Copper homeostasis dysregulation promotes malignant transformation of renal tubular epithelial cells via the ROS/NF-κB/AID signaling axis

Introduction

Renal cell carcinoma (RCC) is a common malignancy (1), and its incidence is rising worldwide (2). Among RCC subtypes, clear cell renal cell carcinoma (ccRCC) is the most common histological subtype, accounting for approximately 70–80% of all RCC cases, and is characterized by aggressive biological behavior, high metastatic potential, and an unfavorable prognosis (3). Despite substantial advances in the management of advanced ccRCC, including immune checkpoint inhibitors (ICIs), tyrosine kinase inhibitors that target the vascular endothelial growth factor receptor, and hypoxia-inducible factor-2α inhibitors, a considerable proportion of patients eventually develop primary or acquired resistance, resulting in tumor progression and limited long-term survival benefits (4-6). Therefore, the molecular mechanisms underlying ccRCC initiation and progression, particularly in the context of chronic renal inflammation and epithelial cell transformation, urgently need to be identified to establish novel diagnostic biomarkers and therapeutic targets.

Copper is an essential trace element that plays critical roles in various biological processes, including mitochondrial respiration, angiogenesis, and redox homeostasis. Cellular copper levels are tightly regulated by copper transport proteins, including the high-affinity importer copper transporter 1 (CTR1) and copper exporters such as ATPase copper-transporting alpha/beta (ATP7A/B), as well as copper chaperones that facilitate intracellular distribution (7-9).

Recent studies have revealed that copper metabolism is dysregulated in multiple cancers, including pancreatic cancer, hepatocellular carcinoma, colorectal cancer, and lung cancer, where elevated copper levels are often associated with tumor progression and poor prognosis (10-16). Research has shown that copper regulates tumor proliferation, metastasis, angiogenesis, oxidative stress responses, and immune remodeling through multiple intracellular signaling pathways (8,17,18). In ccRCC, copper has been shown to directly drive metabolic remodeling and tumor progression (8). Emerging concepts such as cuproplasia and cuproptosis further underscore the dual role of copper in promoting tumor growth and regulating cell death pathways, highlighting copper homeostasis as a promising therapeutic target in cancer (10,17).

One of the major consequences of copper accumulation is the induction of oxidative stress. Copper ions can catalyze Fenton-like reactions, leading to excessive production of reactive oxygen species (ROS) (19-22). ROS are highly reactive molecules that play dual roles in cellular physiology and pathology. Under pathological conditions, excessive ROS accumulation induces oxidative stress, leading to DNA damage, genomic instability, and activation of oncogenic signaling pathways (23-25).

Notably, ROS act as upstream regulators of the NF-κB signaling pathway. Under oxidative stress, Inhibitor of kappa B (IκB) undergoes phosphorylation and degradation, allowing the nuclear factor kappa B p65 subunit (p65) to translocate into the nucleus and regulate the transcription of genes involved in inflammation, cell survival, and tumor progression (17,21). Thus, the ROS/NF-κB axis represents a critical link between inflammation and cancer. Copper-mediated oxidative stress has emerged as an important activator of inflammatory signaling pathways, including NF-κB, signal transducer and activator of transcription 3 (STAT3), and hypoxia-inducible factor 1 alpha (HIF-1α), which collectively contribute to tumor progression and therapeutic resistance (17).

AID is a DNA-editing enzyme that is normally restricted to germinal center B cells, where it mediates somatic hypermutation and class switch recombination (26). However, aberrant expression of AID has been detected in multiple non-lymphoid cancers, including gastric, liver, esophageal, and bladder cancers, where it contributes to genomic instability and tumor progression (27). In a previous study, we demonstrated that AID is significantly upregulated in ccRCC tissues and is closely associated with tumor grade, stage, and poor prognosis; Functional experiments further showed that AID knockdown attenuated the malignant biological behaviors of ccRCC cells, supporting an important role for AID in ccRCC progression (28). Recent research suggests that AID expression can be induced by inflammatory signaling pathways and is closely associated with NF-κB activation in tumor-associated microenvironments, supporting a role for NF-κB-dependent regulation of AID in cancer progression (29) and suggesting a potential mechanistic link between inflammation and AID-driven tumorigenesis.

Based on these findings, we hypothesized that copper dyshomeostasis may promote renal tubular epithelial cell malignant transformation through a ROS/NF-κB/AID signaling axis. The aforementioned studies independently linked copper accumulation, oxidative stress, and NF-κB activation to tumor progression (8,17,19,28); however, research on the integrated regulatory network connecting copper metabolism with AID-mediated genomic instability in ccRCC is limited.

