M sodium hydroxide (Sigma‐Aldrich) was added. Absorbance was measured at 405 nm on a FLUOstar Omega microplate reader. Cell viability graphs (non‐linear regression‐curve fitting) and bar graphs for the IC_50_ were plotted using GraphPad Prism (GraphPad Software, CA, USA).
Western Blotting for Protein Analysis
2.4
20 × 10^4^ cells were grown for 24 h prior to drug treatment (48 h). Nitrocellulose membranes were incubated with primary antibody for 1 h (1:1000; Cell Signaling Technology, 12741, LC3 A/B; 5114, p62; 8558, ATG7; 12994, ATG5, p53 (7F5), 2527, Phospho‐p53 (S15), 9284) at 4°C. Binding of HRP‐linked secondary antibody (1:2000; Cell Signaling Technology, 7074) was visualized by enhanced chemiluminescence using Li‐Cor Odyssey Fc machine and quantified using Image Studio (Licor, Cambridge, UK). Normalization for total protein was performed by reprobing the membrane with mouse anti‐mouse β‐actin (1:1000, Mouse IgG; Abcam) for 1 h at RT, followed by goat anti‐mouse IgG HRP (1:2000; Biorad) for 1 h at RT.
Immunofluorescence for LC3 Punctate
2.5
1 × 10^4^ cells were seeded in 8‐well chamber slide (Sigma Aldrich) and left to attach overnight. Cells were treated the following day and incubated for an additional 48 h. Cells were fixed in ice‐cold 100% methanol and blocked in 5% Normal Goat Serum for 1 h at RT. LC3 A/B (D11) XP Rabbit mAb (Cell Signaling Technologies) was added and the cells incubated overnight at 4°C before being incubated with Alexa Fluor 488 Conjugate secondary antibody (Cell Signaling Technologies) and mounted in Antifade Mounting Medium with DAPI (Vector Labs). Images were acquired with Confocal Microscopy Leica DMI4000 B (Leica Microsystems).
RNP‐CRISPR/Cas9 KO of ATG7 in HOS‐143B Cells
2.6
For specific Knock out (KO) of the ATG7 gene, Single guide RNAs (sgRNAs) targeting Exon 11 were obtained from the predesigned TrueGuide Synthetic gRNA library (ThermoFisher Scientific, CRISPR1032818_SG); 5′‐TTCCAATAGCTGGGCAGCAA‐3′. TrueGuide sgRNA Negative Control (ThermoFisher Scientific) 5′‐AAAUGUGAGAUCAGAGUAAU‐3′ was used as a non‐targeting control (NTC). The ATG7 KO cell line was generated by reverse transfecting TrueGuide sgRNA and TrueCut Cas9 Protein v2 (ThermoFisher Scientific) using Lipofectamine CRISPRMAX Cas9 Transfection Reagent (ThermoFisher Scientific) as per manufacturer's protocol. The first step of the transfection involved the preparation of the Cas9 protein/gRNA/Cas9 Plus reagent solution in Opti‐MEM I Reduced Serum Medium, called the RNP complex. Then, 1.5 μL lipofectamine CRISPRMAX was diluted in 25 μL Opti‐MEM for 1 min, mixed with the RNP complex and incubated for 15 min at RT. Then, 50 μL of RNP‐Lipofectamine mix followed by 8 × 10^4^ cells/wells were added to each well of a 24‐well plate to reach a total volume of 200 μL/well. Cells were incubated at 37°C for 72 h and then harvested for cleavage detection and downstream analysis. GeneArt Genomic Cleavage Detection Kit (ThermoFisher Scientific) was used for cleavage detection, following the manufacture's protocol. Single cell clonal isolation was carried out by limiting dilution method. Cells were plated in a full 96‐well plate at a seeding density of 8 cells/mL. Clones were selected based on single colony growth and were later expanded into 6 well plates. After obtaining confluency of 50%–70%, protein lysates were retrieved from each clone and were probed for ATG7 expression by western blotting. Clones were selected based on no ATG7 expression and this was confirmed by qRT‐PCR (primers: ATG7 F: 5′GGCGGAGGCACCAAATGAT3′ R: 5′CCACATCCAAGGCACTGCTA3′; GAPDH: F: 5′AATCCCATCACCATCTTCCA3′ R: 5′TGGACTCCACGACGTACTCA3′).
