The drug-tolerant persister (DTP) state enables cancer cells to evade cytotoxic stress from anticancer therapy. However, the mechanisms governing DTP generation remain poorly understood. Here, we observed that lung adenocarcinoma (LUAD) cells and organoids entered a quiescent DTP state to survive MAPK inhibitor treatment. DTP cells following MAPK inhibition underwent a metabolic switch from glycolysis to oxidative phosphorylation (OXPHOS). PTEN-induced kinase 1 (PINK1), a serine/threonine kinase that initiates mitophagy, was upregulated to maintain mitochondrial homeostasis during DTP generation. PINK1-mediated mitophagy supported DTP cell survival and contributed to poor prognosis. Mechanistically, MAPK pathway inhibition resulted in MYC-dependent transcriptional upregulation of PINK1, leading to mitophagy activation. Mitophagy inhibition using either clinically applicable chloroquine or depletion of PINK1 eradicated drug tolerance and allowed complete response to MAPK inhibitors. This study uncovers PINK1-mediated mitophagy as a novel tumor protective mechanism for DTP generation, providing a therapeutic opportunity to eradicate DTP and achieve complete responses.

Significance:

DTP cancer cells that cause relapse after anticancer therapy critically depend on PINK1-mediated mitophagy and metabolic reprogramming, providing a therapeutic opportunity to eradicate persister cells to prolong treatment efficacy.

Targeted therapies have been wildly applied in cancer treatment. Although they initially elicit striking success, the emergence of resistance after prolonged tumor regression is frequent. Most studies on understanding resistance assume that cancer cells are either sensitive or resistant. But there is actually a quiescent “drug-tolerant persister (DTP)” state that lies between sensitivity and resistance (1, 2). DTP cancer cells represent a reservoir for resistance development. Unlike irreversible resistant cells, which are driven by genetically inheritable mechanisms, DTP cancer cells are supported by nongenetic mechanisms and are hopeful to be reversed (3, 4). Targeting DTP cancer cells may eventually prevent the development of resistance. However, applicable therapeutic vulnerabilities are hindered by the fact that mechanisms governing DTP generation remain poorly elucidated.

Therapeutic agent treatment causes fatal cellular stresses related to apoptosis initiation, as well as metabolic and oxidative stress (5). To survive under these circumstances, DTP cells must co-opt adaptive responses to deal with stress (6, 7). Thus, determining how DTP cancer cells adapt to the harsh environment may be key to identifying their vulnerabilities. Mitochondria are crucial for cellular homeostasis (8). The complex interplay of mitochondrial signaling makes mitochondria confluent in many cellular processes, such as cell death, regulation of metabolism, and generation of reactive oxygen species (ROS; ref. 9). Stressful environmental conditions, including cytotoxic agents, can lead to mitochondrial dysfunction. These dysfunctional mitochondria are lethal for cell survival (10). Mitophagy, an evolutionarily conserved cellular process that eliminates dysfunctional mitochondria, serves as a protective mechanism that ensures cell survival (11). The contribution of mitophagy to functional mitochondria is also essential for the metabolic switch between aerobic glycolysis and mitochondrial respiration (12, 13). However, as a key confluence of stress response and metabolic alteration, whether and how mitophagy is involved in DTP remains unknown.

The MAPK/ERK pathway (hereafter referred to as MAPK) drives nearly one third of the tumors. Targeting this signaling has been applied to cure tumors (14). In patients receiving MAPK inhibitor therapy, the clinical benefit is usually temporary and resistance inevitably occurs (15). In this study, we identified that MAPK inhibitor treatment induced a quiescent DTP state in lung adenocarcinoma (LUAD) cancer cells and LUAD patient-derived organoids (PDO). These DTP cells exhibited OXPHOS-dependent metabolism. During DTP state generation, mitophagy was activated by PINK1 upregulation, supporting DTP cells to accomplish metabolic switch and maintain homeostasis. PINK1-mediated mitophagy supported DTP cell survival and contributed to poor prognosis. Mitophagy inhibition using either clinically approved chloroquine or depletion of PINK1 enhanced the initial efficacy of MAPK inhibitors and led to prolonged treatment responses. Together, we identified PINK1-mediated mitophagy as a novel vulnerability of DTP cells during MAPK pathway–targeting therapy, which could be targeted by clinically applicable chloroquine.

Cell lines, PDO culture, and transfection

Human cancer cell lines A549, H460, HCT116, and HT29 were obtained from the Shanghai Cell Bank of the Chinese Academy of Sciences. MiaPaCa-2 and Panc-1 were kind gifts from Dr. Yaqing Li. All cell lines were cultured in DMEM (Vigonob) supplemented with 10% fetal bovine serum (Lonsera) and 1% penicillin/streptomycin (Beyotime) with 5% CO2 at 37°C. All cell lines were authenticated by short tandem repeat analysis and examined every 6 months for Mycoplasma. Two cases of LUAD tissues for organoid culture were derived from LUAD patients after surgery at the Sun Yat-Sen Memorial Hospital. After surgery, LUAD tissues were cut into pieces (1–2 mm) and enzymatically processed for 1 hour at 37°C. The released floating malignant cells were collected by centrifugation, resuspended in Matrigel (Corning), plated in 24-well plates, and expanded as 3D organoids, as previously described (16). The 3D PDOs were passaged weekly and maintained in Advanced DMEM/F12 (Life Technology) supplemented with EGF (50 ng/mL, Life Technologies), FGF (10 ng/mL; Life Technologies), Y-27632 (10 μmol/L, MedChemExpress), N2 (1X; Life Technologies), B27 (1X, Life Technologies). Plasmid transfection was performed using ViaFect transfection reagent (Promega). siRNA transfection was performed using Lipofectamine RNAiMAX reagent (Life Technologies) according to the manufacturer's instructions. siRNA were purchased from GenePharma. siRNA sequences are shown in Supplementary Table S1.

Reagents, plasmid, and generation of stable cells using lentiviral infection

The product code and suppliers for all commercial antibodies and chemical reagents used in this study are listed in Supplementary Table S2. The dilution ratios of antibodies used for immunoblotting, immunofluorescence, and IHC were 1:1,000, 1:200, and 1:100, respectively. Unless specified otherwise, chemical reagents were used at a concentration of 100 nmol/L for trametinib and PD0325901; 10 μmol/L for dabrafenib, vemurafenib, and ARS1620; 50 μmol/L for chloroquine and 3-MA; and 100 ng/mL for doxycycline. The pLVX-H2B-GFP plasmid was a gift from Dr. Kaishun Hu. The pHAGE-mt-mkeima plasmid was a gift from Richard Youle (RRID: Addgene_131626). The pEGFP-parkin-WT plasmid was a gift from Edward Fon (RRID: Addgene_45875). Full-length cDNAs coding for MYC and PINK1 were obtained from the human cancer cell line A549 by PCR and then cloned into the pLVX vector. The promoter region of PINK1 (∼1,000 bp) was synthesized in vitro and cloned into the luciferase reporter vector pGL3-basic vector (Promega). shRNAs targeting PINK1 were synthesized in vitro (IGE Biotechnology). shPINK1 #1 was cloned into the pLKO.1 and pLKO-TER vector, and shPINK1 #2 was cloned into the pLKO.1 vector. The PINK1 targeting sequences are listed in Supplementary Table S1. To generate stable cells by lentiviral infection, the destination vectors were packaged into lentiviruses with packaging plasmids pMD2.G (RRID: Addgene_12259) and psPAX2 (RRID: Addgene_12260) by transfecting into 293T cells. A549 and H460 cells were infected with lentiviruses supernatant followed by selection with media containing puromycin (2 μg/mL) or FACS (for cells stably expressing histone-GFP).

