Targeting PGAM1 Y119 phosphorylation resensitizes non-small-cell lung cancer to cisplatin therapy by promoting oxidative stress and DNA damage

Wenxia Zhang; Yanzhao Qu; Xin Wang; Lihua Wang; Xin Jin; Jiahui Kuang; Qian Wang; Shunxin Han; Min Wei; Yang Wang; Jia Liu

doi:10.1016/j.jbc.2026.113506

. 2026 Sep 1;302(10):113506. doi: 10.1016/j.jbc.2026.113506

Targeting PGAM1 Y119 phosphorylation resensitizes non-small-cell lung cancer to cisplatin therapy by promoting oxidative stress and DNA damage

Wenxia Zhang ^1,^‡, Yanzhao Qu ^1,^‡, Xin Wang ¹, Lihua Wang ¹, Xin Jin ¹, Jiahui Kuang ¹, Qian Wang ¹, Shunxin Han ¹, Min Wei ¹, Yang Wang ^1,^∗, Jia Liu ^1,^∗

PMCID: PMC13635264 PMID: 42680101

Abstract

Cisplatin resistance remains a primary challenge in the clinical management of non-small-cell lung cancer (NSCLC), yet the regulatory targets underlying this resistance remain largely unknown. It is well established that cisplatin kills tumor cells through the induction of DNA damage and the accumulation of reactive oxygen species, which exacerbate DNA damage. Here, we identify the glycolytic metabolic enzyme PGAM1, and specifically its elevated activity in cisplatin-resistant tumors, as a pivotal metabolic driver of this resistance. Y119 phosphorylation, which reflects increased PGAM1 activity, is significantly elevated in NSCLC patient tissues and further amplified in cisplatin-resistant cell lines. Mutation of the PGAM1 Y119 phosphorylation site (Y119F) resensitizes resistant cells to cisplatin both in vitro and in vivo. Mechanistically, Y119-phosphorylated PGAM1 enhances flux through the pentose phosphate pathway and the serine synthesis pathway. This metabolic reprogramming promotes nucleotide biosynthesis and NADPH generation, thereby alleviating cisplatin-induced DNA damage and oxidative stress. In vivo, a PGAM1-derived Y119 phosphorylation -mimetic cell-permeable peptide (Y119E-TAT) that competitively disrupts PGAM1 binding to its histidine kinase, thereby inhibiting PGAM1 activity, potently inhibits cisplatin-resistant NSCLC tumor growth. Together, these findings reveal a novel mechanism by which PGAM1 Y119 phosphorylation drives chemoresistance and suggest that targeting this phosphorylation event represents a potential therapeutic strategy to overcome cisplatin resistance in NSCLC.

Keywords: cisplatin resistance (CR), DNA damage, metabolic reprogramming, phosphoglycerate mutase 1 (PGAM1), phosphorylation, ROS

Lung cancer, the leading cause of cancer-related mortality worldwide, is broadly categorized into non-small-cell lung cancer (NSCLC) and small-cell lung cancer, accounting for over one million annual deaths globally (1, 2, 3). NSCLC represents approximately 80% of all cases and is histologically subdivided into adenocarcinoma, squamous cell carcinoma, and large cell carcinoma (4, 5). Platinum-based chemotherapy (e.g., cisplatin and carboplatin) remains a first-line treatment for advanced NSCLC (6, 7, 8, 9). It is well established that platinum drugs kill tumor cells through the formation of DNA crosslinks that trigger genotoxic stress, together with the generation of reactive oxygen species (ROS) that act synergistically to exacerbate DNA damage (10, 11, 12). However, the inevitable development of chemoresistance significantly limits the long-term therapeutic efficacy of platinum drugs.

Metabolic reprogramming has emerged as a fundamental hallmark of cancer (13, 14, 15, 16, 17). The best-characterized metabolic alteration in cancer cells is Warburg effect, also termed aerobic glycolysis, characterized by a preference for glycolysis over oxidative phosphorylation even under oxygen-rich conditions (18, 19, 20). In most tumors, upregulated glycolysis concurrently activates ancillary metabolic pathways, particularly the pentose phosphate pathway (PPP) and the serine synthesis pathway (SSP). These shunt pathways generate crucial metabolites, including ribose-5-phosphate (R5P), one-carbon units, and NADPH, which collectively support nucleotide synthesis and alleviate oxidative stress, thereby facilitating DNA replication and damage repair (21, 22, 23, 24). Notably, these pro-survival metabolic adaptations directly counteract the mechanisms of platinum drugs, suggesting that the Warburg effect may increase tumor resistance to platinum drugs.

PGAM1, a critical glycolytic enzyme frequently overexpressed and hyperactive in various malignancies including lung, bladder, and pancreatic cancers, has emerged as a promising therapeutic target (25, 26, 27, 28). Genetic or pharmacological inhibition of PGAM1 has been shown to suppress glycolytic flux and anabolic biosynthesis, ultimately impairing tumor growth. Mechanistically, PGAM1 occupies a pivotal position in cancer metabolism that its substrate 3-phosphoglycerate (3PG) competitively inhibits 6-phosphogluconate dehydrogenase (6PGD) in the PPP, while its product 2-phosphoglycerate activates 3-phosphoglycerate dehydrogenase in the SSP. The characteristic elevation of PGAM1 activity in tumors leads to decreased 3PG and increased 2-phosphoglycerate levels, thereby simultaneously reducing PPP flux while enhancing SSP flux (29, 30). Our previous work in NSCLC identified a novel regulatory mechanism of PGAM1 activity wherein pyruvate kinase M2 (PKM2) phosphorylates PGAM1 at histidine 11 (H11), activating it to enhance glycolytic shunts in cancer, with Y119 phosphorylation (pY119) being essential for PKM2 binding (31). Inhibition of PGAM1 Y119 phosphorylation reduces PGAM1 activity, thereby disrupting the coordination between glycolysis and PPP/SSP to suppress nucleotide synthesis and ROS accumulation. Therefore, increased PGAM1 activity promotes the Warburg effect; however, whether this is associated with platinum drug resistance remains unknown.

Here, we identify that Y119 phosphorylation of PGAM1, which enhances PGAM1 activity, serves as a critical metabolic checkpoint in NSCLC. By directing metabolic flux toward glycolytic shunts including PPP and SSP, this phosphorylation event confers cisplatin resistance by enhancing DNA damage repair and reducing ROS production. Importantly, in vivo administration of a PGAM1-derived pY119-mimetic cell-permeable peptide to competitively inhibit the Y119 phosphorylation-mediated increase in PGAM1 activity potently inhibits the growth of cisplatin-resistant NSCLC tumors.