Wound Healing Migration Assay
2.7
To determine the effect of ATG7 KO on migration activity in osteosarcoma, 2 × 10^5^ cells were seeded in a 6 well plate and allowed to attach overnight. Following treatment with DOX (0.75 μM) or CIS (5 μM) for 48 h, the confluent cell monolayer was scratched down the center of each well using a 200 μL tip. Subsequently, the monolayer was washed with PBS and fresh culture media was replaced in each well. The wounds were assessed under 10× objective at 0 and 24 h and photographed. The distance migrated was analyzed from 3 points for each sample [22].
Proteome Profiler Human Phospho‐Kinase Array
2.8
The effect of ATG7 KO on phospho‐kinase signaling pathways was investigated using the Proteome Profiler Human Phospho‐Kinase Array (R&D Systems) for simultaneous detection of phosphorylation levels in 43 protein kinases across multiple pathways. In order to isolate proteins, 5 × 10^5^ cells were seeded in 75 cm^2^ flasks and incubated for 72 h. After incubation, the detection of phosphorylation levels of kinases/related proteins was carried out following the manufacturer's instructions. Membranes were imaged on the Odyssey Fc Imaging system (Licor, Cambridge, UK).
Cell Death Analysis Using Flow Cytometry
2.9
Cell death analysis was performed using the Annexin V/PI double staining kit (ThermoFisher Scientific). The cells were trypsinised, washed twice with 1 × PBS, then resuspended in 1 × binding buffer. The cell suspension (1 × 10^5^ cells in 100 μL) was transferred into a FACS tube and stained with 5 μL of PI and 5 μL of Annexin V for 15 min at RT. Further, an extra 400 μL binding buffer was added to each tube prior to analysis using the BD FACSCalibur (BD Biosciences, NJ, USA) alongside CellQuest Pro software for analysis. A minimum of 10,000 cells were acquired for each sample.
Statistical Analysis
2.10
All data were expressed as means ± standard deviation (SD)/standard error of mean (SEM). Error bars in all figures indicate SD/SEM. Statistical significance for the differences between pairs of conditions was analyzed by a two‐sample t‐test. One‐way ANOVA was used to compare more than two conditions. The criterion for statistical significance for an experiment that was independently and successfully repeated three or more times was considered as p‐value less than or equal to 0.05.
Results
3
High Levels of Autophagy Relate to Reduced Survival in High‐Grade OS
3.1
Immunohistochemical staining of OS core biopsies for LC3 protein levels (A/B subtypes) revealed a clear pattern of autophagy levels corresponding to OS tumorigenesis and progression. The highest LC3 levels were observed in grade 3 OS tumors followed by grade 2 and normal cortical bone tissue, as shown in Figure 1A,B. By further grouping the results based on stage, LC3 was expressed at highest levels in the most advanced tumor subtypes followed by less aggressive subtypes and normal bone (Figure 1A,C). These OS biopsies reflected patients between the age of 13 to 64, with no observed difference in gender and tumor location. Samples categorized as grade 2 and grade 3 were T1N0M0 and T2N0M0 as per the TNM staging system, respectively, indicating no record of nearby lymph node dissemination and metastasis.
*LC3 correlates to OS grade and disease progression. (A) Representative images of immunohistochemical detection of LC3 A/B in primary OS tumor tissue of varying grades and stages and normal cortical bone tissue. (B) Quantification analysis of positive LC3 A/B area coverage (%) based on tumor grade (NB, n = 4; G2, n = 42; G3, n = 38). (C) Quantification analysis of positive LC3 A/B area coverage (%) based on tumor stage (NB, n = 4; IA, n = 12; IB, n = 32; IIA, n = 2; IIB, n = 32; IIIB, n = 2; IVB, n = 2). (D) Kaplan–Meier curve of MAP1LC3B gene expression and overall survival probability between OS patient group (GSE42352) with high (blue, n = 62) and low (red, n = 26) expressing MAP1LC3B tumors (p = 0.039). (E) Kaplan–Meier curve of MAP1LC3B gene expression and metastasis‐free survival probability between OS patient group (GSE42352) with high (blue, n = 62) and low (red, n = 26) expressing MAP1LC3B tumors (p = 0.0095). Data is presented as mean ± SEM. *p < 0.01. G2, grade 2; G3, grade 3; NB, normal cortical bone.