Generation of DTP cells and viability assay

For LUAD cell lines (A549 and H460), cells that reached a density of approximately 80% in 100 mm dishes (approximately 5,000,000 cells) were treated with 100 nmol/L trametinib, and thereafter fresh drug-containing complete culture medium was changed every 2 days. For two cases of LUAD PDOs, drug-naïve organoids were dissociated with TrypLE Express (Gibco) and single cells were resuspended in Matrigel (Corning; around 500 cells per μL). The 20-μL drops containing organoid cells were seeded on a prewarmed 24-well plate and allowed to grow until the majority of organoids reached nearly 100 μm. Then, the organoids were cultured in a complete culture medium containing 100 nmol/L trametinib, and fresh drug-containing complete culture medium was changed every 2 days. After being treated with trametinib for 10 to 30 days, most cells (∼90%) were killed or detached by the prescribed treatment, and the residual cells gradually exhibited a stable slow-cycling phenotype under drug treatment. These surviving cells were defined as persister cancer cells and used for follow-up exploration. To explore the phenotypic change after drug withdrawal, persister cells were recultured in a drug-free medium for at least 3 days. Cell viability assays were performed using the Cell Counting Kit 8 (CCK8, Fudebio) for cancer cell lines and the CellTiter-Glo 3D cell viability kit (Promega) for PDOs according to the manufacturer's instructions.

Carboxyfluorescein diacetate succinimidyl ester assay

Cell lines or PDOs were labeled with carboxyfluorescein diacetate succinimidyl ester (CFSE; 25 μmol/L) at 37°C for 20 minutes. The stained cells were quenched with fresh culture medium and incubated with 5% CO2 at 37°C. Cultured cells were dissociated with trypsin and subjected to flow cytometry to measure the fluorescence intensity of CFSE on days 1, 3, and 5 after CFSE labeling. The CFSE median fluorescence intensity on each day was normalized to that on day 1.

H2B-GFP cell labeling and imaging

Cancer cells were engineered to stably express histone H2B-GFP by lentivirus-mediated transduction. Cells were counted, seeded at 5,000 cells per dish, and subjected to trametinib treatment at the indicated time points. Cells were imaged every 24 hours. Six adjacent images were captured for each dish per experiment. Cells are counted through the GFP signal.

Immunofluorescence staining

Cells or organoids grown on glass coverslips were washed once with PBS and fixed with 4% paraformaldehyde for 20 minutes followed by preextraction with 0.5% Triton X-100 for 8 minutes at room temperature. After being blocked with 5% bovine serum albumin (BSA) for 1 hour, samples were incubated with indicated antibodies at 4°C overnight and washed with PBST for 3 times followed by secondary antibodies staining at room temperature for 1 hour. After being stained with DAPI, the samples were subjected to confocal imaging using Zeiss LSM 800 (RRID: SCR_015963). Primary antibodies were summarized in Supplementary Table S2.

For BrdUrd analysis, cells or organoids grown on glass coverslips were incubated with complete culture medium containing 10 μmol/L BrdUrd for 24 hours before immunofluorescence staining. During the immunofluorescence staining procedure, the samples for BrdUrd staining were additionally incubated in 1 M HCL for 1 hour and neutralized with 0.1 M sodium borate buffer pH 8.5 for 30 minutes at room temperature before being blocked with 5% BSA.

Cell-cycle analysis

Cells or organoids were dissociated with trypsin, washed once with PBS, and fixed in 70% ice-cold ethanol overnight. Fixed samples were washed twice with PBS, followed by RNase A treatment and propidium iodide (0.05 mg/mL) staining. PI-labeled samples were analyzed in a flow cytometer (CytoFLEX, Beckman, RRID: SCR_019627) with excitation at 488 nm laser and emission at 585/42 nm filters.

Seahorse metabolic flux analysis

Cells were plated at respective optimal densities in Seahorse XF well plated 1 day prior to the measurement. These cells were subjected to XFe96 Extracellular Flux Analyzer (RRID: SCR_019545) for measuring oxygen consumption rate (OCR) and extracellular acidification rate (ECAR). The cellular energy metabolism phenotype was determined using an energy phenotype test kit (Agilent) according to the manufacturer's instructions. This test simultaneously measures mitochondrial respiration and glycolysis under baseline and stressed conditions induced by oligomycin (1 μmol/L) and FCCP (0.5 μmol/L). The mitochondrial stress test was performed using the mitochondrial stress test kit (Agilent) according to the manufacturer's instructions. This test measures mitochondrial respiration levels under basal conditions and following the addition of oligomycin (1 μmol/L), FCCP (0.5 μmol/L) and a mixture of rotenone/antimycin A (0.5 μmol/L). Data were normalized to cellular protein levels.

Label-free phosphoproteomics

Sample preparation, LC-MS/MS analysis, and pairwise normalization for label-free quantitative phosphoproteomics were performed according to a previously published protocol (17). Phosphorylated peptides were enriched using a High-Select Phosphopeptide Enrichment kit (Thermo Fisher) according to the manufacturer's instructions. Data were analyzed using the “Wu Kong” platform (https://www.omicsolution.com/wkomics/main; ref. 18).

RNA isolation and quantitative RT-PCR

Total RNA was extracted using RNAiso Plus (Takara), and 500-ng aliquots were used for cDNA synthesis using the PrimeScript RT reagent (Takara). The cDNA templates were subjected to amplification using SYBR qPCR Master Mix (Vazyme) in a CFX96 qPCR System (Bio-Rad). Data were analyzed by the threshold cycle (Ct) comparative method and normalized to GAPDH. Primers used are listed in Supplementary Table S1.

Western blot analysis

Immunoblots were performed from whole-cell lysates prepared using RIPA buffer supplemented with protease and phosphatase inhibitors (Bimake). After boiling in 2× SDS loading buffer, the lysates were resolved on SDS-PAGE assay and transferred to PVDF membranes (Millipore). After blocking with 5% milk in PBST, the membranes were incubated with primary antibodies overnight at 4°C, except for the primary antibodies against PINK1 (Proteintect) and CD133 (Proteintect) for 1.5 hours at room temperature. The membranes were incubated with secondary antibodies for 1 hour at room temperature. Indicated antibodies used are listed in Supplementary Table S2.

Mito-keima mitophagy analysis

Cells were infected with a lentivirus harboring the mito-mkeima and grown for several days. After indicated treatment, cells expressing mito-mkeima were trypsinized, resuspended in the PBS buffer, and analyzed by flow cytometer. Measurements of lysosomal mito-keima were made dual-excitation ratiometric pH measurements at 405 nm (pH 7) and 640 nm (pH 4) lasers with 660/20 nm emission filters. For each sample, 10,000 events were collected. Data were analyzed and visualized using FlowJo (RRID: SCR_008520).

ChIP-qPCR

ChIP assay was performed as previously described (19). Cells were fixed with 1% formaldehyde, and nuclei were extracted. The chromatin/DNA complex was sheared by a sonicator. The sonicated lysates were cleared and incubated overnight at 4°C with magnetic beads coupled with an anti-MYC antibody (Proteintect). DNA was eluted and analyzed by qPCR. Primers used are listed in Supplementary Table S1.

Mitochondrial membrane potential analysis and apoptosis assay

Cells were subjected to designated treatments and then collected for JC-10 and annexin V analyses. The mitochondrial membrane potential analysis was performed using the JC-10 probe (4A BIOTECH). Cells were incubated for 20 minutes at 37°C in serum-free DMEM containing 20 μmol/L JC-10 dye. After incubation, cells were washed with PBS once and analyzed by flow cytometer with excitation at 488 nm laser and emission filters at both 525/40 nm and 585/42 nm. For apoptosis analysis, annexin V assay was performed using the annexin V–FITC Apoptosis Detection Kit (UElandy) according to the instruction of the manufacture. Briefly, cells were incubated for 10 minutes at room temperature in a binding buffer containing 5 μL FTIC-conjugated annexin V antibody and 5 μL propidium iodide. After incubation, cells were analyzed by flow cytometer with excitation at 488 nm laser and emission filters at both 525/40 nm and 585/42 nm.

ROS detection

The ROS levels were assessed by measuring intracellular oxidation of 2,7-dichlorodihydrofluorescence (DCFH, Beyotime). Cells were incubated with DCFH-DA probes for 20 minutes at 37°C and washed with serum-free medium. The fluorescence signal was measured by flow cytometer with excitation at 488 nm laser and emission filters at 525/40 nm.

NADP+/NADPH measurement

The NADP+/NADPH redox ratio was measured using the NADP+/NADPH assay kit with WST-8 (Beyotime) according to the manufacturer's protocol. Cells were harvested and lysed according to instructions. Samples were separated into two parts. One part underwent 60°C for 30 minutes to eliminate NADP+. Samples were treated with a working solution containing GAPDH for 10 minutes at 37°C and incubated with a working solution containing WST-8 for 30 minutes at 37°C in dark. The absorption values were detected at 450 nm wavelength.