Results

PGAM1 activity is elevated in NSCLC and associated with cisplatin resistance

We initially examined PGAM1 expression in paired tumor and adjacent normal tissues from 30 NSCLC patients. Immunohistochemistry (IHC) and quantitative analysis revealed that PGAM1 protein expression was significantly elevated in tumor tissues compared with adjacent normal tissues (Fig. 1A). Given our previous finding that Y119 phosphorylation is essential for PGAM1 enzymatic activity (31), we next assessed pY119 status as a surrogate for PGAM1 activity in lung cancer tissues. Using a previously validated and characterized PGAM1 pY119 antibody (31) (Fig. S1), IHC demonstrated a significant increase in PGAM1 Y119 phosphorylation in NSCLC tumor tissues relative to adjacent normal tissues (Fig. 1B). These findings indicate a potential clinical association between elevated PGAM1 expression/activity and NSCLC.

Because PGAM1 activation promotes NADPH and nucleotide biosynthesis, pathways that counteract cisplatin-induced ROS and DNA damage, we reasoned that increased Y119 phosphorylation might reduce cisplatin efficacy and thereby contribute to resistance. Therefore, we examined PGAM1 pY119 levels in three pairs of parentals and acquired cisplatin-resistant NSCLC cell lines (A549 versus A549/CR, H460 versus H460/CR, PC-9 versus PC-9/CR; CR, cisplatin resistance). To characterize the degree of cisplatin resistance, we first performed dose-response assays across a range of cisplatin concentrations (0–128 μM) and calculated GI50 values. The results showed that A549/CR, H460/CR and PC-9/CR cells exhibited resistance indices of 7.43-, 9.64-, and 15.76-fold, respectively, compared with their parental counterparts (Fig. 1, C–E), confirming substantial resistance in all three sublines. We next examined whether PGAM1 expression or its phosphorylation status correlated with the resistant phenotype. While total PGAM1 protein levels remained largely unchanged in resistant cells, Y119 phosphorylation was significantly elevated in all three resistant sublines (Fig. 1F), suggesting that PGAM1 activity, rather than protein abundance, is associated with cisplatin resistance. Functionally, upon cisplatin treatment (20 μM, 24 h), an effective concentration for inducing cytotoxicity (32, 33, 34), the A549, H460, and PC-9 parental cell lines exhibited a markedly increased apoptotic response and significantly reduced cell viability (Fig. 1, G–J). In contrast, the corresponding resistant sublines (A549/CR, H460/CR, and PC-9/CR), which display elevated PGAM1 pY119, showed minimal apoptotic response and no significant change in cell viability (Fig. 1, G–J). While cisplatin-resistant cells necessarily harbor numerous molecular alterations that collectively contribute to the resistant phenotype, the consistent elevation of pY119 across three independently derived resistant sublines suggests that increased PGAM1 activity mediated by elevated Y119 phosphorylation is functionally relevant to cisplatin resistance.

Abrogation of PGAM1 Y119 phosphorylation resensitizes cisplatin-resistant NSCLC cells

To determine whether PGAM1 Y119 phosphorylation contributes to cisplatin resistance, we generated stable A549/CR cells in which endogenous PGAM1 was depleted and then reconstituted with either WT PGAM1 (PGAM1^WT) or a non-phosphorylatable mutant (PGAM1^Y119F). Immunoblot analysis confirmed comparable expression levels of PGAM1^WT and PGAM1^Y119F in the rescued cell lines (Fig. 2A). Upon exposure to cisplatin (20 μM, 24 h), PGAM1-knockdown A549/CR cells exhibited significantly increased apoptosis compared with A549/CR cells, as determined by flow cytometry following Annexin V/PI staining (Fig. 2B). Rescue with PGAM1^WT largely restored resistance to the level of non-targeting control cells, whereas rescue with PGAM1^Y119F failed to do so, resulting in apoptosis levels comparable to those of PGAM1-knockdown cells (Fig. 2B). Consistently, Cell Counting Kit-8 (CCK-8) cell viability assays demonstrated that PGAM1 knockdown or Y119F mutation significantly enhanced cisplatin-induced cytotoxicity, reducing the viability of resistant cells to a level comparable to that of cisplatin-sensitive parental cells (Fig. 2C). These results indicate that mutation of Y119 effectively reverses the apoptotic resistance of A549/CR cells, thereby restoring their sensitivity to cisplatin. To further validate these observations, we performed siRNA-mediated knockdown of endogenous PGAM1 in H460/CR and PC-9/CR cells, followed by rescue with either Flag-tagged PGAM1^WT or PGAM1^Y119F via transient transfection. Immunoblot analysis confirmed efficient knockdown and comparable reconstitution of WT and mutant PGAM1 in both cell lines (Fig. 2, D and E). Notably, similar to the observations in A549/CR cells, upon cisplatin treatment, both H460/CR and PC-9/CR cells with PGAM1 knockdown or rescued with PGAM1^Y119F displayed significantly higher apoptosis rates (Fig. 2, F and G) and reduced cell viability (Fig. 2, H and I) compared with those rescued with PGAM1^WT. These results confirm that loss of Y119 phosphorylation, which reduces PGAM1 activity, sensitizes resistant cells to cisplatin.

To determine whether increased PGAM1 activity is sufficient to confer cisplatin resistance, we overexpressed PGAM1^WT and the phosphomimetic mutant PGAM1^Y119E in cisplatin-sensitive A549 parental cells. Both significantly reduced cisplatin-induced apoptosis and maintained cell viability compared with vector controls (Fig. S2, A and B). These gain-of-function data, together with the loss-of-function results above, establish that PGAM1 activity, specifically through Y119 phosphorylation, is both necessary and sufficient to contribute cisplatin resistance in NSCLC cells.