MAP1LC3B profile analysis from an integrative study of 88 mixed OS tumors (mesenchymal) (GSE42352) obtained from R2 Genomics Analysis and Visualization platform (https://hgserver1.amc.nl/cgi‐bin/r2/main.cgi) revealed a strong link between MAP1LC3B levels and OS patient prognosis. As shown in Figure 1D, the patient group with significantly higher MAP1LC3B gene expression levels (n = 62) was associated with a lower overall survival compared to the low MAP1LC3B expressing group (n = 26) (p = 0.039). Additionally, patients with high MAP1LC3B levels had less probability for metastasis‐free survival (p = 0.0095), shown in Figure 1E. Collectively, these results suggest a possible link between autophagy activation and OS disease outcome.
Chemotherapy Induces Autophagy In Vitro
3.2
A dose response assay was carried out to determine the IC_50_ for cisplatin or doxorubicin hydrochloride in HOS‐143B cells. For cisplatin the IC_50_ value was 5 μM (Figure 2A). For doxorubicin hydrochloride treatment, 0.75 μM of the drug was required to reduce the cell population of HOS‐143B cells to half (Figure 2B). HOS‐143B cells were treated with the respective IC_50_ dose of chemotherapy for western blotting analysis of autophagy markers. Figure 2C showed an increase in autophagic flux, characterized by the increased conversion of LC3‐I to LC3‐II. The LC3‐II:LC3‐I protein levels increased 9.13‐fold (p‐value: 0.039) with cisplatin treatment and 27.63‐fold (p‐value: 0.016) with doxorubicin hydrochloride treatment versus the untreated control (Figure 2D). The downregulation of p62 protein levels following chemotherapy confirmed autophagic clearance through the activation of autophagy. A 4.34‐fold (p‐value: 0.040) and 5.62‐fold (p‐value: 0.041) reduction in p62 protein levels was observed following cisplatin and doxorubicin hydrochloride treatment, respectively in comparison to the untreated HOS‐143B cells (Figure 2E). To further investigate the effects of chemotherapy treatment on autophagy induction, drug‐treated HOS‐143B cells were probed for LC3 puncta formation to detect changes in autophagic flux. Cells treated with doxorubicin or cisplatin displayed an increase in LC3 puncta formation, which is an index for the number of autophagosomes contained in the cytoplasm. Together, these results confirm that autophagy is activated with chemotherapy in HOS‐143B cells (Figure 2F).
*Activation of autophagy in chemotherapy‐treated HOS‐143B cells. (A) Dose response curve for cisplatin treatment. (B) Dose response curve for doxorubicin hydrochloride treatment. The HOS‐143B cells were dosed with either cisplatin or doxorubicin hydrochloride in increasing concentrations up to 200 or 20 μM, respectively. (C) Western blotting images of LC3‐I, LC3‐II, p62, and β‐actin protein levels in 5 μM cisplatin‐treated or 0.75 μM doxorubicin hydrochloride‐treated HOS‐143B cells. Untreated HOS‐143B cells served as the control. β‐actin is used as an internal control. Graphical representation of fold changes in protein levels (D) LC3‐II:LC3‐I and (E) p62 protein levels in cisplatin‐treated and doxorubicin hydrochloride‐treated HOS‐143B cells relative to the control. (F) FITC‐LC3 and DAPI immunofluorescence images of HOS‐143B cells treated with cisplatin (10 μM) or doxorubicin hydrochloride (1 μM) for 48 h. Data are expressed as mean ± SD. For all data n = 3 cell culture replicates run in triplicate; p ≤ 0.05.
ATG7 KO Disrupts Autophagy
3.3
To further define the fundamental role of autophagy in response to chemotherapeutic agents, ATG7 expression was stably knocked out (KO) by CRISPR/Cas9 and downstream responses quantified. For gene disruption, gRNA was targeted to ATG7 gene at Exon 11 of HOS‐143B cells (Figure 3Ai). The cleavage efficiency in the pooled HOS‐143B cell population following ATG7 KO was 36% as shown in Figure 3Aii. Single cell clonal isolation was performed to select specific clones with constitutive knockout of ATG7. Following clonal selection, ATG7 gene and protein levels from a single cell clonal isolate showed a significant reduction in ATG7 ^ −/− ^ compared to wild‐type (WT) HOS‐143B cells (14.8‐fold p < 0.001 and 34,429‐fold, p < 0.0001 respectively; Figure 3B–D). To further explore the effect of ATG7 KO on autophagy, levels of key biomarkers of autophagy (LC3‐II:LC3‐I and p62) were analyzed alongside ATG5. ATG7 KO inhibited autophagy in HOS‐143B cells as indicated by a significant decrease in LC3‐II:LC3‐I (2.9‐fold, p = 0.0006; Figure 3C,E) and an increase in p62 levels (2.7‐fold p < 0.0001; Figure 3B,F) in comparison to WT HOS‐143B cells. ATG5‐ATG12 complex formation was inhibited in ATG7 ^ −/− ^ HOS‐143B cells as indicated by the increase in protein levels of unconjugated ATG5 (63‐fold, p < 0.0001; Figure 3B,H) coupled with a decrease in conjugated ATG5‐ATG12 (913‐fold, p < 0.0001; Figure 3B,G) in ATG7 ^ −/− ^ HOS‐143B cells compared to WT HOS‐143B cells. Additionally, protein levels were also measured in HOS‐143B cells administered with Cas9 alongside a non‐targeting sgRNA sequence that was used as (NTC). The non‐targeting sgRNA does not recognize any sequence in the human genome. The insignificant changes observed between the WT and NTC HOS‐143B cells implies that observed effects were specific to the ATG7 sgRNA and not an off‐target effect caused by Cas9 delivery. This confirmed the role of the E1‐like enzymatic activity of ATG7 in the formation of the two conjugates, LC3‐II and ATG5‐ATG12 that is crucial in the autophagy process.