Single-cell sequencing data analysis

Single-cell RNA-seq data were obtained from the GEO database (GSE134839 and GSE116237). Data were processed by quality control, data mapping and normalization, unsupervised clustering, and aggregated scoring, according to a published procedure (20). In brief, three parameters were used for assessing cell quality: nFeature_RNA, nCount_RNA, and percent.MT. Cells with poor quality were filtered. After read quality assessment, UMI counts were converted to transcripts-per-10,000 using the Seurat package with the “LogNormalize” method. Unsupervised clustering analyses were performed on gene-expression matrices using R package Seurat version 3.1.2. UMAPs and feature plots were generated using Seurat. Aggregated gene set score was calculated using “AddModuleScore()” based on the averaged expression.

Clonogenic assay

For clonogenic assay under the indicated drug treatment conditions, 5,000 cells per well were seeded into 12-well plates and allowed to adhere overnight. The cells were subjected to the indicated drug treatment for 48 hours, cultured in a drug-free medium for 7 days, and then fixed using 4% PFA for 15 minutes followed by 0.1% crystal violet staining for 15 minutes. The clones stained with crystal violet were dissolved in acetic acid and detected at the absorbance of 570 nm.

Animal studies

All BALB/c nude mice (6-week-old, average body weight 18 g) were obtained from the Laboratory Animal Center of Sun Yat-Sen University. All mice were housed in cages containing aspen chip bedding in rooms under standard conditions in accordance with the institutional guidelines of the Animal Care and Use Committee of Sun Yat-Sen University. For the subcutaneous xenograft model, A549 cells (5,000,000 cells/mouse) and H460 cells (3,000,000 cells/mouse) were suspended in cold PBS, mixed at a 1:1 dilution of Matrigel solution (BD Biosciences), and then subcutaneously administered to the rear flank of the mice. Tumor size was measured using digital calipers and evaluated using the formula: tumor volume (mm3) = length × width × width/2. When the volume reached ∼100 mm3, the mice were randomized into the indicated groups and received the indicated treatment. To determine the impact of doxycycline-induced PINK1 depletion on trametinib therapeutic effect in vivo, doxycycline (20 mg/kg body weight) or saline as negative control were injected intraperitoneally twice per week. Trametinib was given in a daily diet at 2 ppm (2 mg of trametinib was mixed per kg of chow). Treatments began on day 12 after inoculation and lasted for 6 to 8 weeks. To determine the impact of chloroquine on trametinib therapeutic effect in vivo, chloroquine (50 mg/kg body weight) or saline as negative control was injected intraperitoneally twice per week. Trametinib was administrated in a daily diet at 2 ppm. Treatments began on day 12 and stopped on day 45 after inoculation. The tumor size and body weight of mice were recorded twice a week.

For the response evaluation criteria in solid tumors (RECIST) assessment, we compared the percentage change in tumor volume during the period from the beginning to the end of drug treatment. Mice whose tumor volume increased more than 20% during this period were defined as progressive disease (PD). Mice whose tumor volume increased no more than 20% or decreased no more than 30% were defined as stable disease (SD). Mice whose tumor volume decreased more than 30% were defined as partial response (PR). Mice whose tumors were undetectable at drug cessation were defined as complete response (CR).

IHC

After deparaffinization with xylene, rehydration in graded ethanol, immersion in 0.3% hydrogen peroxide, and heat-mediated antigen retrieval in citric acid at pH 6.0, tissue sections were incubated with primary antibodies at 4°C overnight, and labeled with HRP-conjugated secondary antibody at room temperature for 20 minutes. Finally, sections were developed in DAB solution under microscopic observation and counterstained with hematoxylin. According to both the proportion and intensity of positive tumor cells, the expression of PINK1 in LUAD tissues was scored as negative (score 1), weakly positive (score 2), moderately positive (score 3), and strongly positive (score 4). Scores 1 and 2 were defined as low PINK1 levels, and scores 3 and 4 were defined as high PINK1 levels.

Synergy assay

To evaluate synergy in vitro, cells were seeded into 96-well plates and allowed to adhere overnight. After the indicated drug treatment for 72 hours, cell viability was measured. Data were analyzed using Combenefit software (21).

Statistical analyses

Statistical analyses were performed using the GraphPad Prism (RRID: SCR_002798) version 7. P values <0.05 were considered statistically significant. Data are presented as the mean ± SEM. All statistical tests and sample sizes are described in the figure legends.

Study approval

The study was approved by the medical ethics committee of Sun Yat-Sen Memorial Hospital, and written informed consent was obtained from all patients. All animal experiments were conducted according to protocols approved by the Institutional Animal Care and Use Committee of Sun Yat-Sen University.

Data availability

The data generated in this study are available in the article and its supplementary files. Additional data is available upon request from the corresponding author.

MAPK pathway inhibitor treatment induces a quiescent but reversible DTP state

To model the MAPK inhibitors response and inefficacy, we examined the therapeutic response to MAPK pathway inhibition therapy in both LUAD cell lines and PDOs (Fig. 1A). Although most cells were killed by trametinib (MEK inhibitor) treatment, a few residual cancer cells persisted after 10 to 30 days of treatment (Fig. 1B). These residual cancer cells remained in a nonproliferative state throughout the treatment, and we named them “persister.” Compared with the parental cells, the persister cells exhibited tolerance to trametinib as shown by the cell viability assay (Supplementary Fig. S1A). These persister cells were similar to those described as DTP cells, which enter a quiescent state to escape cytotoxic stress from anticancer therapy (1). We further verified the quiescent character of these persister cells using the proliferation-dependent fluorescent cell-tracking dye CFSE, a dye whose reduction in fluorescence intensity reflects cell cycling. The persister cells showed slow-cycling during trametinib treatment (Fig. 1C). The slow-cycling state of the persister cells was reversible, because they began to proliferate following drug withdrawal (Fig. 1C). Live cells expressing histone 2B-GFP (H2B-GFP) imaging also suggested that the persister cells exhibited significant growth suppression under trametinib treatment, but resumed growth upon drug withdrawal (Fig. 1D). BrdUrd labeling showed that persister cells could not be labeled with BrdUrd, suggesting that their DNA synthesis was inactive (Fig. 1E; Supplementary Fig. S1B). Ki67 staining showed that the persister cells treated with trametinib were mostly negative for Ki67, indicating that they were arrested in the G0 phase (Fig. 1F; Supplementary Fig. S1C). Likewise, cell-cycle analysis demonstrated that persister cells underwent G0–G1 arrest under trametinib treatment (Supplementary Fig. S1D). Cell-cycle arrest of persister cells was also confirmed by increased p21 and p27 expression (Supplementary Fig. S1E). DTP state has been characterized to increase expression of stemness markers and activate epithelial–mesenchymal transition (EMT; ref. 22). We observed the upregulation of CD44 and CD133, as well as induction of epithelial–mesenchymal transition (EMT), in persister cells (Supplementary Fig. S1E). Taken together, our observations in both cell lines and PDOs indicate that LUAD cells enter a quiescent but reversible DTP state to survive under MAPK inhibitor treatment.

Figure 1.

MAPK inhibitor treatment induces a quiescent but reversible DTP state, accompanied by a metabolic shift from glycolysis to OXPHOS. A, Schematic diagram showing generation of DTP state by 10 to 30 days treatment with 100 nmol/L trametinib to two LUAD cell lines as well as two LUAD PDOs. B, Image of parental cells or persister cells. Scale bars, 100 μm. C, Cell-tracking CFSE dye, whose fluorescence intensity gradually decreases during cell division, was used to measure the relative rate of cell proliferation. D, Persister cells expressing H2B-GFP were subjected to trametinib (Tra) treatment or drug washout at the indicated time points. Cell numbers were quantified over 25 days. E and F, Immunofluorescence staining of BrdUrd (E) and Ki67 (F) among the parental cells, the persister cells, and the regrown cells after drug washout. For BrdUrd labeling, cells were treated with 10 μmol/L BrdUrd for 24 hours. Scale bars, 10 μm. G, Cell energy phenotype analyses of the indicated cells through real-time quantifications of ECAR and OCR at baseline or stressed with oligomycin/FCCP. H, Mitochondrial respiration was measured in the parental cells, the persister cells, and the regrown cells after drug washout. Maximum respiration and spare respiration were quantified based on OCR measurements. I, Cell viability assays were performed after 48 hours of either 2-DG treatment (5 mmol/L) or rotenone treatment (20 μmol/L). Data are presented as the mean ± SEM. n = 3 for C, G, and H; n = 6 for D and I. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by ANOVA (C) or t test (G–I).