PGAM1 Y119 phosphorylation correlates with suppression of cisplatin-induced oxidative stress and DNA damage

It is well established that cisplatin exerts its antitumor effects primarily through the induction of genotoxic stress via DNA crosslink formation and the generation of ROS, which synergistically amplify DNA damage and trigger apoptotic cell death (10, 11, 12). To investigate whether PGAM1 Y119 phosphorylation contributes to cisplatin resistance by mitigating these cytotoxic effects, we measured intracellular ROS levels and DNA damage markers in A549/CR cells expressing either PGAM1^WT or PGAM1^Y119F following cisplatin treatment. We employed flow cytometric analysis using the ROS-sensitive probe DCFH-DA and found that PGAM1^Y119F-expressing cells accumulated significantly higher levels of ROS upon cisplatin exposure compared with PGAM1^WT-expressing cells, which maintained low ROS levels similar to those in untreated cells (Fig. 3A). These results indicate that loss of Y119 phosphorylation correlates with elevated ROS levels. We next assessed DNA damage by immunostaining for γH2AX, a well-established marker of DNA double-strand breaks. Following cisplatin treatment, PGAM1^Y119F-expressing A549/CR cells exhibited a substantial increase in the number of γH2AX foci per nucleus compared with PGAM1^WT-expressing cells (Fig. 3B). Quantitative flow cytometric analysis of γH2AX fluorescence intensity confirmed these findings, showing a marked elevation in DNA damage specifically in cells lacking Y119 phosphorylation (Fig. 3C). Together, these data indicate that loss of Y119 phosphorylation is associated with increased cisplatin-induced ROS accumulation and DNA damage.

**PGAM1 Y119 Phosphorylation Suppresses Cisplatin-Induced Oxidative Stress and DNA Damage.**A, assay for the ROS levels in PGAM1^WT or PGAM1^Y119F cell lines upon cisplatin treatment. PGAM1^WT and PGAM1^Y119F rescue A549/CR cells were treated with or without cisplatin (20 μM) for 24 h. ROS levels were assessed by flow cytometry. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗∗∗p < 0.0001; ns, not significant. B and C, assay for DNA damage in in PGAM1^WT or PGAM1^Y119F cell lines upon cisplatin treatment. PGAM1^WT and PGAM1^Y119F rescue A549/CR cells were treated with or without cisplatin (20 μM) for 24 h. Cells were pulse-labeled with γH2AX and analyzed for DNA damage by immunofluorescence (B) or flow cytometry (C). Representative images (*left*) and associated quantifications (*right*) were displayed. Five random fields from each of the three experiments. Mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗∗∗p < 0.0001; ns, not significant. D–I, assay for the ROS levels and DNA damage in PGAM1^WT or PGAM1^Y119F cell lines upon cisplatin treatment. PGAM1^WT and PGAM1^Y119F rescue H460/CR (D, F and H) or PC-9/CR (E, G and I) cells were treated with or without cisplatin (20 μM) for 24 h. ROS levels were assessed by flow cytometry (D and E). Cells were pulse-labeled with γH2AX and analyzed for DNA damage by immunofluorescence (F and G) or flow cytometry (H and I). Representative images (*left*) and associated quantifications (*right*) were displayed. Five random fields from each of the three experiments. Mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗∗∗p < 0.0001; ns, not significant. ROS, reactive oxygen species.

To further confirm the generality of this observation, we performed similar analyses in H460/CR and PC-9/CR cells following siRNA-mediated knockdown of endogenous PGAM1 and rescue with either PGAM1^WT or PGAM1^Y119F. In both cell lines, expression of PGAM1^Y119F resulted in significantly increased ROS accumulation (Fig. 3, D and E) and enhanced γH2AX signal, as detected by both immunofluorescence (Fig. 3, F and G) and flow cytometry (Fig. 3, H and I), compared with PGAM1^WT controls. These results extend the correlation between Y119 phosphorylation and suppression of oxidative stress and DNA damage to multiple cisplatin-resistant NSCLC cell lines.

Y119 phosphorylation modulates PGAM1-dependent metabolic flux to confer cisplatin resistance in NSCLC

To investigate whether Y119 phosphorylation alters PGAM1-dependent metabolic flux to support cisplatin resistance, we performed comprehensive metabolomic profiling comparing A549 parental cells and A549/CR cells using UPLC-MS/MS. Compared with A549 cells, A549/CR cells exhibited significantly increased metabolic flux through the PPP, as evidenced by elevated levels of R5P, erythrose-4-phosphate, and sedoheptulose-7-phosphate (Fig. 4A). Similarly, intermediates in the SSP, including 3PG, phosphoserine (p-Ser), serine, and glycine, were markedly enriched in A549/CR cells (Fig. 4B). To determine whether this metabolic adaptation depends on Y119 phosphorylation, we compared the metabolomic profiles of A549/CR cells expressing PGAM1^WT versus PGAM1^Y119F. Expression of the Y119F mutant reversed the metabolic alterations observed in resistant cells. Levels of PPP intermediates (R5P and erythrose-4-phosphate) and SSP metabolites (serine, glycine, and downstream one-carbon units) were significantly reduced in PGAM1^Y119F-expressing cells compared with PGAM1^WT controls (Fig. 4, C and D), indicating that loss of Y119 phosphorylation reduces both PPP and SSP flux.

**Y119 Phosphorylation Modulates PGAM1-Dependent Metabolic Flux.**A and B, metabolomics analysis of cellular metabolic profiles in human lung cancer cell lines and the CR sublines. A549 and A549/CR cells were subjected to metabolomics analysis by UPLC-MS/MS. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001. C and D, metabolomics analysis of cellular metabolic profiles in PGAM1^WT or PGAM1^Y119F cell lines. PGAM1^WT or PGAM1^Y119F rescued A549/CR cells were subjected to metabolomics analysis by UPLC-MS/MS. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001. A.U., arbitrary units. E and F, assay for NADPH/NADP⁺ levels. NADPH/NADP⁺ levels were detected with the Enhanced NADP⁺/NADPH Assay Kit with WST-8 in A549 and A549/CR cells (E) or PGAM1^WT or PGAM1^Y119F rescued A549/CR cells (F). Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗p < 0.05; ∗∗∗p < 0.001. UPLC-MS/MS, ultra-high performance liquid chromatography-tandem mass spectrometry.

Given that the PPP is a major source of NADPH, we next examined NADPH levels. A549/CR cells exhibited increased NADPH levels compared with parental A549 cells (Fig. 4E), whereas PGAM1^Y119F expression substantially reduced NADPH levels relative to PGAM1^WT (Fig. 4F). These results suggest that Y119 phosphorylation promotes NADPH production through enhanced PPP flux. To determine whether Y119 phosphorylation mediates ROS scavenging specifically through this metabolic pathway rather than through activation of other ROS-scavenging systems, we treated PGAM1^WT- and PGAM1^Y119F-rescued A549/CR, H460/CR, and PC-9/CR cells with 6PGD-IN-1, a specific inhibitor of 6PGD, the rate-limiting enzyme of the oxidative PPP, in the presence of cisplatin. In PGAM1^WT cells, 6PGD-IN-1 treatment significantly reduced NADPH levels (Fig. 5, A–C), elevated ROS accumulation (Fig. 5, D–F), increased apoptosis (Fig. 5, G–I), and decreased cell viability (Fig. 5, J–L), phenocopying the effects of the Y119F mutation. Importantly, in PGAM1^Y119F cells, 6PGD-IN-1 did not further exacerbate these phenotypes (Fig. 5, A–L), indicating that Y119 phosphorylation and 6PGD function within the same linear pathway. These results demonstrate that Y119 phosphorylation supports ROS detoxification through enhanced PPP flux and NADPH production, establishing a direct mechanistic link between Y119 phosphorylation of PGAM1 and suppression of cisplatin-induced oxidative stress.