*KO of ATG7 by CRISPR/Cas9 ablates autophagy in HOS‐143B cells. (Ai) Schematic illustration of the binding site of TrueGuide sgRNA on Exon 11 of ATG7 gene. The sgRNA guides the Cas9 endonuclease to the target site for enzymatic double stranded cleavage (ThermoFisher.com). (Aii) Gel electrophoresis of PCR product for the cleavage assay shows a 36% reduction of ATG7 expression in the pooled cell population following ATG7 KO. (B) qRT‐PCR relative expression of ATG7 normalized to GAPDH in ATG7
−/− , NTC and WT HOS‐143B cells following single cell clonal isolation for constitutive knockout of ATG7. (C) Western blotting images of ATG7, LC3‐I, LC3‐II, p62, ATG5‐ATG12, ATG5 and β‐actin protein levels in ATG7
−/− , NTC and WT HOS‐143B cells following single cell clonal isolation for constitutive knockout of ATG7. Graphical representation of fold changes in relative protein levels of (D) ATG7, (E) LC3‐II:LC3‐I, (F) p62, (G) ATG5‐ATG12, (H) ATG5 in ATG7
−/− and NTC compared to the WT HOS‐143B cells. Data are expressed as mean ± SD. For all data n = 3 biological replicates run in triplicate; ****p < 0.0001; ***p < 0.001; *p < 0.01 (One‐way ANOVA).
Autophagy Inhibition Enhances Chemosensitivity and Reduces HOS‐143B Cell Migration
3.4
Levels of autophagy markers suggested that autophagy inhibition mediated by ATG7 knockout is sustained following chemotherapy in HOS‐143B cells characterized by the accumulation of p62 and decreased conversion of LC3‐I to LC3‐II (Figure 4A). Figure 4B–E demonstrates the dose response curves and graphical representation of the IC_50_ values of DOX or CIS for ATG7 ^ −/− ^, NTC and WT HOS‐143B cells. The IC_50_ values for CIS in ATG7 ^ −/− ^ HOS‐143B cells are 4.5 μM ± 0.21 and in WT HOS‐143B are 7.0 μM ± 0.69 (1.6‐fold decrease, p = 0.0036). These results confirmed that ATG7 knockout/autophagy inhibition significantly sensitized HOS‐143B cells to CIS treatment. The IC_50_ of DOX in ATG7 ^ −/− ^ and WT HOS‐143B cells was 0.7 and 0.6 μM respectively with no statistically significant fold change. Hence, ATG7 KO and subsequent autophagy inhibition had no effect on the sensitivity of DOX in HOS‐143B cells.
*Chemosensitivity and migration of ATG7 knockout HOS‐143B cells. (A) Western blotting images of LC3‐I, LC3‐II, p62 and β‐actin protein levels in ATG7
−/− , NTC and WT HOS‐143B cells treated with 5 μM of cisplatin or 0.75 μM of doxorubicin hydrochloride. Untreated cells served as the control. To assess chemosensitivity, ATG7
−/− , NTC and WT HOS‐143B cells were dosed with DOX and CIS in increasing concentrations up to 200 μM and 20 μM, respectively. (B) The dose response curve for DOX treatment. (C) Graphical representation of IC50 of DOX. (D) The dose response curve for CIS treatment. (E) Graphical representation of IC50 of CIS. To assess migration, a scratch assay was carried out and migratory distance was measured after 48 h of incubation. (F) Graphical representation of migratory distance (μm) of ATG7
−/− , NTC and WT HOS‐143B, in DOX or CIS‐treated and untreated conditions. Data are expressed as mean ± SD. For all data n = 3 biological replicates run in triplicate; **p < 0.01; p < 0.05 (One‐way ANOVA).