Figure 1.

MAPK inhibitor treatment induces a quiescent but reversible DTP state, accompanied by a metabolic shift from glycolysis to OXPHOS. A, Schematic diagram showing generation of DTP state by 10 to 30 days treatment with 100 nmol/L trametinib to two LUAD cell lines as well as two LUAD PDOs. B, Image of parental cells or persister cells. Scale bars, 100 μm. C, Cell-tracking CFSE dye, whose fluorescence intensity gradually decreases during cell division, was used to measure the relative rate of cell proliferation. D, Persister cells expressing H2B-GFP were subjected to trametinib (Tra) treatment or drug washout at the indicated time points. Cell numbers were quantified over 25 days. E and F, Immunofluorescence staining of BrdUrd (E) and Ki67 (F) among the parental cells, the persister cells, and the regrown cells after drug washout. For BrdUrd labeling, cells were treated with 10 μmol/L BrdUrd for 24 hours. Scale bars, 10 μm. G, Cell energy phenotype analyses of the indicated cells through real-time quantifications of ECAR and OCR at baseline or stressed with oligomycin/FCCP. H, Mitochondrial respiration was measured in the parental cells, the persister cells, and the regrown cells after drug washout. Maximum respiration and spare respiration were quantified based on OCR measurements. I, Cell viability assays were performed after 48 hours of either 2-DG treatment (5 mmol/L) or rotenone treatment (20 μmol/L). Data are presented as the mean ± SEM. n = 3 for C, G, and H; n = 6 for D and I. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by ANOVA (C) or t test (G–I).

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DTP cells arising from MAPK pathway inhibition undergo metabolic switch from glycolysis to oxidative phosphorylation

Metabolic plasticity and cancer growth are closely related and reciprocally regulated. Cancer cells hijack flexible metabolic phenotypes to survive in unfavorable environments (23). We simultaneously monitored two major energy-producing pathways, glycolysis and oxidative phosphorylation (OXPHOS), by measuring the ECAR and OCR. Compared with the parental cells, the persister cells exhibited higher OCR both at baseline and under stress (oligomycin/FCCP), indicating that they were undergoing a metabolic shift toward OXPHOS (Fig. 1G; Supplementary Fig. S2A). In contrast, the regrown cells after drug washout switched their metabolism back to glycolysis (Fig. 1G; Supplementary Fig. S2A). The metabolic shift toward OXPHOS in persister cells was confirmed by the mitochondrial stress assay, which showed that the maximum respiration and spare respiration capacity were increased in persister cells (Fig. 1H; Supplementary Fig. S2B). Also, the persister cells showed decreased glucose uptake and lactate production compared with their corresponding parental cells and regrown cells after drug washout (Supplementary Fig. S2C and S2D). To explore whether the metabolic shift is crucial for the survival of DTP cells, we examined cell viability after treatment with either 2-DG (glycolysis inhibitor) or rotenone (OXPHOS inhibitor). Both parental cells and the regrown cells after drug washout showed sensitivity to glycolysis inhibition. Conversely, persister cells showed an increased sensitivity to OXPHOS inhibition (Fig. 1I). These results indicate that DTP cells arising from MAPK inhibition exhibit OXPHOS-dependent metabolism.

The establishment of DTP state following MAPK inhibition is dependent on activation of PINK1-mediated mitophagy

To dissect cellular processes and/or key effectors that govern the establishment of the DTP state after MAPK inhibitor treatment, we conducted phosphoproteomics on drug-naïve parental cells and drug-tolerant persister cells (Fig. 2A). As previously stated, the initial response to drug treatment favors DTP generation (24). Therefore, we characterized the phosphoproteomic responses to short-term (72 hours) treatment with trametinib (Fig. 2A). A total of 5,157 phosphopeptides were identified, of which 2,781 were quantified (Supplementary Fig. S3A; Supplementary Table S3). Most of the differentially phosphorylated peptides (|fold change| ≥ 1.5, P < 0.05) changed both in the short-term treated cells and persister cells (Fig. 2B). Phosphorylated ubiquitin (serine 65), a substrate of PINK1 (25), was one of the top candidates for upregulated phosphoproteins (Fig. 2B). Gene ontology (GO) enrichment analysis was performed on these phosphoproteins. As anticipated, Ras signaling and the MAPK cascade were enriched in downregulated phosphoproteins. In accordance with the slow-cycling phenotype of DTP cells, the downregulated phosphoproteins corresponded to G1–S transition and DNA replication (Fig. 2C). Macroautophagy and PINK1/parkin-mediated mitophagy were significantly enriched in the upregulated phosphoproteins (Fig. 2C). Macroautophagy, also known as autophagy, is involved in the removal of misfolded proteins and damaged organelles. PINK1-mediated mitophagy is a pivotal autophagy pathway that mediates the removal of damaged mitochondria. The contribution of PINK1-mediated mitophagy to mitochondrial homeostasis may establish a metabolic switch between aerobic glycolysis and OXPHOS. Autophagy inhibitors including chloroquine or 3-methyladenine (3-MA), disrupted the DTP state induced by MAPK inhibition (Supplementary Fig. S3B). Likewise, PINK1 knockdown through doxycycline-inducible shRNA targeting PINK1 abolished the DTP state induced by MAPK inhibition (Supplementary Fig. S3C and S3D). These results reveal that cancer cells rely on PINK1-mediated mitophagy to enter the DTP state following MAPK inhibition.

Figure 2.

PINK1-mediated mitophagy is induced in response to MAPK pathway inhibitor treatment. A, Schematic procedure of label-free quantitative phosphoproteomics to identify perturbation in phosphoproteome during establishing MAPK inhibitor tolerance. Untreated A549 cells act as a control. B, Scatter plot displays the log2 of fold change in phosphopeptides abundance of trametinib-treated cells for 72 hours vs. parental cells (72 hours vs. control) and persister cells vs. parental cells (persister vs. control). Dotted lines represent |fold change| = 1.5. C, Gene ontology analysis of the upregulated phosphopeptides and downregulated phosphopeptides that change consistently in trametinib-treated cells for 72 hours and the persister cells. D, qRT-PCR analysis of PINK1 in both parental and persister cells treated with trametinib. E, Immunoblotting of PINK1 in both parental and persister cells treated with trametinib for the indicated time. F, Immunoblotting of PINK1 in A549 cells introduced with shRNA targeting either scramble or PINK1. G and H, A549 cells stably expressing shRNA targeting scramble or PINK1 were transfected with GFP-parkin, then treated with either vehicle control or trametinib (100 nmol/L) for 72 hours. Mitochondria were visualized through immunostaining of TOMM20. The colocalization between parkin and TOMM20 was analyzed by confocal microscopy. Right panels in G show the pixel intensity of parkin and TOMM20 from a line. I and J, A549 cells stably expressing shRNA targeting scramble or PINK1 were infected with mito-Keima and then treated with vehicle control or trametinib (100 nmol/L) for 72 hours. FACS was performed to analyze the lysosomal-positive mito-Keima, which was displayed in the upper gate and represented mitophagy levels. Data are presented as the mean ± SEM. n = 3 for D, H, and J. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test.

Figure 2.

PINK1-mediated mitophagy is induced in response to MAPK pathway inhibitor treatment. A, Schematic procedure of label-free quantitative phosphoproteomics to identify perturbation in phosphoproteome during establishing MAPK inhibitor tolerance. Untreated A549 cells act as a control. B, Scatter plot displays the log2 of fold change in phosphopeptides abundance of trametinib-treated cells for 72 hours vs. parental cells (72 hours vs. control) and persister cells vs. parental cells (persister vs. control). Dotted lines represent |fold change| = 1.5. C, Gene ontology analysis of the upregulated phosphopeptides and downregulated phosphopeptides that change consistently in trametinib-treated cells for 72 hours and the persister cells. D, qRT-PCR analysis of PINK1 in both parental and persister cells treated with trametinib. E, Immunoblotting of PINK1 in both parental and persister cells treated with trametinib for the indicated time. F, Immunoblotting of PINK1 in A549 cells introduced with shRNA targeting either scramble or PINK1. G and H, A549 cells stably expressing shRNA targeting scramble or PINK1 were transfected with GFP-parkin, then treated with either vehicle control or trametinib (100 nmol/L) for 72 hours. Mitochondria were visualized through immunostaining of TOMM20. The colocalization between parkin and TOMM20 was analyzed by confocal microscopy. Right panels in G show the pixel intensity of parkin and TOMM20 from a line. I and J, A549 cells stably expressing shRNA targeting scramble or PINK1 were infected with mito-Keima and then treated with vehicle control or trametinib (100 nmol/L) for 72 hours. FACS was performed to analyze the lysosomal-positive mito-Keima, which was displayed in the upper gate and represented mitophagy levels. Data are presented as the mean ± SEM. n = 3 for D, H, and J. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test.