**PGAM1 Y119 phosphorylation enhances ROS detoxification through the PPP-NADPH metabolic axis.**A–C, assay for NADPH/NADP⁺ levels. A459/CR cells (PGAM1 KD rescued with PGAM1^WT or PGAM1^Y119F) and H460/CR and PC-9/CR cells (siPGAM1 rescued with Flag-tagged PGAM1^WT or PGAM1^Y119F) were treated with or without cisplatin (20 μM) or 6PGD-IN-1 (20 μM) for 24 h. NADPH/NADP⁺ levels were detected with the Enhanced NADP⁺/NADPH Assay Kit with WST-8. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001; ns, not significant. A.U., arbitrary units. D–F, assay for the ROS levels. A459/CR cells (PGAM1 KD rescued with PGAM1^WT or PGAM1^Y119F) and H460/CR and PC-9/CR cells (siPGAM1 rescued with Flag-tagged PGAM1^WT or PGAM1^Y119F) were treated with or without cisplatin (20 μM) or 6PGD-IN-1 (20 μM) for 24 h. ROS levels were assessed by flow cytometry. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗p < 0.01; ∗∗∗∗p < 0.0001; ns, not significant. G–I, assay for the apoptosis rate. A459/CR cells (PGAM1 KD rescued with PGAM1^WT or PGAM1^Y119F) and H460/CR and PC-9/CR cells (siPGAM1 rescued with Flag-tagged PGAM1^WT or PGAM1^Y119F) were treated with or without cisplatin (20 μM) or 6PGD-IN-1 (20 μM) for 24 h. Cell apoptosis rate (PI and AnnexinⅤ staining) were assessed by flow cytometry. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001; ns, not significant. J–L, analysis of cell viability. A459/CR cells (PGAM1 KD rescued with PGAM1^WT or PGAM1^Y119F) and H460/CR and PC-9/CR cells (siPGAM1 rescued with Flag-tagged PGAM1^WT or PGAM1^Y119F) were treated with or without cisplatin (20 μM) or 6PGD-IN-1 (20 μM) for 24 h. Cell viability was assessed using the CCK-8 Assay Kit. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001; CCK-8, Cell Counting Kit-8; ns, not significant; ROS, reactive oxygen species.

In parallel, R5P derived from the PPP and serine from the SSP provide one-carbon units essential for nucleotide biosynthesis. Accordingly, metabolites involved in nucleotide biosynthesis, including AMP, adenosine, uridine monophosphate, uridine, orotic acid and guanosine, were significantly elevated in A549/CR cells compared with parental controls (Fig. 6A), and these increases were reversed by the Y119F mutation (Fig. 6B). This suggests that Y119 phosphorylation may enhance DNA damage repair capacity in cisplatin-resistant cells by promoting nucleotide production. Moreover, enhanced nucleotide synthesis not only facilitates DNA damage repair but also promotes DNA replication during rapid proliferation of cisplatin-resistant cells. To test whether enhanced nucleotide synthesis supports DNA replication, we performed EdU incorporation assays. Cisplatin-resistant A549/CR, H460/CR, and PC-9/CR cells all exhibited significantly higher rates of EdU incorporation compared with their parental counterparts (Fig. 6, C–E). Importantly, expression of PGAM1^Y119F in each of these resistant cell lines markedly reduced EdU incorporation to levels comparable to those of sensitive parental cells (Fig. 6, F–H), demonstrating that Y119 phosphorylation supports DNA synthesis in the resistant context. Collectively, these results establish that Y119 phosphorylation modulates PGAM1-dependent metabolic flux to promote both NADPH production and nucleotide biosynthesis. This dual metabolic reprogramming simultaneously alleviates cisplatin-induced oxidative stress and supports DNA repair and replication, thereby contributing to cisplatin resistance and rapid proliferation in NSCLC cells.

**Y119 Phosphorylation Modulates PGAM1-Dependent Metabolic Flux to Confer Cisplatin Res**istance in NSCLC.A and B, metabolomic analysis of cellular metabolic profiles in human lung cancer cell lines and the CR sublines. A549 and A549/CR cells (A) or PGAM1^WT or PGAM1^Y119F rescued A549/CR cells (B) were subjected to metabolomics analysis by UPLC-MS/MS. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA: ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001; ns, not significant. C–E, assay for DNA replication in human lung cancer cell lines and the CR sublines. A549 and A549/CR (C), H460 and H460/CR (D), PC-9 and PC-9/CR (E) cells were pulse-labeled with EdU and analyzed for DNA synthesis by immunofluorescence. Representative images (*left*) and associated quantifications (*right*) were displayed. The scale bar represents, 25 μm. Five random fields from each of the three experiments. Mean ± SD, n = 3. Student’s t test, ∗∗∗∗p < 0.0001. F–H, assay for DNA replication in PGAM1^WT or PGAM1^Y119F cell lines. PGAM1^WT or PGAM1^Y119F rescued A549/CR (F), H460/CR (G) and PC-9/CR (H) cells were pulse-labeled with EdU and analyzed for DNA synthesis by immunofluorescence. Representative images (*left*) and associated quantifications (*right*) were displayed. The scale bar represents, 25 μm. Five random fields from each of the three experiments. Mean ± SD, n = 3. Student’s t test, ∗∗∗∗p < 0.0001. UPLC-MS/MS, ultra-high performance liquid chromatography-tandem mass spectrometry.