The wound healing migration assay revealed no difference in the migratory distance between ATG7 ^ −/− ^, NTC, and WT HOS‐143B cells under normal conditions. However, treatment with DOX and CIS displayed a 93.2% and 97.2% decrease in migratory distance in the ATG7 ^ −/− ^ compared to NTC HOS‐143B cells, respectively (p < 0.05; Figure 4F). These results imply that autophagy inhibition reduced the migratory potential of chemotherapy‐treated highly aggressive HOS‐143B cells.
Pharmacological Inhibition of ATR Enhanced CIS Sensitivity
3.5
The semi‐quantitative phosphorylation levels of 45 different kinases/related proteins associated with various signaling pathways were assessed between autophagy‐inhibited (ATG7 ^ −/− ^ HOS‐143B) and NTC HOS‐143B cells. The data as shown in Figure 5A,B revealed a decrease in phosphorylation of p53 at S15 in autophagy‐inhibited ATG7 ^ −/− ^ HOS‐143B cells compared to NTC HOS‐143B cells. This led to the identification of small molecule inhibitors of ATM and ATR kinases that are responsible for the phosphorylation of p53 at S15 called KU‐60019 and VE‐821 respectively. Figure 5C,D shows the dose response curve and IC_50_ values for DOX with/without of either 1 μM KU‐60019 or VE‐821. No significant differences were noted. However, a 2.00‐fold significant decrease (p‐value = 0.007) in IC_50_ for CIS was observed with 1 μM VE‐821 compared to the control (Figure 5E,F). This implies that a lower dose of CIS is required to reduce the population of HOS‐143B cells to half when combined with ATR inhibitor (ATRi) VE‐821. No significant changes in CIS sensitivity were observed with ATM inhibitor (ATMi) KU‐60019.
*Chemosensitivity of HOS‐143B cells following co‐administration with ATMi/ATRi. (A) Image showing the relative levels of phospho‐p53 (S15; boxed) in ATG7
−/− and NTC HOS‐143B cells. (B) Graphical representation of relative phosphorylation of p53 (S15). For the dose response assay, HOS‐143B cells were dosed with CIS or DOX in increasing concentrations up to 200 μM or 10 μM respectively alongside 1 μM of either ATMi/KU‐60019 or ATRi/VE‐821. (C) Dose response curve for DOX with/without 1 μM ATMi/ATRi. (D) Graphical representation of IC50 of DOX with/without 1 μM ATMi/ATRi. (E) Dose response curve for CIS with/without 1 μM ATMi/ATRi. (F) Graphical representation of IC50 values of CIS with/without 1 μM ATMi/ATRi. (G) Dose response curve of CIS combined with/without ATRi A–D. (H) Graphical representation of IC50 values of CIS with/without ATRi A–D. VE‐821/ATRi A, BAY1895344/ATRi B, AZD6738/ATRi C and AZ20/ATRi D. 1 μM of ATRi A, ATRi C and ATRi D and 0.1 μM of ATRi B. Data are expressed as mean ± SD. For all data n = 3 biological replicates run in triplicate; **p < 0.01; p < 0.05 (two‐sample t‐test vs. control).
To confirm that the increase in sensitivity to CIS following VE‐821/ATRi A (1 μM) treatment is a class effect, three other small molecule ATR inhibitors, namely, BAY1895344/ATRi B (0.1 μM), AZD6738/ATRi C (1 μM), and AZ20/ATRi D (1 μM), were assessed. The concentration of inhibitors was determined following an initial toxicity screen based on negligible effects on cellular growth (0.001–10 μM; 10‐fold dilutions). The IC_50_ values for ATRi A–D were 2.15 μM ± 0.45, 0.63 μM ± 0.03, 0.83 μM ± 0.14, and 0.68 μM ± 0.01, respectively (CIS IC_50_ = 3.10 μM ± 0.26). All investigated ATR inhibitors significantly increased sensitivity to CIS treatment, as shown in Figure 5G,H. The fold changes and p‐values are listed in Table 1. Interestingly, 0.01 μM BAY1895344/ATRi B was most efficient at enhancing growth inhibition in CIS‐treated HOS‐143B cells. Hence, ATRi B was used for further investigation.