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MAPK pathway inhibition induces PINK1 expression and thus activates mitophagy

Previous studies have shown that PINK1 acts as a sensor for damaged mitochondria and plays a central role in regulating mitophagy (26). To validate the activation of PINK1-mediated mitophagy during DTP establishment, we examined whether MAPK inhibition affected the expression of PINK1. PINK1 levels were higher in persister cells than in parental cells (Fig. 2D and E). We observed that PINK1 levels were upregulated after trametinib treatment in both parental cancer cells and persister cells (Fig. 2D and E). Elevated PINK1 levels following MAPK pathway inhibitor treatments were confirmed in various cancer cell lines and PDOs (Supplementary Fig. S4A and S4B). During mitophagy activation, parkin is recruited to the mitochondria to amplify mitophagy signals. Trametinib treatment resulted in increased recruitment of parkin to the mitochondria (Fig. 2FH). The mitochondrial recruitment of parkin induced by trametinib treatment was reduced in PINK1-deficient cells (Fig. 2FH). Similar results were obtained when examining the colocalization of mitochondria and LC3B, an autophagosome marker (Supplementary Fig. S4C and S4D), suggesting that trametinib-induced mitophagy is inhibited in PINK1-deficient cells. We further used Mito-Keima, a useful tool in the assessment of acidic mitochondrial events (27), to evaluate mitophagy flux (Supplementary Fig. S4E). In response to trametinib treatment, a remarkable increase in mitophagy flux was detected in control cells, which was not observed in PINK1-deficient cells (Fig. 2I and J). These data suggest that inhibition of the MAPK pathway activates mitophagy by stimulating PINK1 expression.

MAPK pathway inhibition relieves the MYC-engaged transcriptional repression of PINK1

Given that PINK1 expression was induced at the mRNA levels by MAPK pathway inhibitor treatments, we speculated PINK1 that was transcriptionally regulated following MAPK pathway inhibition. We predicted potential transcription factors (TF) binding to the PINK1 promoter by overlaying the candidates from three public databases (Fig. 3A). Among these putative TFs, MYC protein stability has been reported to depend on MAPK pathway activation (28). The MYC protein levels decreased after treatment with various MAPK inhibitors (Supplementary Fig. S5A). In addition, we found that the MYC consensus motif was located in a relatively conserved sequence at the transcriptional start site of PINK1 (Supplementary Fig. S5B). Binding peaks of MYC on the promoters of PINK1 were observed in various cancer cells (Fig. 3B) and were validated by ChIP-qPCR assays (Fig. 3C). The enrichment of MYC in PINK1 promoters was blocked by trametinib treatment (Fig. 3C). Both the mRNA and protein levels of PINK1 were elevated when MYC was silenced (Supplementary Fig. S5C) and were suppressed when MYC was overexpressed (Supplementary Fig. S5D). We also observed an inverse correlation between MYC and PINK1 expression in three independent LUAD data sets (Supplementary Fig. S5E; refs. 29, 30). And the expression of “Hallmark MYC Target” gene set enriches in LUAD with low PINK1 expression (Supplementary Fig. S5F). We cloned the PINK1 promoter into the pGL3 luciferase reporter vector and found that the activity of the PINK1 promoter decreased when MYC was overexpressed (Fig. 3D). These findings clarify the MYC-engaged transcriptional repression of the PINK1 promoter. We also noted that silencing MYC promoted trametinib-induced PINK1 upregulation (Fig. 3E), whereas overexpression of MYC blocked PINK1 expression and mitophagy activation following trametinib treatment (Fig. 3FH). Together, we identified a previously unknown mechanism by which PINK1 expression is controlled by MYC-mediated transcriptional repression in the absence of drug treatment. MYC depletion following MAPK pathway inhibitor treatment attenuates the transcriptional repression of PINK1, thus elevating PINK1 expression and activating mitophagy.

Figure 3.

MAPK pathway inhibition relieves the MYC-engaged transcriptional repression of PINK1. A, Venn diagram indicating the overlaps of TFs predicted to regulate PINK1 transcription by three independent databases. B, ChIP-seq signal traces of MYC in PINK1 locus. C, ChIP-qPCR shows the enrichment of MYC on PINK1 promoter in cancer cells treated with either vehicle control or trametinib (100 nmol/L) for 72 hours. The qPCR primers are designed to anneal to the sequence within the binding peak of MYC on the PINK1 promoter. D, Luciferase reporter activity of promoter constituents of PINK1 measured in cancer cells stably expressing either empty vector (vector) or plasmid encoding MYC (oeMYC). Renilla luciferase control plasmid was cotransfected with either pGL3-basic vector or pGL3-PINK1 promoter plasmid and acted as a normalization control. E, qRT-PCR analysis and immunoblotting of PINK1 in cancer cells transfected with either siNC or siMYC after 72 hours of vehicle or trametinib (100 nmol/L) treatment. F, qRT-PCR analysis and immunoblotting of PINK1 in cancer cells stably expressing either empty vector or plasmid encoding MYC after 72 hours of either vehicle or trametinib (100 nmol/L) treatment. G and H, Mitophagy levels measured in A549 and H460 stably expressing either empty vector or plasmid encoding MYC after 72 hours of vehicle or trametinib (100 nmol/L) treatment. Data are presented as the mean ± SEM. n = 8 for C; n = 3 for D, E, F, and H. **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test.

Figure 3.

MAPK pathway inhibition relieves the MYC-engaged transcriptional repression of PINK1. A, Venn diagram indicating the overlaps of TFs predicted to regulate PINK1 transcription by three independent databases. B, ChIP-seq signal traces of MYC in PINK1 locus. C, ChIP-qPCR shows the enrichment of MYC on PINK1 promoter in cancer cells treated with either vehicle control or trametinib (100 nmol/L) for 72 hours. The qPCR primers are designed to anneal to the sequence within the binding peak of MYC on the PINK1 promoter. D, Luciferase reporter activity of promoter constituents of PINK1 measured in cancer cells stably expressing either empty vector (vector) or plasmid encoding MYC (oeMYC). Renilla luciferase control plasmid was cotransfected with either pGL3-basic vector or pGL3-PINK1 promoter plasmid and acted as a normalization control. E, qRT-PCR analysis and immunoblotting of PINK1 in cancer cells transfected with either siNC or siMYC after 72 hours of vehicle or trametinib (100 nmol/L) treatment. F, qRT-PCR analysis and immunoblotting of PINK1 in cancer cells stably expressing either empty vector or plasmid encoding MYC after 72 hours of either vehicle or trametinib (100 nmol/L) treatment. G and H, Mitophagy levels measured in A549 and H460 stably expressing either empty vector or plasmid encoding MYC after 72 hours of vehicle or trametinib (100 nmol/L) treatment. Data are presented as the mean ± SEM. n = 8 for C; n = 3 for D, E, F, and H. **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test.

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PINK1 is essential for functional mitochondria and redox homeostasis in the DTP cells

To investigate the role of PINK1 in DTP development, we monitored the levels of PINK1 and OXPHOS at various time points during DTP development. PINK1 levels started to increase 2 days after trametinib treatment and remained high thereafter (Supplementary Fig. S6A). OCR levels initially decreased in response to trametinib treatment and started to recover 4 to 5 days after drug treatment, suggesting the OXPHOS phenotype was impaired at the initial stage of drug treatment (Supplementary Fig. S6B). Based on these observations, we assumed that PINK1 might be important for the metabolic switch during DTP generation. To validate this hypothesis, we generated PINK1 overexpressed and empty vector control cell lines from naïve parent cells (Supplementary Fig. S7A) and assessed the effect of PINK1 overexpression on mitochondrial status. Trametinib treatment induced the loss of mitochondrial membrane potential (MMP), which was recovered by PINK1 overexpression (Supplementary Fig. S7B). PINK1 overexpressed cells also exhibited an increased mitochondrial respiration capacity (Supplementary Fig. S7C). We examined MMP in persister cells transfected with a doxycycline-inducible shRNA targeting PINK1. There was little change in MMP in persister cells following trametinib treatment (Fig. 4A), indicating that they were capable of handling mitochondrial damage. In contrast, PINK1-depleted persister cells failed to maintain MMP after trametinib treatment (Fig. 4A). Both PINK1 knockdown and mitophagy inhibition impaired mitochondrial respiration capacity (Fig. 4B; Supplementary Fig. S7D). These results suggest that PINK1 upregulation is essential for maintaining functional mitochondria, which is a prerequisite of persister cells to complete the metabolic shift toward OXPHOS.