A PGAM1 pY119-mimetic cell-permeable peptide overcomes cisplatin resistance in vivo

It is known that PGAM1 Y119 phosphorylation facilitates binding to PKM2, which in turn phosphorylates H11 to generate the active form of PGAM1. To further validate the relationship between Y119 phosphorylation and cisplatin resistance, we designed a cell-permeable therapeutic peptide designated Y119E-TAT. This peptide consists of a PGAM1-derived sequence harboring a Y119E mutation, a common approach that uses the acidic residue glutamate to mimic phosphorylation (35, 36, 37), fused to the HIV-TAT cell-penetrating domain (Fig. 7A). Theoretically, upon treatment with the Y119E-TAT peptide, it competitively disrupts the binding of PGAM1 to its histidine kinase PKM2, thereby inhibiting PGAM1 activation. Our results show that the Y119E-TAT peptide, but not the TAT control, disrupted the interaction between PGAM1 and PKM2 in A549/CR cells. This disruption led to decreased phosphorylation at PGAM1 H11, which is the active site of PGAM1 (Fig. 7B). These results indicate that Y119E-TAT indeed inhibits intracellular PGAM1 activity.

To evaluate the therapeutic potential of Y119E-TAT in vivo, we established subcutaneous xenograft tumors in athymic nude mice using A549/CR cells (Fig. 7C). Beginning 10 days after tumor implantation, mice received intraperitoneal injections of either control TAT or Y119E-TAT peptide (10 mg/kg) every other day. After a 10-days treatment period, when tumors in the control TAT group had reached approximately 500 mm³, the tumors were harvested. Both tumor volume and mass were significantly reduced in mice treated with Y119E-TAT compared with those receiving the TAT peptide (Fig. 7, D–G). To confirm target engagement in vivo, we performed immunohistochemical analysis of PGAM1 Y119 phosphorylation in tumor sections. Tumors from Y119E-TAT-treated mice exhibited significantly reduced pY119 staining compared with TAT-treated controls (Fig. 7, G and I), confirming that the peptide effectively inhibits PGAM1 Y119 phosphorylation in the tumor microenvironment. This was accompanied by decreased Ki-67 staining (Fig. 7, G and H), indicating reduced tumor cell proliferation. These results demonstrate that Y119E-TAT peptide effectively engages its target in vivo and suppresses tumor growth. These results demonstrate that targeted disruption of PGAM1 Y119 phosphorylation-mediated function using the Y119E-TAT peptide effectively inhibits the growth of cisplatin-resistant NSCLC tumors in vivo, suggesting that PGAM1 activity, as reflected by upregulated Y119 phosphorylation, contributes to tumor cell cisplatin resistance.

Discussion

The involvement of glycolytic shunts in maintaining redox homeostasis and supporting DNA synthesis and repair processes suggests that Warburg effect may increase tumor resistance to cisplatin (10, 11, 18, 19, 20). While PGAM1 uniquely integrates glycolysis with its metabolic shunts (25, 26, 27, 28, 29, 30), whether it contributes to cisplatin resistance has yet to be determined. Here, we identify that Y119 phosphorylation, which enhances PGAM1 activity, acts as a critical metabolic checkpoint in NSCLC. By directing metabolic flux toward glycolytic shunts and nucleotide synthesis, it confers cisplatin resistance through reducing ROS production and enhancing DNA damage repair. In vivo, administration of a PGAM1-derived pY119-mimetic cell-permeable peptide to competitively inhibit the Y119 phosphorylation-mediated increase in PGAM1 activity potently inhibits the growth of cisplatin-resistant NSCLC tumors.

PGAM1, a pivotal glycolytic node enzyme, bridges glycolysis and biosynthetic shunts in cancer, playing a dual role in redox homeostasis and DNA damage repair (25, 26, 27, 28, 29, 30). In platinum-resistant NSCLC, elevated PGAM1 activity suppresses ROS accumulation and mitigates cisplatin-induced oxidative DNA damage, reinforcing its role as a key regulator of treatment-refractory malignancies. Beyond its metabolic functions, PGAM1 exerts non-enzymatic roles in cancer progression. For example, its enzymatic activity stabilizes CtIP to promote homologous recombination-mediated DNA repair (38, 39, 40), while its enzymatically inactive form enhances metastasis by interacting with actin alpha 2 to regulate actin dynamics and cell migration (28). A novel allosteric inhibitor, HKB99, targets PGAM1 by disrupting both its catalytic conformational changes and its interaction with actin alpha 2, thereby suppressing tumor growth, metastasis, and overcomes erlotinib resistance in NSCLC by amplifying oxidative stress and modulating key signaling pathways (41, 42). Together, these findings establish PGAM1 as a central metabolic orchestrator in chemoresistance and highlight its therapeutic potential in aggressive cancers.

Our previous work demonstrated that PKM2-mediated phosphorylation of PGAM1 at H11 drives glycolytic flux and promotes tumor progression. This process depends on priming phosphorylation at Y119, which facilitates PKM2 binding and subsequent H11 activation (31). Here, we extend these findings by showing that Y119 phosphorylation orchestrates a metabolic reprogramming that directly fuels cisplatin resistance in NSCLC. Specifically, in cisplatin-resistant cells, elevated Y119 phosphorylation enhances flux through PPP and SSP. This metabolic reprogramming boosts NADPH production and nucleotide biosynthesis, as evidenced by increased levels of PPP/SSP intermediates and nucleotide precursors. The resulting metabolic state simultaneously alleviates cisplatin-induced oxidative stress and promotes DNA damage repair. Consequently, resistant cells survive cisplatin treatment, whereas genetic (Y119F mutation) or pharmacological (Y119E-blocking peptide) inhibition of Y119 phosphorylation restores chemosensitivity both in vitro and in vivo. Collectively, these findings position PGAM1 Y119 phosphorylation as a critical metabolic checkpoint that reprograms cellular metabolism to confer cisplatin resistance. Targeting this phosphorylation event not only sensitizes treatment-naïve NSCLC cells to cisplatin but also overcomes acquired resistance in refractory cells.

Previous studies have established PGAM1 as a promising therapeutic target in oncology, driving intensive research into the development of PGAM1 inhibitors (26, 41, 43, 44). MJE3 was identified through in situ proteome reactivity profiling, though its moderate PGAM1 inhibition and weak anti-proliferative effects precluded further therapeutic exploration (44). Similarly, while EGCG exhibited potent enzymatic inhibition, its limited cellular efficacy and promiscuous targeting of multiple tumorigenic pathways undermined its selectivity (45). Subsequent work expanded PGMI-004A, HKB99 and 12r as PGAM1 inhibitors, yet only HKB99 demonstrated robust suppression of tumor growth and metastasis in vivo (41, 42, 46). Parallel efforts have advanced peptide-based strategies, exemplified by PD-L1-blocking peptides for immune checkpoint modulation and dimeric 123B9 peptide targeting EphA2, which synergized with paclitaxel to inhibit breast cancer lung metastasis (47). Our group previously contributed synthetic PGAM1 pY119 peptides that antagonize PGAM1-PKM2 binding and impede tumor progression, but their clinical potential was constrained by rapid phosphotyrosine dephosphorylation (31). To address this, in the present study, we engineered a Y119E phosphomimetic peptide, replacing tyrosine with glutamate to replicate phosphorylation-dependent steric and charge properties while achieving exceptional stability. This innovation enabled sustained PGAM1-PKM2 disruption and significant tumor suppression in NSCLC xenografts, prompting future studies to evaluate Y119E combinations with immune checkpoint inhibitors for targeting metabolic-immune crosstalk in refractory cancers.