Pharmacological Inhibition of ATR Enhanced Cellular Apoptosis and Blocked p53 Phosphorylation and Autophagy
3.6
Figure 6A–C demonstrates flow cytometry analysis used to determine the percentage of Annexin V‐positive and/or PI‐ positive in HOS‐143B cells treated with/without CIS and/or ATRi. CIS treatment induced cell death as indicated by a significant increase in the early/late apoptotic cell population; 5.51% in untreated control versus 67.27% in CIS‐treated HOS‐143B cells. The combination of ATR inhibitor and CIS‐treatment significantly increased the early/late apoptotic cell population further by 23.27% (p‐value: 0.003, CIS vs. CIS + ATRi). This suggests that ATR inhibition significantly enhanced CIS‐induced apoptosis in HOS‐143B cells. The effect of ATR inhibition on phospho‐53 (S15) and autophagy levels is shown in Figure 6D–J, respectively. Treatment significantly increased the phosphorylation of p53 in comparison to untreated HOS‐143B cells (16.86‐fold; p‐value: < 0.0001). However, treatment with 0.01 μM ATRi significantly reduced the protein levels of total p53 (463‐fold; p‐value: < 0.0001) as well as phospho‐p53 (13.32‐fold; p‐value: < 0.0001) compared to HOS‐143B cells treated with CIS only. This implies that treatment increased phosphorylation of p53 at S15 and ATR inhibition whilst blocking CIS‐treatment blocked the phosphorylation of p53 and reduced total p53 protein levels in HOS‐143B cells. The protein levels of ATG7 were significantly reduced following treatment with CIS and ATRi in comparison to CIS treatment alone (p‐value: < 0.0001). Thus, ATR inhibition depleted ATG7 protein levels in CIS‐treated HOS‐143B cells. Furthermore, a significant decrease in LC3‐I, LC3‐II and p62 was recorded with ATR inhibition in CIS‐treated HOS‐143B cells (p‐values: < 0.0001). Together, this suggests that the autophagy pathway is inhibited via ATR inhibition in CIS‐treated HOS‐143B cells.
*Pharmacological inhibition of ATR enhanced apoptosis and blocked p53 phosphorylation and autophagy. (A) Annexin V/PI staining of HOS‐143B cells with/without 5 μM CIS and/or 0.01 μM ATRi, BAY‐1895344. Quadrant guide: Lower left = live cells; Lower right, early apoptosis; Upper left, necrosis; Upper right, late apoptosis. (B) Percentage of the live, early and late apoptotic and necrotic cell population. (C) Statistical analysis of live, early and late apoptotic and necrotic cell population. (D) Western blotting images of total p53, phospho‐p53 and β‐actin protein levels in HOS‐143B cells treated with either 5 μM CIS or 0.01 μM ATRi, or a combination of both. Untreated cells served as the control. Graphical representation of fold changes in protein levels of (E) Total p53 and (F) phospho‐p53 relative to the control. (G) Western blotting images of ATG7, LC3‐I, LC3‐II, and β‐actin protein levels in HOS‐143B cells treated with either 5 μM CIS or 0.01 μM ATRi, or a combination of both. Untreated cells served as the control. Graphical representation of fold changes in protein levels of (H) ATG7 (I) LC3‐II:LC3‐I, and (J) p62 relative to the control. Data are expressed as mean ± SD. For all data n = 3 biological replicates run in triplicate; ****p < 0.0001; ***p < 0.001; **p < 0.01; p ≤ 0.05 (flow cytometry: One‐way ANOVA; Western blot: two‐sample t‐test vs. control).
Discussion
4
Autophagy is an evolutionarily conserved pathway that is used by cells to recycle damaged cellular components as a mechanism of adaptation to adverse environmental stimuli including cancer cell replication stress as well as exposure to chemotherapeutic agents. This study showed that higher autophagy levels are associated with lower metastasis‐free survival as well as overall survival in OS patients. Moreover, in vitro studies showed higher autophagy levels in OS cells treated with chemotherapy, and blocking autophagy through the KO of ATG7 increased cisplatin sensitivity and reduced OS cell migration. Further, p15 phosphorylation on S15 was shown to be reduced in the autophagy inhibited model of HOS‐143B OS cells, and blocking this phosphorylation activity via small molecule inhibitors mimicked the ATG7 KO effect in HOS‐143B cells, increasing sensitivity to cisplatin.