Figure 4.

PINK1 is essential for functional mitochondria and redox homeostasis in the persister cells. A, MMP detection through JC-10 probe dying in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. A representative FACS spectrum is shown in which the population of cells in the right and bottom quadrants contain decreased MMP. B, Mitochondrial respiration was measured in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, before and after 72 hours of stimulation with 100 ng/mL doxycycline. C, Apoptosis was measured in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. D, Cell viability of control persister cells (−dox) and doxycycline-induced PINK1 knockdown persister cells (+dox) treated with H2O2 for 24 hours. E, ROS measurement in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. F, Redox balance between NADP/NADPH was monitored in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. Data are presented as the mean ± SEM. n = 3 for A, B, C, D, E, and F. *, P < 0.05; **, P < 0.01; ***, P < 0.001, as determined by the t test.

Figure 4.

PINK1 is essential for functional mitochondria and redox homeostasis in the persister cells. A, MMP detection through JC-10 probe dying in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. A representative FACS spectrum is shown in which the population of cells in the right and bottom quadrants contain decreased MMP. B, Mitochondrial respiration was measured in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, before and after 72 hours of stimulation with 100 ng/mL doxycycline. C, Apoptosis was measured in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. D, Cell viability of control persister cells (−dox) and doxycycline-induced PINK1 knockdown persister cells (+dox) treated with H2O2 for 24 hours. E, ROS measurement in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. F, Redox balance between NADP/NADPH was monitored in the persister cells introduced with a doxycycline-inducible shRNA targeting PINK1, in response to indicated treatments for 72 hours. Data are presented as the mean ± SEM. n = 3 for A, B, C, D, E, and F. *, P < 0.05; **, P < 0.01; ***, P < 0.001, as determined by the t test.

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We also noted that PINK1 knockdown resulted in substantial apoptosis following trametinib treatment in persister cells (Supplementary Fig. S3C and S3D; Fig. 4C), indicating that PINK1 expression was essential for the survival of DTP cells. OXPHOS produces the majority of ROS. The metabolic shift toward OXPHOS should cause greater exposure of persister cells to oxidative stress. DTP generation has been shown to depend critically on antioxidant programs (31, 32). Thus, we investigated the role of PINK1-mediated mitophagy with respect to redox homeostasis in persister cells. When exposed to H2O2, an exogenous source of oxidative stress, PINK1 knockdown persister cells failed to buffer the cytotoxicity of H2O2 (Fig. 4D). We also found increased ROS production and disturbed redox balance after trametinib treatment in PINK1-knockdown persister cells, whereas little change was observed in control persister cells (Fig. 4E and F). The by-effect of doxycycline was excluded in persister cells without tetracycline-responsive shRNA (Supplementary Fig. S8A and S8B). These findings suggest that activation of PINK1-mediated mitophagy is necessary for redox homeostasis and cell survival in MAPK inhibitor-tolerant persisters.

Next, we evaluated the association between PINK1 expression and the quiescent phenotype of the DTP state. PINK1 overexpression inhibited cellular proliferation in a mitochondrial activity-dependent manner, as measured by CFSE staining, Ki67 expression, and BrdUrd labeling (Supplementary Fig. S9A–S9C). DTP state has been characterized by G0-like and embryonic diapause-like transcriptional programs (2). Using the gene-expression data from two independent data sets (GSE31210 and GSE10245), we found that hallmarks of “Toshihiko G0 Down” (33) and “Rehman Diapause Down” (1) gene sets decreased in LUAD with low PINK1 expression (Supplementary Fig. S9D and S9E), suggesting that PINK1 expression positively correlates with the hallmarks of DTP state. We also analyzed single-cell RNA sequencing data sets of DTP cells persisting MAPK inhibitors and EGFR-TKI (20, 34). Compared with untreated cells, DTP cells were enriched in cell population with high expression of PINK1. In addition, the DTP population with high PINK1 expression showed upregulation of the G0 gene set (35), OXPHOS gene set, and mitophagy gene set (Supplementary Fig. S9F–S9I).

Overall, these observations indicate that PINK1-mediated mitophagy supports a quiescent DTP state, by maintaining mitochondrial homeostasis.

PINK1 contributes to MAPK inhibitors' therapeutic inefficacy and tumor relapse

These findings prompted us to characterize the role of PINK1 in MAPK inhibitor inefficacy. Utilizing data from the Genomics of Drug Sensitivity in Cancer (GDSC), we classified LUAD cell lines based on their PINK1 expression and queried their IC50 value to various MAPK inhibitors. Cells with high PINK1 expression (top 10) were less responsive to MAPK inhibitors than cells with low PINK1 expression (bottom 10; Fig. 5A). PINK1 expression was positively correlated with IC50 values of various MAPK pathway inhibitors (Supplementary Fig. S10A). Although PINK1 depletion did not cause an obvious change in cell proliferation in the absence of MAPK pathway inhibitors, it resulted in increased therapeutic responses to various MAPK pathway inhibitors (Fig. 5B; Supplementary Fig. S10B and S10C). In addition, PINK1 knockdown cells showed greater apoptosis after treatment with MAPK pathway inhibitors (Fig. 5C). To study the impact of PINK1 depletion on the DTP state in vivo, cell-derived xenograft models were generated by subcutaneously inoculating A549 cells transfected with doxycycline-inducible shRNA targeting PINK1 (Supplementary Fig. S10D). Trametinib treatment in vivo led to durable tumor inhibition during the entire 5-week treatment period (Fig. 5D). The doxycycline-free tumors started to regrow soon after trametinib therapy was stopped, which was consistent with the presence of DTP cells in vivo. In comparison, the doxycycline-induced PINK1 knockdown tumors maintained a decreased regrowth capacity and significantly smaller tumors after trametinib cessation (Fig. 5D), consistent with the impaired DTP state after PINK1 depletion. Activation of PINK1-mediated mitophagy following trametinib treatment occurred in vivo, manifested by upregulated PINK1 levels (Fig. 5E and F) and increased serine 65-phosphorylated parkin (Fig. 5E and G). The decreased regrowth capacity after withdrawal in PINK1-depleted tumors was also evident at the level of tumor cell proliferation (measured by Ki67 staining; Fig. 5E and H). We also confirmed that PINK1 depletion disrupted redox homeostasis and resulted in apoptosis in vivo, as indicated by NADP/NADPH (Fig. 5I) and terminal-deoxynucleoitidyl transferase-mediated deoxyuridine triphosphate nick and labeling (TUNEL), respectively (Fig. 5E and J). Collectively, our in vitro and in vivo studies suggest that PINK1 facilitates the drug tolerance and therapeutic inefficacy of MAPK pathway inhibitors.

Figure 5.