Thus, this study aimed to systematically investigate copper accumulation in ccRCC tissues and cell models, and to further elucidate its role in regulating ROS production, NF-κB activation, AID expression, and epithelial-mesenchymal transition (EMT). This study also evaluated the effects of copper chelation, ROS scavenging, and NF-κB inhibition on tumor-associated phenotypes. Our findings provide novel insights into copper-driven tumorigenesis and identify potential therapeutic targets for ccRCC. We present this article in accordance with the MDAR reporting checklist (available at https://tau.amegroups.com/article/view/10.21037/tau-2026-0795/rc).

Methods

Patient selection and tissue preparation

A total of six patients diagnosed with ccRCC at The First Affiliated Hospital, Hainan Medical University between March 2026 and June 2026 were enrolled in the study. Of the patients, five were male and one was female. The age of the patients ranged from 30 to 70 years.

An experienced pathologist identified the renal cancer tissue specimens and their corresponding inflammatory and non-tumor renal tissues. The samples were stored in liquid nitrogen or fixed in 4% formalin before Western blotting, immunohistochemistry (IHC), hematoxylin and eosin (HE) staining, or immunofluorescence (IF) analysis. This study was conducted in accordance with the Declaration of Helsinki and its subsequent amendments. The study was approved by the Ethics Committee of The First Affiliated Hospital, Hainan Medical University (No. 2025-KYL-153). All participants provided informed consent for the use of their samples and data.

Cell culture and treatments

Representative human renal tubular epithelial cells (HK-2) and two human renal cancer cell lines (786-O and 769-P) were obtained from ASY Biotechnology Ltd., Corp (Wuhan, China). Unless otherwise indicated, all cells were cultured in 89% Roswell Park Memorial Institute 1640 medium (Life Technologies, Gibco, USA) supplemented with 10% fetal bovine serum (FBS; Life Technologies) and 1% penicillin-streptomycin (Life Technologies). The cells were incubated in a culture flask at 37 ℃ in a humidified incubator with 5% CO₂ until 85–95% confluency was reached. The cells were then trypsinized and harvested for subsequent experiments.

For the in vitro experiments, the cells were divided into the following treatment groups after reaching approximately 70–80% confluence:

• HK-2 cells: (I) control group; (II) Copper (II) chloride (CuCl₂)-treated group, in which a chronic Cu exposure model was established by gradually increasing Cu concentrations over 2 weeks (7.5, 15, 22.5, and 30 μmol/L), followed by maintenance in Dulbecco’s modified eagle medium supplemented with 10% FBS and 30 μmol/L Cu; (III) hydrogen peroxide (H₂O₂)-treated group, treated with 8 μmol/L H₂O₂ for 2 h; and (IV) lipopolysaccharide (LPS)-treated group, treated with 20 μg/L LPS for 72 h.
• 786-O and 769-P cells: (I) control group; (II) triethylenetetramine dihydrochloride (TETA)-treated group, treated with 20 μmol/L TETA for 24 h; (III) N-acetyl-L-cysteine (NAC)-treated group, treated with 10 mmol/L NAC for 24 h; and (IV) pyrrolidine dithiocarbamate (PDTC)-treated group, treated with 20 μmol/L PDTC for 24 h.

CuCl₂ (#751944-25G), H₂O₂ (#1.08600), LPS (#L2630), NAC (#A9165), ammonium PDTC (#P8765), and TETA (#90460) were obtained from Sigma-Aldrich (USA).

Measurement of intracellular copper levels and ROS detection

In brief, tissue protein supernatants were collected, and tissue copper concentrations were measured using a copper ion assay kit (#E-BC-K300-M, Elabscience, Wuhan, China). The cells were lysed on ice for 30 min using radioimmunoprecipitation assay (RIPA) buffer (#P0013C, Beyotime Biotechnology, Shanghai, China), a protease inhibitor cocktail (#P1005, Beyotime Biotechnology), and phenylmethylsulfonyl fluoride (PMSF; #ST505, Beyotime Biotechnology) at a volume ratio of 100:4:1. Protein concentrations were determined using a bicinchoninic acid (BCA) protein assay kit (#P0010S, Beyotime Biotechnology). Intracellular copper concentrations were measured using a copper ion assay kit (#E-BC-K775-M，Elabscience, Wuhan, China). The copper ion concentration was expressed as micromoles per gram of protein (μmol/g prot).

For ROS detection, fresh surgical kidney tissues were harvested and washed in pre-cooled phosphate-buffered saline (PBS) to remove residual blood. The tissues were minced into ~1 mm³ pieces and digested in digestion buffer containing collagenase I (1 mg/mL, #ST3511, Beyotime Biotechnology) and Deoxyribonuclease I (DNase I; 50 U/mL, #D7073, Beyotime Biotechnology) at 37 ℃ with gentle shaking for 30 min. After digestion, the cell suspension was filtered through a 70-μm cell strainer and centrifuged at 300 ×g for 5 min. After removal of the supernatant, the cells were resuspended in PBS to prepare single-cell suspensions.