In the current study, it was evident through the staining of LC3 that higher autophagy levels were present in advanced OS tumors. Furthermore, high MAP1LC3B gene expression was associated with poorer survival outcomes and increased metastasis in OS patients. These results agree with a study carried out by White et al. [23], who have shown that higher levels of autophagy proteins were measured in high‐grade glioma in comparison to low‐grade tissues.
Several studies have shown that chemotherapy can upregulate autophagy in tumors as a means to protect transformed cells from organelle damage and nutrient deprivation [12, 24, 25]. In the current study, analysis of DOX ‐ CIS‐treated HOS‐143B cells suggested that autophagy is activated as a cellular coping mechanism following DNA damage induced by chemotherapy. Therefore, targeting autophagy could be a therapeutic strategy that may improve the efficacy of the standard of care anti‐cancer treatment. ATG7, being an E1 ligase, plays a crucial role in autophagy regulation and it has been reported that ATG7‐dependent autophagy modulates the progression of several types of cancer [26, 27, 28]. In this study, a significant increase in conversion of LC3‐I to lipidated LC3‐II and increased clearance of p62 was observed in CRISPR/Cas9 ATG7 ^ −/− ^ cells in comparison to WT and NTC HOS‐143B cells.
This data agrees with the observations made in ATG7 knockout mutants by Cui et al. [29], which showed high p62 protein levels and lower levels of LC3‐II proteins in the ATG7 knockout mutants compared to the controls (WT and empty vector control) suggesting that ATG7 knockout efficiently impeded autophagy. In addition to the role of the ATG7 enzymatic activity in the formation of the LC3‐I‐PE conjugate, it is also involved in the formation of the ATG5‐ATG12 conjugate. In the current study, the ATG5‐ATG12 conjugate was not formed in ATG7 ^ −/− ^ HOS‐143B cells and ATG5 was present in its free form, confirming ablation of the autophagy process.
To gain a mechanistic insight into the consequences of autophagy inhibition on chemosensitivity of OS, a chemotherapy dose response assay showed that CIS sensitivity increased in ATG7 ^ −/− ^ HOS‐143B cells, characterized by a reduction in the IC_50_ value. These findings agree with a study conducted by Piya et al. [30] who showed that suppression of ATG7 in acute myeloid leukemia enhanced the sensitivity to chemotherapeutic agents marked by an increase in DNA damage and cellular apoptosis. CIS induces DNA damage, and preventing the repair of damaged DNA through autophagy inhibition leads to increased cell death [31]. On the other hand, DOX induces cell death through multiple pathways, and autophagy inhibition does not affect these pathways significantly, as shown in the current study [32, 33].
Several different kinases as well as phosphatases are responsible for regulating autophagy through phosphorylation activity [34]. In the current study, variations in phosphorylation levels of p53 (S15) were revealed in ATG7 ^−/−^ HOS‐143B cells in comparison to NTC HOS‐143B cells. The phosphorylation of p53 at S15 serves as a critical trigger for the DDR in response to DNA damage inducing agents. ATM and ATR are two protein kinases involved in the phosphorylation of p53 at S15, recognizing double‐strand DNA breaks and replication stress, respectively [35, 36].
Currently, pharmacological inhibitors of ATM and ATR are in preclinical and clinical trials. The data obtained from preclinical studies have shown promising results for their use in combination with standard of care treatments, paving the way for clinical testing [37]. This study utilized two small molecule inhibitors for specific inhibition of ATM and ATR kinase activity. The dose response analysis revealed a statistically significant increase in sensitivity to CIS in HOS‐143B cells treated with ATR but not ATM inhibitors. As CIS‐induced DNA damage primarily activates ATR and not ATM, it explains these findings and suggests that ATR inhibition dictates drug synergy for CIS treatment in OS [38]. Furthermore, in the current study Western Blotting analysis confirmed a decrease in phospho‐p53 following ATR inhibition with BAY‐1895344 in CIS‐treated HOS‐143B cells. This suggests that ATR is one of the central kinases involved in the phosphorylation of p53 at serine 15 and its inhibition can independently decrease the phospho‐p53 (S15) levels [37]. Furthermore, Western Blotting analysis revealed a decrease in ATG7 protein levels in BAY‐1895344 and CIS‐treated HOS‐143B cells. This suggests a possible association between phospho‐p53‐mediated DDR and ATG7/autophagy in OS chemoresistance. A study by Lee et al. [39], showed that ATG7 and p53 bind to each other directly and as such, not only does p53 regulate autophagy but ATG7 can regulate p53. However, this needs to be investigated in the context of OS chemoresistance. Interestingly however, the addition of BAY‐1895344 and CIS resulted in a down‐regulation in p62 levels. During autophagy, p62 is degraded and serves as a marker of autophagic flux. However, it has been shown that VX‐970 increases p62 through an ATR‐independent mechanism, as this effect was seen only with VX‐970 and the precursor another ATR inhibitor VE‐821 but not with other ATR inhibitors such as BAY‐1895344 [40]. This suggests off‐target activity linked to their shared structure, with inhibitor assays further suggesting that VX‐970 disrupts late‐stage autophagy.