In summary, this study demonstrates that increased PGAM1 activity enhances ROS detoxification and DNA damage repair during rapid proliferation of NSCLC cells. We further establish that PGAM1-driven metabolic reprogramming, mediated by Y119 phosphorylation, constitutes a central axis of cisplatin resistance. Therefore, targeting PGAM1 activity via disrupting Y119 phosphorylation represents a promising therapeutic strategy to overcome cisplatin resistance in NSCLC.

Experimental procedures

Cell culture

Human A549 lung cancer cells and cisplatin-resistant A549/CR cells were cultivated in Ham's F-12 medium (Sigma), supplemented with 10% fetal bovine serum (FBS, Procell) and 1% penicillin/streptomycin (Sigma). Human H460, H460/CR, PC-9 and PC-9/CR lung cancer cells were cultivated in RPMI 1640 medium (Sigma), supplemented with 10% FBS, Procell and 1% penicillin/streptomycin (Sigma). Human embryonic kidney 293T (HEK-293T) cells were maintained in DMEM medium (Sigma) with 10% FBS and 1% penicillin/streptomycin. All cell lines were sourced from the Cell Bank of Type Culture Collection of the Chinese Academy of Sciences and the cell lines were authenticated using short tandem repeat (STR) profiling by the suppliers. Mycoplasma contamination was not detected in any of the cell lines.

DNA constructs and mutagenesis

Human PGAM1 cDNA was amplified via PCR and cloned into p3×Flag-CMV-10 or pCDH-CMV vectors. The PGAM1 Y119F mutation was introduced using the QuickChange Site-directed Mutagenesis Kit (Takara). To create a PGAM1-depleted cell line, pLVTHM PGAM1 shRNA was constructed with the oligonucleotide CGCCTCAATGAGCGGCACTAT targeting the PGAM1 transcript. The control pLVTHM was generated with the oligonucleotide GCTTCTAACACCGGAGGTCTT. PGAM1 siRNA was constructed with the oligonucleotide CAAGAACTTGAAGCCTATCAA and CGCCTCAATGAGCGGCACTAT targeting the PGAM1 transcript.

Stable cell line generation

A PGAM1-depleted cell line was established through lentiviral transduction in A549 cells. Lentivirus particles were produced in HEK-293T cells by cotransfecting psPAX and Pmd2.G plasmids along with pLVTHM-shPGAM1. The virus supernatant, supplemented with polybrene (Sigma), was used to infect A549 cells. Infected cells were selected with medium containing puromycin (2 μg/ml, Sigma) for 1 week before testing for expression. For PGAM1 rescue cell lines, WT or mutant pCDH-PGAM1 plasmids were cotransfected with psPAX and Pmd2.G into HEK-293T cells to prepare lentiviruses. Cells depleted for PGAM1 were infected with these lentiviruses in the presence of polybrene and selected with puromycin (2 μg/ml, Sigma) and hygromycin (400 μg/ml, Sigma) for 2 weeks before testing for expression.

Immunoprecipitation and Western blot

Cells were lysed in cold WB-IP lysis buffer (Beyotime) for 30 min and centrifuged at 12,000g for 15 min at 4 °C to remove debris. The supernatants, serving as whole-cell lysates, were used for immunoprecipitation with indicated antibodies coupled to Protein A/G PLUS Agarose beads for 3 h at 4 °C. Nonspecific mouse IgG antibody coupled to Protein A/G PLUS Agarose beads was used as a negative control. Except for PGAM1 H11 phosphorylation assays, immunoprecipitated proteins were boiled off with SDS-PAGE loading buffer (pH 6.8) for 5 min and analyzed by Western blot using standard procedures. Primary antibodies used included anti-PGAM1 (1:200 for WB and 1:10 for IP, Santa Cruz, 130334), anti-Flag (1:1000 for WB and 1:50 for IP, Sigma, F1804), anti-Actin (1:1000 for WB, Sigma, A4700), anti-Ki-67 (1:100 for IHC, CST, 34330), and PGAM1 pY119 site-specific antibody (1:1000 for WB and 1:100 for IHC) generated by Abclonal Technology. The PGAM1 phospho-mimic peptide KIWRRSEDVP and non-phosphopeptide KIWRRSYDVP were synthesized and conjugated with KLH, respectively. Rabbit serum antibodies were produced by immunizing with the synthetic phosphopeptide KIWRRSEDVP after five immunizations. Non-phospho-specific antibodies were removed by chromatography using non-phosphopeptide KIWRRSYDVP, and the PGAM1 pY119 site-specific antibody was purified by affinity chromatography using epitope-specific KIWRRS(p)YDVP phosphopeptide. Secondary antibodies included goat anti-mouse IgG HRP-conjugated (SAB, L3032, 1:5000) and goat anti-rabbit IgG HRP-conjugated (SAB, L3012, 1:5000). Mouse IgG (Sigma, I8765, 1:50) and Rabbit IgG (Sigma, I8140, 1:50) were also used.

Peptide dot blot

Peptide dot blots were used to test the specificity of PGAM1 pY119 antibody. The peptides including phosphorylated PGAM1 Y119 peptide (KIWRRSpYDVP) and unphosphorylated PGAM1 Y119 peptide (KIWRRSYDVP) were dissolved in water to reach the concentration of 2 mg/ml. In the assay, serial dilutions (200, 100, 50, 10, 5, and 1 ng/μl) were prepared and 1 μl of each peptide was spotted on nitrocellulose. After drying for 10 min at RT, membranes were blocked with blocking buffer for 1 h at RT and incubated with PGAM1 pY119 antibody (diluted 1:1000 in blocking buffer) for 1 h at RT as described in “Western blot” in “Experimental procedures”.