PINK1 contributes to the therapeutic inefficacy of MAPK inhibitors and tumor recurrence. A, IC50 of indicated MAPK inhibitors in the top 10 LUAD cells with high PINK1 expression compared with that in the bottom 10 LUAD cells with low PINK1 expression. The data was obtained from the GDSC database. B, Colony formation of cancer cells introduced with shRNA targeting scramble or PINK1 under treatment with indicated agents. Ctrl, vehicle control; Tra, trametinib; 0325901, PD0325901; Dabra, dabrafenib; Vemura, vemurafenib; 1620, ARS1620. C, Apoptosis was measured in cancer cells introduced with shRNA targeting scramble or PINK1, in response to indicated treatments for 72 hours. D, Tumor growth monitoring in mice bearing doxycycline-induced PINK1 knockdown A549 cells xenograft tumors, under indicated treatments. Detailed treatment procedure is shown in Supplementary Fig. S10D. EJ, The xenograft tumors from D at the endpoint were detected for PINK1, phospho-parkin (serine 65), Ki67, TUNEL, and NADP/NADPH. Scale bar, 100 μm. K, Overall survival (OS) and recurrence-free survival (RFS) of LUAD patients with either low or high PINK1 levels. L, Representative IHC staining of PINK1 in LUAD tissues from patients with or without recurrence. Scale bars, 100 μm. M, Percentage of recurrence events in LUAD patients with indicated PINK1 levels. N, Progression-free survival (PFS) of melanoma patients who received MAPK inhibitor plus anti–PD-L1 therapy, stratified to high and low groups based on PINK1 expression. Data derived from GSE158403. Data are presented as the mean ± SEM. n = 3 for B and C; n = 6 for F, G, H, I, and J. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test for B, C, F, G, H, I, and J, log-rank test for K and N, or Χ2 test for M.

Figure 5.

PINK1 contributes to the therapeutic inefficacy of MAPK inhibitors and tumor recurrence. A, IC50 of indicated MAPK inhibitors in the top 10 LUAD cells with high PINK1 expression compared with that in the bottom 10 LUAD cells with low PINK1 expression. The data was obtained from the GDSC database. B, Colony formation of cancer cells introduced with shRNA targeting scramble or PINK1 under treatment with indicated agents. Ctrl, vehicle control; Tra, trametinib; 0325901, PD0325901; Dabra, dabrafenib; Vemura, vemurafenib; 1620, ARS1620. C, Apoptosis was measured in cancer cells introduced with shRNA targeting scramble or PINK1, in response to indicated treatments for 72 hours. D, Tumor growth monitoring in mice bearing doxycycline-induced PINK1 knockdown A549 cells xenograft tumors, under indicated treatments. Detailed treatment procedure is shown in Supplementary Fig. S10D. EJ, The xenograft tumors from D at the endpoint were detected for PINK1, phospho-parkin (serine 65), Ki67, TUNEL, and NADP/NADPH. Scale bar, 100 μm. K, Overall survival (OS) and recurrence-free survival (RFS) of LUAD patients with either low or high PINK1 levels. L, Representative IHC staining of PINK1 in LUAD tissues from patients with or without recurrence. Scale bars, 100 μm. M, Percentage of recurrence events in LUAD patients with indicated PINK1 levels. N, Progression-free survival (PFS) of melanoma patients who received MAPK inhibitor plus anti–PD-L1 therapy, stratified to high and low groups based on PINK1 expression. Data derived from GSE158403. Data are presented as the mean ± SEM. n = 3 for B and C; n = 6 for F, G, H, I, and J. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test for B, C, F, G, H, I, and J, log-rank test for K and N, or Χ2 test for M.

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We further examined the clinical relevance of PINK1 expression in patients with LUAD. The clinical features of LUAD patients are presented in Supplementary Table S4. These patients were classified into two groups according to PINK1 levels in the tumor tissues (Supplementary Fig. S11A). Patients with high PINK1 levels exhibited significantly poor overall and recurrence-free survival (Fig. 5K), as validated in two additional data sets (GSE31210 and GSE10245; Supplementary Fig. S11B and S11C). We also found that high PINK1 levels were associated with increased recurrence rates (Fig. 5L and M). In addition, high expression of PINK1 was associated with poor progression-free survival of melanoma patients who had received MAPK inhibitors plus anti–PD-L1 therapy (Fig. 5N; ref. 36). Taken together, these data imply that elevated PINK1 expression correlates with poor prognosis and tumor recurrence.

Pharmacologic mitophagy inhibition synergizes with MAPK pathway inhibition therapy to eradicate tumor growth and to prevent recurrence

These findings suggest that PINK1-mediated mitophagy activation could be an “Achilles heel” for persister cells. Targeting PINK1-mediated mitophagy may enhance the efficacy of MAPK targeting therapy and lead to prolonged treatment responses. Currently, there is a lack of specific mitophagy inhibitors. And the most frequently used methods for mitophagy inhibition are autophagy inhibitors, including chloroquine and 3-MA (37). Thus, we investigated whether these two mitophagy inhibition agents enhanced the efficacy of MAPK inhibitors in LUAD cell lines as well as in PDOs. We observed synergistic antiproliferative effects of mitophagy inhibitors in combination with various MAPK inhibition therapies (Fig. 6AC; Supplementary Fig. S12A–S12C). The combination of mitophagy inhibition and MAPK pathway inhibition also resulted in increased cell death compared with either of the individual agents (Fig. 6D; Supplementary Fig. S12D). In vivo, the combination of chloroquine and trametinib resulted in complete responses and cleared tumors that did not recur following drug removal (Fig. 6E and F). In addition, we assessed changes in tumor volume from dosing initiation to discontinuation in terms of RECIST. Most mice that received vehicle therapy developed PD. Although trametinib monotherapy resulted in PD or SD, mice treated with the combination of chloroquine and trametinib showed either PR or CR (Fig. 6G and H). The combination of chloroquine and trametinib induced no toxicity in treated mice (Supplementary Fig. S12E). Together, our in vitro and in vivo findings demonstrate that mitophagy inhibition is a promising strategy for eradicating MAPK inhibitor tolerance.

Figure 6.

Mitophagy inhibition synergizes with MAPK pathway inhibition to eradicate tumor growth and to prevent recurrence. AC, LUAD cell lines and LUAD PDOs were treated for 72 hours with indicated agents and then analyzed for cell viability. Synergy graphs were generated utilizing Combenefit Software. CQ, chloroquine; Tra, trametinib; Dabra, dabrafenib; Tra/dabra, trametinib plus dabrafenib. D, LUAD cell lines and LUAD PDOs were treated for 72 hours with indicated vehicle control or chloroquine (50 μmol/L) in combination with either trametinib (100 nmol/L), dabrafenib (10 μmol/L), or trametinib plus dabrafenib, and then analyzed for apoptosis. E and F, A549-derived (E) and H460-derived (F) xenograft nude mice received vehicle, chloroquine (50 mg/kg), trametinib (2 ppm), or the combination of chloroquine and trametinib. The treatment started at day 12 and stopped at day 42. Com, combination. G and H, Waterfall plot showing the response of A549 xenografts (G) and H460 xenografts (H) to indicated treatments. Percentage change of tumor volume was calculated by dividing the change of tumor volume during the entire treatment period by that at the beginning of treatment and was used to evaluate the responses to indicated therapy based on RECIST criteria. Data, mean ± SEM in D. n = 3 for D; n = 6 for E and F. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test.

Figure 6.

Mitophagy inhibition synergizes with MAPK pathway inhibition to eradicate tumor growth and to prevent recurrence. AC, LUAD cell lines and LUAD PDOs were treated for 72 hours with indicated agents and then analyzed for cell viability. Synergy graphs were generated utilizing Combenefit Software. CQ, chloroquine; Tra, trametinib; Dabra, dabrafenib; Tra/dabra, trametinib plus dabrafenib. D, LUAD cell lines and LUAD PDOs were treated for 72 hours with indicated vehicle control or chloroquine (50 μmol/L) in combination with either trametinib (100 nmol/L), dabrafenib (10 μmol/L), or trametinib plus dabrafenib, and then analyzed for apoptosis. E and F, A549-derived (E) and H460-derived (F) xenograft nude mice received vehicle, chloroquine (50 mg/kg), trametinib (2 ppm), or the combination of chloroquine and trametinib. The treatment started at day 12 and stopped at day 42. Com, combination. G and H, Waterfall plot showing the response of A549 xenografts (G) and H460 xenografts (H) to indicated treatments. Percentage change of tumor volume was calculated by dividing the change of tumor volume during the entire treatment period by that at the beginning of treatment and was used to evaluate the responses to indicated therapy based on RECIST criteria. Data, mean ± SEM in D. n = 3 for D; n = 6 for E and F. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001, as determined by the t test.

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Oncogene-targeted therapies are the effective care for cancer treatment. However, these therapies are rarely curative. Many patients initially achieve dramatic responses to targeted therapy, but still retain minimal residual disease (MRD), which eventually leads to relapse. These observations suggest that residual tumor cells persist throughout treatment. DTP state has been widely accepted to account for MRD (38). However, the mechanism by which DTP cells persist remains largely unexplored.