Flow cytometry analysis of Annexin V/PI‐stained HOS‐143B cells showed that although CIS treatment caused apoptotic cell death, combining an ATR inhibitor alongside CIS treatment significantly increased the incidence of apoptosis. This suggests that ATRi may potentiate the effects of CIS treatment through reversal of the phospho‐p53‐mediated DDR and autophagy in HOS‐143B OS cells (Figure 7). ATR inhibitors have a significant impact on cells active within the cell cycle, such as cancer cells, leading to a sharp increase in the number of apoptotic cells. However, this treatment is ineffective, let alone beneficial for the survival of noncycling cells, such as healthy normal cells [41]. A proof‐of‐principle study shows that ATR inhibition in pancreatic ductal adenocarcinoma xenografts increased sensitivity to radiation as well as gemcitabine treatment without exerting cytotoxicity to normal cells [42].
ATRi blocked phosphorylation of p53 (S15) and enhanced cell death in CIS‐treated OS cells.
BAY‐1895344 is an orally administered ATR inhibitor by Bayer AG (Germany) that entered clinical trials for the treatment of lymphomas and solid cancers in 2017. Currently, ClinicalTrials.gov shows planned clinical trials investigating the safety and efficacy of BAY‐1895344 for treatment of several other cancers, such as advanced pancreatic and ovarian cancer [38, 43]. ATR inhibitors show promise as adjuvant therapies in OS, but several challenges must be addressed for clinical translation. These include potential toxicity to normal proliferating cells, OS tumor heterogeneity, and compensatory resistance mechanisms such as ATM or CHK1 activation.
The current study has shown that autophagy may play a role in determining the fate of OS patients, as higher levels of autophagy are detected in advanced OS tumors and are also found to be associated with poorer survival and metastasis. Chemotherapy increases autophagy levels, restoring cellular homeostasis that allows for OS tumor progression. The inhibition of autophagy by ATG7 KO as well as the inhibition of phospho‐p53 (S15) via inhibitors of ATR protein kinase enhanced CIS sensitivity in OS. These findings could ultimately lead to novel adjuvant therapy for OS, lowering the dosage of CIS required by combining it with an ATR kinase inhibitor. This could reduce the harsh side effects of CIS treatment as well as overcome chemoresistance and tumor recurrence, potentially improving the overall survival of OS patients.
Author Contributions
Janice S. Pereira: data curation (equal), formal analysis (equal), investigation (lead), methodology (equal), project administration (equal), validation (equal), writing – original draft (lead), writing – review and editing (equal). Gabriel Rosa: data curation (equal), formal analysis (equal), investigation (equal), methodology (equal), writing – original draft (equal), writing – review and editing (equal). Aine Pears: data curation (equal), formal analysis (equal), investigation (equal), writing – original draft (equal), writing – review and editing (equal). Beata Burczynska: formal analysis (equal), investigation (equal), methodology (equal), software (equal), supervision (equal), writing – original draft (equal), writing – review and editing (equal). Britta Stordal: data curation (equal), formal analysis (equal), funding acquisition (equal), investigation (equal), methodology (equal), supervision (equal), writing – original draft (equal), writing – review and editing (equal). Scott J. Roberts: conceptualization (lead), data curation (supporting), formal analysis (supporting), investigation (lead), methodology (lead), supervision (supporting), writing – original draft (supporting), writing – review and editing (supporting). Helen C. Roberts: conceptualization (lead), data curation (supporting), formal analysis (supporting), funding acquisition (lead), investigation (lead), methodology (lead), project administration (lead), resources (lead), supervision (lead), validation (lead), writing – original draft (supporting), writing – review and editing (supporting).
Funding
This research was part‐funded by the Bone Cancer Research Trust.
Ethics Statement
This research was approved by the Middlesex University Natural Sciences Research Ethics Committee (Reference 7653).
Conflicts of Interest
The authors declare no conflicts of interest.
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