ROS levels were detected using a ROS assay kit (#S1105S, Beyotime Biotechnology). Single-cell suspensions were adjusted to 1×10⁶ cells/mL and incubated with 10 μmol/L 2’,7’-dichlorodihydrofluorescein diacetate (DCFH-DA) at 37 ℃ in the dark for 30 min, with gentle mixing every 5 min. The cells were then washed three times with pre-cooled PBS and analyzed by flow cytometry (BD Accuri™ C6 Plus). Dichlorofluorescein (DCF) fluorescence was excited at 488 nm and acquired in the fluorescein isothiocyanate (FITC) channel (530±15 nm). Cell debris and aggregates were excluded based on forward scatter and side scatter parameters, and at least 10,000 events were collected per sample. Mean fluorescence intensity was analyzed using FlowJo software (version 10.8.1) to quantify intracellular ROS levels.

Intracellular ROS levels were detected by collecting cells from each treatment group and incubating them with 10 μmol/L DCFH-DA (#S1105S, Beyotime Biotechnology) at 37 ℃ in the dark for 30 min. After washing three times with PBS, the cells were stained with 4′,6-diamidino-2-phenylindole (DAPI) for nuclear visualization. Fluorescence intensity was examined using a laser scanning confocal microscope (LSM900, ZEISS, Germany).

Western blot analysis

The cells were lysed on ice for 30 min using RIPA buffer (Beyotime Biotechnology, Shanghai, China), a protease inhibitor cocktail (Roche, Basel, Switzerland), and PMSF (Beyotime Biotechnology) at a volume ratio of 100:4:1. Protein concentrations were determined using a BCA protein assay kit (Beyotime Biotechnology). Since intracellular copper measurement was described earlier in the manuscript, this sentence is redundant and deleted.

The protein samples were mixed with 5× sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample loading buffer (Beyotime Biotechnology). In addition, 30 μg of total protein from each sample was isolated by 15% SDS-PAGE (Biotechwell, Shanghai, China) and transferred onto polyvinylidene difluoride (PVDF) membranes (#IPFL00010; Millipore, USA). The PVDF membranes were then blocked with 5% skimmed milk (#232100; BD Biosciences, USA) for 2 h, and incubated with primary antibodies at 4 ℃ overnight. Next, the samples were incubated with secondary antibodies for 2.5 h. The Chemiluminescence Imaging System (JP-K600, JIAPENG, China) was used to obtain the experiment results. The grayscale value of each experiment was analyzed using ImageJ software.

The primary antibodies used for Western blotting were anti-CTR1 (#67221-1-Ig, Proteintech), anti-ATP7A (#DF8506, Affinity), anti-E-cadherin (#14472; Cell Signaling Technology, USA), anti-phosphorylated NF-κB p65 (p-p65; #3033, Cell Signaling Technology, USA), anti-p65 (#D14E12; Cell Signaling Technology, USA), anti-vimentin (#5741; Cell Signaling Technology, USA), and anti-AID (#4975; Cell Signaling Technology, USA).

The membranes were then washed and incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (#A0208 and #A0216, Beyotime Biotechnology) for 1 h at room temperature. Protein bands were visualized using enhanced chemiluminescence reagents (#P0018S, Beyotime Biotechnology) and quantified using ImageJ software. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH; #AF2819, Beyotime Biotechnology) was used as an internal control.

IHC and HE staining

The paraffin-embedded tissue sections were deparaffinized in xylene and rehydrated through a graded ethanol series. Antigen retrieval was performed using citrate buffer in a microwave oven for 10 min, followed by cooling at room temperature for 30 min.

Endogenous peroxidase activity was blocked with 3% H₂O₂ for 10 min. The sections were blocked with 5% bovine serum albumin (BSA) for 1 h and incubated overnight at 4 ℃ with primary antibodies against CTR1, ATP7A, P-P65, AID, E-cadherin, and vimentin.

After incubation with HRP-conjugated secondary antibodies (#A0208 and #A0216, Beyotime Biotechnology) for 30 min, staining was visualized using diaminobenzidine (DAB). The sections were counterstained, dehydrated, and mounted.

For HE staining, ccRCC patient tissue samples were fixed in 4% paraformaldehyde (PFA), embedded in paraffin, and sectioned. The sections were deparaffinized in xylene for 20 min, rehydrated through graded ethanol for 30 min, and stained with HE.

Images were acquired using an orthophoto microscope (BX63; Olympus, Japan).