Metabolomics assay

Cultured cells were rapidly washed twice with ice-cold PBS and quenched with 80% prechilled methanol (−80 °C). Cells were scraped, and the suspension was transferred to precooled tubes. Internal standards were added for quality control. Following vortexing and centrifugation (15,000g, 15 min, 4 °C), the supernatant was collected and dried under a gentle stream of nitrogen. The dried extracts were reconstituted in 50% acetonitrile for ultra-high performance liquid chromatography-high resolution mass spectrometry analysis. Metabolite separation was performed on a U3000 ultra-high performance liquid chromatography system (Thermo Fisher Scientific) coupled to a Q Exactive HF Hybrid Quadrupole-Orbitrap Mass Spectrometer. A hydrophilic interaction liquid chromatography column (e.g., Thermo Fisher Scientific Accucore hydrophilic interaction liquid chromatography column, 2.1 × 100 mm, 2.6 μm) was maintained at 40 °C. The mobile phases consisted of (A) 0.2% formic acid in water and (B) 0.2% formic acid in acetonitrile. The flow rate was 0.3 ml/min with the following gradient: 0 to 3 min, 95% B; 3 to 12 min, 95% B to 50% B; 12 to 16 min, hold at 50% B; 16 to 16.1 min, 50% B to 95% B; 16.1 to 20 min, re-equilibration at 95% B. The injection volume was 5 μl. Mass detection was carried out in both positive and negative ionization modes with a heated electrospray ionization source. Key parameters were: spray voltage, ±3.5 kV; capillary temperature, 320 °C; sheath gas, 40 arb; aux gas, 10 arb; probe heater temperature, 350 °C. Full MS scans were acquired from m/z 60 to 900 at a resolution of 70,000. Data-dependent MS/MS scans were triggered for the top 10 most intense ions per scan cycle at a resolution of 17,500, using stepped normalized collision energy (20, 40, 60). Raw data were processed using Compound Discoverer software (Thermo Fisher Scientific). This included peak detection, alignment, and integration. Metabolites were identified by matching accurate mass (mass error < 5 ppm) and MS/MS spectra against commercial databases (e.g. mzCloud).

Intracellular NADPH level assay

The intracellular NADPH level was measured using the NADP⁺/NADPH Quantification Kit (K347, Biovision). Briefly, samples and standards from the kit were heated at 60 °C for 30 min to decompose NADP⁺, then incubated with the reaction mix (NADP⁺/NADPH Extraction Buffer, NADP⁺ Cycling Buffer, NADP⁺ Cycling Enzyme Mix, NADPH Developer, and Stop Solution) in a 96-well plate. Absorbance at 450 nm was measured using a microplate reader, and NADPH concentrations were calculated based on the absorbance of the standards.

EdU Click-iT assay

Cells were seeded on glass coverslips for 24 h and treated with EdU (40 μM) for 20 min. After washing, cells were fixed in 4% paraformaldehyde for 7 min. EdU staining was performed using the Click-iT Cell Reaction Buffer Kit (Invitrogen, C10269) and Alexa Fluor 647 Azide (Invitrogen, A10277) according to the manufacturer's instructions. Cell nuclei were stained with DAPI (Sigma, D9542). Images were captured using a fluorescent confocal microscope (Carl Zeiss LSM880) and analyzed with NIS-Elements AR version 5.01 software (Nikon) (https://www.microscope.healthcare.nikon.com/en_AOM/products/software/nis-elements/nis-elements-advanced-research). Statistical analysis was conducted using Prism 8 (GraphPad software) (https://www.graphpad.com/).

Intracellular ROS production

Intracellular ROS levels were quantified using the DCFH-DA Flow Cytometry Assay Kit (Sigma, D6883). Cell-permeable reagents in the kit are non-fluorescent or weakly fluorescent in a reduced state and exhibit strong fluorescence upon oxidation by ROS. Briefly, DCFH-DA Reagent was added to cells at a final concentration of 5 μM and incubated for 30 min at 37 °C. Cells were then harvested and analyzed by flow cytometry (BD, FACSCantoII) at excitation/emission = 640/665 nm.

DNA damage assay

Cells were seeded in 6-well plates at a density of 3 × 10⁵/well and treated with cisplatin for 24 h after overnight culture. For immunofluorescence analysis, cells were transferred to a new plate on ice, washed sequentially with precooled PBS, CSK permeate, CSK-Triton, and PBS, and then fixed with 4% paraformaldehyde, blocked with 5% BSA-PBST, and incubated with primary and secondary antibodies at 4 °C and room temperature, respectively. After DAPI staining and sealing with Mowiol 488, slides were imaged using a fluorescence microscope. For flow cytometry, cells were seeded in 12-well plates at a density of 1.25 × 10⁵/well, treated with or without drugs for 48 h, and processed through trypsin digestion, washing, permeabilization, fixation, and antibody incubation. Finally, cells were resuspended in PBS for flow cytometry detection.

Cell apoptosis assay

Cells were seeded in 12-well plates at a density of 1.25 × 10⁵/well and cultured in normal complete medium for 1 day, then treated with or without drugs for 48 h. The old culture medium was retained when collecting cells, which were digested with trypsin without EDTA for about 5 min, stopped with a small amount of serum, transferred to a centrifuge tube, and centrifuged at 300g for 5 min at 4 °C. After suspension in complete culture medium, cells were cultured at 37 °C for 30 min. Subsequently, cells were washed twice with precooled PBS and centrifuged for 5 min at 300g each time. Finally, cells were resuspended in 100 μl 1 × Annexin V binding buffer, stained with 3 to 5 μl YF647-Annexin V and PI in the dark for 10 to 15 min. After staining with YF647-Annexin V and PI, cells were resuspended in 400 μl PBS and analyzed using a BD FACSCanto II flow cytometer. For each sample, at least 10,000 events were acquired. Debris and cell aggregates were excluded by gating on forward scatter (FSC) and side scatter area parameters, followed by doublet discrimination using FSC-height versus FSC-width. Compensation was performed using single-stained controls (YF647-Annexin V-only and PI-only) to correct for spectral overlap. Apoptotic cells were identified on a quadrant gate set on YF647-Annexin V (APC channel, 660/20 nm) versus PI (PE channel, 585/42 nm) dot plots: viable cells (YF647-Annexin V^-/PI^-), early apoptotic cells (YF647-Annexin V⁺/PI⁻), late apoptotic/necrotic cells (YF647-Annexin V⁺/PI⁺). The total apoptotic rate was calculated as the sum of late apoptotic cells. Data were analyzed using FlowJo 10 software.