The emergence of the DTP state has been shown to be critically dependent on plastic changes, including altered metabolism and antioxidant programs, to escape stress (1, 24, 39). In addition, PINK1-mediated mitophagy has been shown to reduce the efficacy of chemotherapy and targeted therapy in several contexts (40, 41). However, the mechanistic basis of these observations remains unclear. Here we provide a mechanistic link between these two observations. Our studies demonstrate that PINK1-mediated mitophagy is induced following MAPK pathway inhibition. This is due to the alleviation of MYC-engaged transcriptional repression of PINK1 and as such favors a functional mitochondria network allowing the formation of a DTP state (Fig. 7).

Figure 7.

Schematic model showing the mechanism and vulnerability of DTP cells that develop from MAPK inhibitor therapy. Left, the drug-naïve tumor is fast-cycling, adopting glycolysis to meet their proliferative demands. PINK1 expression is controlled by MYC-mediated transcriptional repression. Middle, during MAPK inhibitor treatment, PINK1 expression is induced due to MYC deprivation. PINK1 mediates mitophagy activation to maintain mitochondrial homeostasis, enabling OXPHOS-dependent DTP cells to survive. Right, mitophagy inhibition through clinically approved chloroquine eradicates DTP cell generation, permitting complete responses to MAPK inhibitor therapy.

Figure 7.

Schematic model showing the mechanism and vulnerability of DTP cells that develop from MAPK inhibitor therapy. Left, the drug-naïve tumor is fast-cycling, adopting glycolysis to meet their proliferative demands. PINK1 expression is controlled by MYC-mediated transcriptional repression. Middle, during MAPK inhibitor treatment, PINK1 expression is induced due to MYC deprivation. PINK1 mediates mitophagy activation to maintain mitochondrial homeostasis, enabling OXPHOS-dependent DTP cells to survive. Right, mitophagy inhibition through clinically approved chloroquine eradicates DTP cell generation, permitting complete responses to MAPK inhibitor therapy.

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Cancer cells usually alter their metabolism to suit their proliferation (42). In various contexts, DTP cells have been shown to hijack developmental processes involved in flexible metabolic alterations to become phenotypically plastic and reversibly evolve toward drug-refractory cell identities (43). Although cancer cells generally rely on aerobic glycolysis to fuel their active proliferation, slow-cycling DTP cells exhibit increased dependence on mitochondrial respiration (44, 45). The dependence on mitochondrial respiration in DTP cells is analogous to that in resting normal cells, which also have a low demand for proliferation (46). Functional mitochondria are the foundation of mitochondrial respiration. The contribution of a functional mitochondrial network to cellular homeostasis is necessary for DTP cells to cope with cytotoxic stress. We make it clear that elevated PINK1 expression is essential for functional mitochondria and redox homeostasis in MAPK inhibitor-tolerant cells. Therefore, our observations provide PINK1-mediated mitophagy as a novel protective mechanism and prerequisite for DTP generation.

In addition, our study complements and extends prior knowledge of the biological regulation of PINK1-mediated mitophagy. As a central player in mitophagy initiation, PINK1 has been shown to be downregulated in tumors (47). Our data revealed that MYC causes transcriptional repression of PINK1. While depriving MYC protein stability, MAPK inhibition relieves the transcriptional repression of PINK1. This is consistent with the consensus that DTP is sustained by epigenetic mechanisms. MYC is a global regulator of aerobic glycolysis and biosynthesis (24, 48, 49). Aberrant MYC overexpression is common in malignant fast-growing tumors, resulting in sustained glycolysis to meet the biomass demand for rapid proliferation. In the presence of MAPK inhibitors, the MYC-dependent transcriptional induction of PINK1 establishes a switch between aerobic glycolysis (in fast-growing cancer cells) and mitochondrial respiration (in slow-cycling DTP).

Currently, there are three therapeutic strategies to target DTP: (i) sustaining tumor cells in a quiescent state, (ii) awaking DTP and sensitizing them to drugs targeting proliferative cells, and (iii) forcing quiescent cells to die and eradicating DTP (50). The first two strategies are palliative options. Both sustaining DTP cells in a slow-cycling state and awakening them to enter the active cell cycle are fraught with complications such as unpredictable tumor behavior and oncogenic evolution. Thus, targeting the regulators of DTP seems to be more rational. Our study uncovers that targeting PINK1-mediated mitophagy eradicates DTP generation. Although there is still a lack of a specific inhibitor of PINK1-mediated mitophagy, the combination of autophagy inhibitors and MAPK inhibitors generates cleared tumors that do not reoccur after withdrawal. Similarly, the synergistic efficacy of trametinib and chloroquine has been confirmed in pancreatic ductal adenocarcinoma (51). A recent study provides CDK9 inhibitors as an option for targeting PINK1-mediated mitophagy (52). Another team revealed that DTP cancer cells arising from chemotherapy are preferentially sensitive to CDK9 inhibitors (53). Whether CDK9 inhibitors synergize with MAPK pathway inhibitors needs to be evaluated in the future.

We recognize that our study has several limitations. First, neither in vitro culture models, including cell lines or organoids, nor in vivo xenograft models were able to reproduce the tumor immune microenvironment during DTP development. It will be interesting to further investigate whether the contribution of PINK1 on DTP generation is associated with tumor stroma. Second, we were not able to validate the activation of PINK1-mediated mitophagy in clinical MRD because the specimens of MRD tumors are hardly accessible. In addition, a more extensive characterization of the role of PINK1 in DTP generation in the context of other kinds of therapies and different cancer types is warranted in future work.

In summary, we propose a model where MAPK inhibitors elevate PINK1 expression and activate mitophagy by relieving MYC-mediated transcriptional repression of PINK1. PINK1-mediated mitophagy supported DTP cells' survival and contributed to poor prognosis. Targeting PINK1-mediated mitophagy eradicates DTP cancer cells and permits complete responses to MAPK inhibitors. Our work also complements and extends prior knowledge on the biological regulation of PINK1-mediated mitophagy in cancer.

Y. Li reports a patent for PINK1 as a marker of poor outcome of MAPK-targeted therapy in cancer pending. H. Chen reports grants from the National Natural Science Foundation of China and China Postdoctoral Science Foundation during the conduct of the study; in addition, H. Chen has a patent for PINK1 as a marker of poor outcome of MAPK-targeted therapy in cancer pending. D. Yin reports grants from the National Key Research and Development Program of China, the Natural Science Foundation of China, and the Guangdong Science and Technology Department during the conduct of the study; in addition, D. Yin has a patent for PINK1 as a marker of poor outcome of MAPK-targeted therapy in cancer pending. No disclosures were reported by the other authors.

Y. Li: Conceptualization, formal analysis, investigation, visualization, methodology, writing–original draft. H. Chen: Conceptualization, data curation, software, formal analysis, funding acquisition, validation, writing–original draft. X. Xie: Resources, data curation, validation, methodology. B. Yang: Software, formal analysis, visualization. X. Wang: Software, formal analysis, methodology. J. Zhang: Validation, investigation, methodology. T. Qiao: Software, formal analysis, methodology. J. Guan: Resources, investigation, methodology. Y. Qiu: Resources, formal analysis, investigation. Y.-X. Huang: Resources, formal analysis, visualization. D. Tian: Resources, formal analysis, investigation. X. Yao: Resources, formal analysis, investigation. D. Lu: Writing–review and editing. H.P. Koeffler: Supervision, writing–review and editing. Y. Zhang: Conceptualization, resources, supervision, writing–review and editing. D. Yin: Conceptualization, supervision, funding acquisition, writing–original draft, project administration, writing–review and editing.

The authors appreciate all of the members of Yin's laboratory for their important insights and helpful discussion. Mass spectrometry analysis was performed by the Bioinformatics and Omics Center, Sun Yat-Sen Memorial Hospital, Sun Yat-Sen University. This work was supported by the National Key Research and Development Program of China (2021YFA0909300), National Natural Science Foundation of China (82073067, 81872140, 81621004, 81420108026, and 82102716), China Postdoctoral Science Foundation (2022M713588), and Guangdong Science and Technology Department (2019B020226003, 2021A0505030084, 2020B1212060018, and 2020B1212030004).

The publication costs of this article were defrayed in part by the payment of publication fees. Therefore, and solely to indicate this fact, this article is hereby marked “advertisement” in accordance with 18 USC section 1734.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

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Supplementary data