IF and F-actin staining

The cells were fixed with 4% PFA for 15 min, permeabilized with 0.3% Triton X-100 in PBS containing Tween-20 for 10 min at room temperature, and blocked with 5% BSA for 30 min. The samples were then incubated overnight at 4°C with primary antibodies against CTR1, ATP7A, P-P65, AID, E-cadherin, and vimentin. After washing with PBS, the cells were incubated with fluorescent secondary antibodies for 1 h at room temperature in the dark. The nuclei were counterstained with DAPI, and the slides were mounted using an anti-fade mounting medium. Fluorescent images were captured using a confocal laser scanning microscope (LSM900, Zeiss, Germany).

F- actin was visualized using the Beyotime Actin-Tracker Green-488 Kit (C2221S, Beyotime Biotechnology, China) according to the manufacturer’s instructions. Briefly, the cells were incubated with 1X Actin-Tracker Green-488 working solution (diluted from 100× stock) and Hoechst 33342 (1,000×) for nuclear counterstaining under light-protected conditions. The staining solution was added at 1 mL/well for six-well plates or 100 μL/well for 96-well plates. After incubation, the cells were washed with PBS, followed by overnight incubation with primary antibodies at 4 ℃. After further washing with PBS, the cells were incubated with fluorochrome-conjugated secondary antibodies at room temperature for 1 hour in the dark, and images were then captured using a laser confocal microscope (LSM900, Zeiss, Germany).

Wound-healing assay

The cells were seeded in six-well plates and cultured until reaching approximately 90% confluence. A sterile 200-μL pipette tip was used to create a linear scratch wound. Detached cells were removed by washing with PBS. The cells were then cultured in serum-free medium, and images were captured at 0 and 24 h under a microscope (Axio Observer 3, Zeiss, Germany). Migration ability was quantified using ImageJ software.

Colony formation assay

The cells were seeded in six-well plates at a density of 500 cells per well and cultured for 14 days. Colonies were fixed with 4% PFA and stained with crystal violet. Colonies containing ≥50 cells were counted under a microscope (BZ-X800, KEYENCE, Japan).

Transwell invasion assay

Matrigel (50 μg/mL) was diluted 1:9 with serum-free medium, added to the upper chambers of the Transwell inserts, and incubated at 37 ℃ for 4 h. The cells (8×10⁴ per well) suspended in serum-free medium were seeded into the upper chambers. The lower chambers were filled with medium containing 20% FBS. After incubation for 24 h, the cells were fixed with 4% PFA and stained with 0.1% crystal violet. The non-invading cells were removed, and the invading cells were counted in five random fields under a microscope (BZ-X800, KEYENCE, Japan).

EdU proliferation assay

Cell proliferation was assessed using an EdU incorporation assay kit (#C0078S, Beyotime Biotechnology) according to the manufacturer’s instructions. Fluorescence images were acquired using a confocal laser scanning microscope (LSM900, Zeiss, Germany), and the proportion of proliferating cells was quantified.

Statistical analysis

All in vitro experiments were independently repeated at least three times. All data presented in this study were obtained from biological replicates. Patient tissue samples and independently cultured cell samples were considered independent biological replicates and were used for the statistical analyses. Data are presented as mean ± standard deviation. Statistical analysis was performed using GraphPad Prism 11.0.0.

Differences between two groups were analyzed using the Student’s t-test. Comparisons among multiple groups were performed using one-way or two-way analysis of variance, followed by Tukey’s post hoc test. A P value <0.05 was considered statistically significant.

Results

Copper accumulation in ccRCC is associated with ROS elevation and activation of the NF-κB/AID axis and EMT

To investigate copper homeostasis in ccRCC, copper levels were first measured in ccRCC tissues, chronic nephritis tissues, and adjacent non-tumor tissues. As shown in Figure 1A, copper levels progressively increased from adjacent non-tumor tissues to chronic nephritis tissues, reaching the highest levels in ccRCC tissues (P<0.05), indicating significant copper accumulation during renal disease progression.