Cell viability assay

Cell viability was assessed using the CCK-8 assay. Briefly, cells were seeded in 96-well plates and treated with the indicated compounds. Following incubation, 10 μl of CCK-8 reagent was added to each well, and the plates were further incubated at 37 °C for 1 to 4 h. The absorbance was measured at 450 nm using a microplate reader. For dose-response analysis, cells were seeded in 96-well plates at 5 × 10³ cells/well and treated with increasing concentrations of cisplatin (0–128 μM) for 48 h. Cell viability was assessed using CCK-8 assay as described above. GI50 values (concentration required for 50% growth inhibition) were calculated by nonlinear regression analysis using Prism 8 (GraphPad software). Cell viability was calculated relative to untreated control wells and presented as the mean percentage ± SD from at least three independent experiments.

Tumor xenograft analysis

For Y119E-TAT peptide functional analysis, 1 × 10⁶ A549/CR cells were suspended in 50 μl Matrigel and 100 μl PBS and injected into mice. When tumors reached ∼50 mm³ (day 10 after implantation), mice were randomly assigned to TAT control or Y119E-TAT groups (n = 9 per group) with comparable mean tumor volumes. Then, mice received intraperitoneal injections of TAT or Y119E-TAT (10 mg/kg) every other day for 10 days. Tumor volumes were measured on day 10, 15 and 20 to monitor growth kinetics. Five days after tumor formation, 10 mg/kg of TAT peptide or Y119E-TAT peptide was injected intraperitoneally every other day. Xenograft tumors were sampled and photographed after 20 days. Mice were obtained from Beijing HFK Bioscience Co, Ltd and housed in a 12-h light/dark cycle at 20 °C and 40% humidity. All animal procedures were approved by the Ethics Committee of Science and Technology of Northeast Normal University (202502057).

Immunohistochemistry analysis

Mouse tumor tissues were fixed and prepared for IHC. Paraffin-cut sections of xenograft tumors were stained with H&E and Ki-67. Three random areas of each tissue sample were examined, analyzed, and imaged (Leica DMi8) to quantify positive staining. Human lung cancer tissue sections were stained with indicated antibodies. Tissue sections were blindly scored based on the percentage of positive cells and staining intensity (0 = negative, 1 = weak, 2 = moderate, 3 = strong; proportion scores: 0=<5%, 1 = 6–25%, 2 = 26–50%, 3 = 51–75%, 4=>75%). The IHC score was calculated using the formula: 3 × score of strongly staining area + 2 × score of moderately staining area + 1 × score of weakly staining area (range 0–12). Scores were correlated with overall survival. Lung cancer tissues and matching normal tissues were obtained from the Biobank of China–Japan Union Hospital of Jilin University during surgery and stored at −80 °C. The study was conducted in accordance with the principles of the Declaration of Helsinkihe, and the protocol was approved by the Ethics Committee of Science and Technology of Northeast Normal University and conducted with informed consent from all patients (202502057).

Statistical analysis

For IHC data, Student’s t test (paired, two-tailed) was used for two-group comparisons. Pearson’s correlation test was applied to analyze the correlation significance between proteins in human lung cancer tissue data. Log-rank test was used to assess survival differences between groups. For other data, Student’s t-tests (unpaired) were performed for two groups, and one-way or two-way ANOVA followed by Tukey’s multiple comparison test was used for datasets with three or more groups. Data were analyzed using Prism Version eight software (GraphPad) and presented as mean ± SD. Significance levels were denoted as ∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001; ns, not significant.

Data availability

Data are provided within the manuscript or supporting information files. The metabolomics data have been deposited in the OMIX, China National Center for Bioinformation/Beijing Institute of Genomics, Chinese Academy of Sciences (https://ngdc.cncb.ac.cn/omix), accession number OMIX019080.

Supporting information

This article contains supporting information.

Acknowledgments

Author contributions

W. Z. and Y. Q. visualization; W. Z., Y. Q., Y. W., and J. L. project administration; W. Z., Y. Q., X. W., L. W., X. J., J. K., Q. W., S. H., and J. L. investigation; M. W. and Y. W. writing–review and editing; M. W., Y. W., and J. L. conceptualization; Y. W. supervision; Y. W. and J. L. funding acquisition; J. L. writing–original draft; J. L. validation.

Funding and additional information

This work was supported by the National Natural Science Foundation of China (32571509 and 32500685), the Science and Technology Development Plan Project of Jilin Province, China (20260101029JJ), the Postdoctoral Fellowship Program of CPSF under Grant Number GZC20251777 and the China Postdoctoral Science Foundation under Grant Number 2026M792760.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Reviewed by members of the JBC Editorial Board. Edited by Paul Shapiro

Contributor Information

Yang Wang, Email: wangy422@nenu.edu.cn.

Jia Liu, Email: liuj906@nenu.edu.cn.

Supporting information

Figs. S1 and S2

mmc1.pptx^{(535KB, pptx)}

Supplementary Material

mmc2.ppt^{(138.1MB, ppt)}

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figs. S1 and S2

mmc1.pptx^{(535KB, pptx)}

Supplementary Material

mmc2.ppt^{(138.1MB, ppt)}

Data Availability Statement

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PERMALINK

Targeting PGAM1 Y119 phosphorylation resensitizes non-small-cell lung cancer to cisplatin therapy by promoting oxidative stress and DNA damage

Wenxia Zhang

Yanzhao Qu

Xin Wang

Lihua Wang

Xin Jin

Jiahui Kuang

Qian Wang

Shunxin Han

Min Wei

Yang Wang

Jia Liu

Abstract

Results

PGAM1 activity is elevated in NSCLC and associated with cisplatin resistance

Figure 1.

Abrogation of PGAM1 Y119 phosphorylation resensitizes cisplatin-resistant NSCLC cells

Figure 2.

PGAM1 Y119 phosphorylation correlates with suppression of cisplatin-induced oxidative stress and DNA damage

Figure 3.

Y119 phosphorylation modulates PGAM1-dependent metabolic flux to confer cisplatin resistance in NSCLC

Figure 4.

Figure 5.

Figure 6.

A PGAM1 pY119-mimetic cell-permeable peptide overcomes cisplatin resistance in vivo

Figure 7.

Discussion

Experimental procedures

Cell culture

DNA constructs and mutagenesis

Stable cell line generation

Immunoprecipitation and Western blot

Peptide dot blot

Metabolomics assay

Intracellular NADPH level assay

EdU Click-iT assay

Intracellular ROS production

DNA damage assay

Cell apoptosis assay

Cell viability assay

Tumor xenograft analysis

Immunohistochemistry analysis

Statistical analysis

Data availability

Supporting information

Acknowledgments

Author contributions

Funding and additional information

Conflict of interest

Contributor Information

Supporting information

References

Associated Data

Supplementary Materials

Data Availability Statement

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