Figure 1 Copper accumulation, ROS elevation, NF-κB activation, AID overexpression, and EMT changes in ccRCC tissues. (A) Measurement of copper levels in adjacent non-tumor tissues, chronic nephritis tissues, and ccRCC tissues using a copper detection kit. Copper levels progressively increased from adjacent non-tumor tissues to chronic nephritis tissues, reaching the highest levels in ccRCC tissues. (B) Flow cytometric analysis of intracellular ROS levels in adjacent non-tumor tissues, chronic nephritis tissues, and ccRCC tissues. ROS levels gradually increased across the three tissue groups, reaching the highest level in ccRCC tissues. (C) Western blot analysis of CTR1, ATP7A, phosphorylated NF-κB p65 (p-p65), AID, E-cadherin, and vimentin expression in adjacent non-tumor tissues, chronic nephritis tissues, and ccRCC tissues. (D) Quantitative densitometric analysis of Western blot bands shown in (C). (E) Representative IHC staining of CTR1, ATP7A, p-p65, AID, E-cadherin, and vimentin in adjacent non-tumor tissues, chronic nephritis tissues, and ccRCC tissues (magnifications: 100× and 300×, scale bars: 100 and 50 μm). (F) Quantitative analysis of IHC staining intensity using ImageJ software. *, P<0.05; **, P<0.01; ***, P<0.001. AID, activation-induced cytidine deaminase; AOD, average optical density; ccRCC, clear cell renal cell carcinoma; EMT, epithelial-mesenchymal transition; FITC, fluorescein isothiocyanate; HE, hematoxylin and eosin; IHC, immunohistochemistry; NF-κB, nuclear factor-kappa B; ROS, reactive oxygen species.

Consistently, the flow cytometric analysis (Figure 1B) revealed that intracellular ROS levels were significantly increases in chronic nephritis tissues and further increased in ccRCC tissues compared with adjacent controls (P<0.05), suggesting that oxidative stress is associated with copper dysregulation in renal carcinogenesis.

Western blot analysis (Figure 1C) demonstrated that CTR1, ATP7A, p-p65, p65, AID, and vimentin were progressively upregulated, while E-cadherin was downregulated across adjacent non-tumor tissues, chronic nephritis tissues, and ccRCC tissues. The results of the quantitative densitometry analysis are shown in Figure 1D, confirming statistically significant differences among groups.

IHC analysis (Figure 1E) further validated these expression patterns, showing strong positive staining for CTR1, ATP7A, p-p65, AID, and vimentin in ccRCC tissues, but markedly reduced staining for E-cadherin. Quantification by ImageJ (Figure 1F) confirmed consistent trends.

Collectively, these findings indicate that copper accumulation in ccRCC is closely associated with ROS elevation, NF-κB activation, AID overexpression, and EMT progression.

Copper accumulation activates ROS/NF-κB signaling and promotes malignant phenotypes in renal cancer cells

To further validate copper dysregulation in vitro, intracellular copper levels were measured in HK-2, 786-O, and 769-P cells. As shown in Figure 2A, both renal cancer cell lines exhibited significantly higher copper levels than HK-2 cells (P<0.01).

Figure 2 Copper accumulation, ROS production, signaling activation, cytoskeletal remodeling, and proliferation in renal cells. (A) Measurement of intracellular copper levels in HK-2, 786-O, and 769-P cells as well as LPS-treated HK-2 cells using a copper detection assay. (B) Representative fluorescence images of intracellular ROS levels detected by DCFH-DA staining in different cell groups. (C) Quantification of ROS-associated fluorescence intensity in each group (magnifications: 200×, scale bars: 100 μm). (D) Western blot analysis of CTR1, ATP7A, p-p65, AID, E-cadherin, and vimentin expression in HK-2, LPS-treated HK-2, 786-O, and 769-P cells. (E) Quantitative analysis of Western blot band intensities. (F) IF staining of CTR1, p-p65, AID, E-cadherin, and F-actin expression in different cell groups, including fluorescence intensity correlation and co-localization analyses (magnifications: 400×, scale bars: 50 μm). (G) EdU incorporation assay evaluating cell proliferation capacity in different groups (magnifications: 200×, scale bars: 100 μm). *, P<0.05; **, P<0.01; ***, P<0.001. AID, activation-induced cytidine deaminase; DAPI, 4’,6-diamidino-2-phenylindole; DCFH-DA, 2’,7’-dichlorodihydrofluorescein diacetate; EdU, 5-ethynyl-2'-deoxyuridine; IF, immunofluorescence; LPS, lipopolysaccharide; ROS, reactive oxygen species.

ROS detection using DCFH-DA staining (Figure 2B,2C) showed significantly increased ROS levels in 786-O and 769-P cells compared with HK-2 cells (P<0.01), indicating increased oxidative stress in renal cancer cells.

To mimic inflammatory stimulation, the HK-2 cells were treated with LPS. The LPS treatment moderately increased intracellular copper levels and significantly increased ROS levels (Figure 2B,2C), suggesting that inflammatory NF-κB activation contributes to copper imbalance and oxidative stress.

Western blot analysis (Figure 2D) demonstrated that CTR1, ATP7A, p-p65, p65, AID, and vimentin were significantly upregulated, while E-cadherin was downregulated in renal cancer cells compared with HK-2 cells. The results of the quantitative analysis are shown in Figure 2E.

IF staining (Figure 2F) corroborated these findings and demonstrated consistent expression patterns of AID with NF-κB and E-cadherin. Moreover, IF staining revealed pronounced cytoskeletal remodeling. Compared with HK-2 cells, cancer cells exhibited markedly enhanced F-actin fluorescence signals and a more organized filamentous arrangement.

Correlation analysis (Figure 2F) revealed a positive association between CTR1 and F-actin expression, while p-p65 and AID exhibited strong co-localization, suggesting functional coupling between NF-κB activation and AID expression. In contrast, E-cadherin and vimentin showed opposite expression patterns, consistent with EMT progression.

EdU assays (Figure 2G) demonstrated significantly increased proliferative capacity in 786-O and 769-P cells compared with HK-2 cells, which was further enhanced following inflammatory stimulation.

Copper induces malignant transformation of renal tubular epithelial cells via ROS/NF-κB/AID signaling

To investigate whether copper accumulation promotes malignant transformation, HK-2 cells were treated with CuCl₂, H₂O₂, or LPS to model copper accumulation, oxidative stress, or an inflammatory microenvironment, respectively.

As shown in Figure 3A, all treatment groups showed significantly increased intracellular copper levels, with the CuCl₂ group exhibiting the strongest effect (P<0.05).

Figure 3 Copper-induced malignant transformation of HK-2 cells via ROS-dependent NF-κB/AID signaling. (A) Intracellular copper levels in HK-2 cells after treatment with CuCl₂, H₂O₂, and LPS. (B)Representative fluorescence images of ROS levels detected by DCFH-DA staining in different treatment groups. (C) Quantification of ROS-associated fluorescence intensity in HK-2 cells following different treatments (magnifications: 200×, scale bars: 100 μm). (D) Western blot analysis of CTR1, ATP7A, p-p65, AID, E-cadherin, and vimentin expression in treated HK-2 cells. (E) Quantitative densitometric analysis of Western blot results. (F)IF staining showing expression and localization of CTR1, p-p65, AID, E-cadherin, and F-actin in treated HK-2 cells (magnifications: 400×, scale bars: 50 μm). (G) EdU assay showing the proliferative ability of HK-2 cells under different treatments (magnifications: 200×, scale bars: 100 μm). *, P<0.05; **, P<0.01; ***, P<0.001. AID, activation-induced cytidine deaminase; DAPI, 4’,6-diamidino-2-phenylindole; DCFH-DA, 2’,7’-dichlorodihydrofluorescein diacetate; EdU, 5-ethynyl-2'-deoxyuridine; IF, immunofluorescence; LPS, lipopolysaccharide; NF-κB, nuclear factor-kappa B; ROS, reactive oxygen species.

ROS detection (Figure 3B,3C) revealed that CuCl₂, H₂O₂, and LPS treatment also significantly increased intracellular ROS levels.

Western blot analysis (Figure 3D,3E) showed that all treatments significantly upregulated CTR1, ATP7A, p-p65, AID, and vimentin, while downregulating E-cadherin. The quantitative results are presented in Figure 3E.

IF staining (Figure 3F) confirmed some of these expression changes at the cellular level.

Functional assays demonstrated that the CuCl₂, H₂O₂, and LPS treatments significantly promoted HK-2 cell proliferation (Figure 3G) as assessed by EdU assay, indicating enhanced malignant transformation ability.

Together, these findings indicate that copper overload is associated with increased ROS production and activation of the NF-κB/AID signaling axis, accompanied by EMT and malignant transformation of renal tubular epithelial cells.

Copper depletion reverses ccRCC malignant phenotypes via inhibition of the ROS/NF-κB/AID axis (Figures 4,5)

To investigate whether targeted modulation of copper homeostasis could reverse malignant phenotypes, 769-P and 786-Ocells were treated with TETA, NAC, or PDTC to simulate environments with reduced copper ions, diminished oxidative stress, or suppressed inflammation, respectively.

As shown in Figures 4A,5A, both TETA and NAC significantly reduced intracellular copper levels in the ccRCC cells.

Figure 4 Effects of copper chelation, ROS inhibition, and NF-κB inhibition on the malignant phenotypes of 769-P cells. (A) Intracellular copper levels in 769-P cells after treatment with TETA, NAC, and PDTC. (B)Quantification of ROS-associated fluorescence intensity in treated 769-P cells. (C) Representative DCFH-DA fluorescence images showing ROS levels in different treatment groups (magnifications: 200×, scale bars: 100 μm). (D) Western blot analysis of CTR1, ATP7A, p-p65, AID, E-cadherin, and vimentin expression in 769-P cells. (E)Quantitative analysis of Western blot band intensities. (F) IF analysis of protein expression and cytoskeletal changes involving CTR1, p-p65, AID, E-cadherin, and F-actin (magnifications: 400×, scale bars: 50 μm). (G)Transwell invasion assay showing the invasive capacity of 769-P cells (magnification: 200×, scale bar: 50 μm). (H) Colony formation assay showing the proliferative ability of 769-P cells. (I) Wound-healing assay showing the migratory capacity of 769-P cells (magnifications: 40×, scale bars: 250 μm). (J) Quantitative analysis of Transwell invasion, colony formation, and wound-healing assays. ns, P≥0.05; *, P<0.05; **, P<0.01; ***, P<0.001. AID, activation-induced cytidine deaminase; DAPI, 4’,6-diamidino-2-phenylindole; DCFH-DA, 2’,7’-dichlorodihydrofluorescein diacetate; IF, immunofluorescence; NAC, N-acetyl-L-cysteine; NF-κB, nuclear factor-kappa B; ns, not significant; PDTC, pyrrolidine dithiocarbamate; ROS, reactive oxygen species; TETA, triethylenetetramine.

Figure 5 Effects of copper chelation, ROS inhibition, and NF-κB inhibition on the malignant phenotypes of 786-O cells. (A) Intracellular copper levels in 786-O cells after treatment with TETA, NAC, and PDTC. (B)Quantification of ROS-associated fluorescence intensity in treated 786-O cells. (C) Representative DCFH-DA fluorescence images showing ROS levels in different treatment groups (magnifications: 200×, scale bars: 100 μm). (D) Western blot analysis of CTR1, ATP7A, p-p65, AID, E-cadherin, and vimentin expression in 786-O cells. (E)Quantitative analysis of Western blot band intensities. (F) IF analysis of protein expression and cytoskeletal changes involving CTR1, p-p65, AID, E-cadherin, and F-actin (magnifications: 400×, scale bars: 50 μm). (G)Transwell invasion assay showing the invasive capacity of 786-O cells (magnification: 200×, scale bar: 50 μm). (H) Colony formation assay showing the proliferative ability of 786-O cells. (I) Wound-healing assay showing the migratory capacity of 786-O cells (magnifications: 40×, scale bars: 250 μm). (J) Quantitative analysis of Transwell invasion, colony formation, and wound-healing assays. ns, P≥0.05; *, P<0.05; **, P<0.01; ***, P<0.001. AID, activation-induced cytidine deaminase; DAPI, 4’,6-diamidino-2-phenylindole; DCFH-DA, 2’,7’-dichlorodihydrofluorescein diacetate; IF, immunofluorescence; NAC, N-acetyl-L-cysteine; NF-κB, nuclear factor-kappa B; ns, not significant; PDTC, pyrrolidine dithiocarbamate; ROS, reactive oxygen species; TETA, triethylenetetramine.

ROS assays (Figures 4B,4C,5B,5C) demonstrated that TETA, NAC, and PDTC markedly decreased intracellular ROS production.

Western blot analysis (Figures 4D,4E,5D,5E) revealed that TETA and NAC significantly downregulated CTR1, ATP7A, p-p65, AID, and vimentin, while restoring E-cadherin expression. In contrast, PDTC specifically inhibited p-p65, AID, and vimentin but had no obvious effect on CTR1 or ATP7A, suggesting that NF-κB functions downstream of copper regulation.

IF staining (Figures 4F,5F) further confirmed these protein expression changes and demonstrated partial reversal of EMT-related cytoskeletal remodeling.

Functional assays showed that TETA, NAC, and PDTC significantly inhibited invasion (Figure 4G,5G), proliferation (Figures 4H,5H), and migration (Figures 4I,5I) in both 769-P and 786-O cells. The quantitative analysis results are shown in Figures 4J,5J.

Together, these results indicate that copper depletion suppresses ROS accumulation, inhibits NF-κB activation, downregulates AID expression, and reverses EMT and malignant phenotypes in ccRCC cells.

Working model

Taken together, our data suggest that copper accumulation promotes malignant transformation-related changes in renal tubular epithelial cells and contributes to ccRCC progression through a ROS/NF-κB/AID signaling axis, while copper chelation effectively disrupts this pathway and reverses tumor-associated phenotypes.

Discussion

The global incidence of renal cell carcinoma (RCC) has continued to rise in recent years, ccRCC is the predominant subtype of RCC (1,2). Despite advances in surgical resection, targeted therapies, and ICIs, the prognosis of patients with advanced disease remains poor due to frequent drug resistance and tumor recurrence. Therefore, the molecular mechanisms underlying renal tubular epithelial cell malignant transformation, especially in the context of chronic inflammation and metabolic dysregulation, urgently need to be identified to establish novel biomarkers and therapeutic targets.

In this study, we demonstrated that copper homeostasis imbalance plays a central role in ccRCC progression through the activation of a ROS/NF-κB/AID signaling axis. Our results showed that copper levels were significantly elevated in ccRCC tissues compared with adjacent non-tumor tissues, accompanied by increased ROS production, NF-κB activation, AID overexpression, and enhanced EMT. These findings suggest that copper dysregulation is not merely a metabolic alteration but a driver of malignant transformation in renal epithelial cells.

Increasing evidence indicates that copper metabolism reprogramming represents an important metabolic feature of malignant tumors. Although copper is an essential trace element required for mitochondrial respiration and enzymatic activity, excessive intracellular copper accumulation can promote tumor progression by regulating oxidative stress, metabolic adaptation, and oncogenic signaling pathways. Recent studies have proposed the concept of “cuproplasia” in cancer cells, suggesting that malignant cells actively maintain elevated copper levels to satisfy their increased metabolic demands (18,30).

Dysregulation of copper transporters, particularly CTR1, the major high-affinity copper importer, and ATP7A, an important regulator of intracellular copper trafficking and copper efflux, represents a critical mechanism underlying tumor-associated copper accumulation (9,30-32). We found that both CTR1 and ATP7A were upregulated in tumor tissues and cells. The elevated expression of these two copper-related proteins may indicate enhanced copper metabolic activity, consistent with the increased copper levels observed in ccRCC tissues and cells.

Excess copper accumulation can induce oxidative stress through redox-active reactions and mitochondrial dysfunction, resulting in increased ROS production. However, copper toxicity may also involve ROS-independent effects, including direct interactions with proteins, protein mismetallation, aggregation, and disruption of protein homeostasis (33,34). Thus, while our findings support an important role for ROS in copper-associated signaling, other ROS-independent effects of copper cannot be excluded. Nevertheless, ROS are not merely a byproducts of metabolic stress but also function as signaling molecules capable of activating multiple oncogenic pathways. Previous studies have demonstrated that ROS-mediated activation of NF-κB promotes tumor cell proliferation, survival, EMT, and invasion by regulating inflammatory and metastasis-associated genes (35-37).

In the present study, increased ROS levels were consistently observed in ccRCC tissues and copper-treated renal epithelial cells, while ROS scavenging effectively attenuated downstream signaling activation, supporting an important role for oxidative stress in copper-induced tumorigenic effects.

NF-κB is a central transcription factor that integrates inflammatory and oxidative stress signals (17,21). Under oxidative conditions, NF-κB is activated through the phosphorylation and degradation of IκB, leading to nuclear translocation of p65 and the transcriptional activation of downstream target genes involved in inflammation and tumor progression. Our results showed that NF-κB signaling was significantly activated in ccRCC tissues and copper-stimulated cells. Notably, ROS inhibition markedly suppressed NF-κB activation, suggesting that copper-induced oxidative stress acts upstream of NF-κB signaling in renal epithelial cells.

NF-κB activation may further contribute to tumor evolution by inducing AID, a DNA-editing enzyme involved in inflammation-associated genomic instability. AID is physiologically responsible for antibody diversification in B cells; however, aberrant AID expression in epithelial cells can promote accumulation of DNA mutations and accelerate carcinogenesis (38-40). In a previous study, we demonstrated that AID is significantly upregulated and closely associated with tumor grade, stage, and poor prognosis in ccRCC (28). In the present study, we further confirmed that NF-κB signaling regulates AID expression, consistent with previous reports showing that inflammatory signaling pathways can induce ectopic AID expression in non-lymphoid tissues (9).

Our findings further extend the biological significance of copper dysregulation in ccRCC. Beyond its established role in metabolic remodeling, we identified AID as a previously unrecognized downstream effector of copper-induced ROS/NF-κB signaling. This observation provides a mechanistic link between copper accumulation, inflammatory signaling, genomic instability, and epithelial plasticity during renal carcinogenesis.

Functionally, activation of the Copper/ROS/NF-κB/AID signaling axis was associated with EMT, characterized by the downregulation of E-cadherin and the upregulation of vimentin, as well as enhanced proliferation, migration, and invasion of renal epithelial cells. Conversely, copper chelation (TETA), ROS inhibition (NAC), and NF-κB blockade (PDTC) effectively suppressed these malignant phenotypes, supporting the functional involvement of this signaling cascade in ccRCC progression.

Taken together, our findings suggest that copper accumulation contributes to renal tubular epithelial cell malignant transformation and ccRCC progression potentially through ROS-associated activation of the NF-κB/AID signaling axis. This mechanism provides a potential link between metal metabolism dysregulation, oxidative stress, inflammatory signaling, and genomic instability in renal carcinogenesis. However, the present findings do not exclude additional ROS-independent effects of copper ions, and further studies are needed to clarify the relative contributions of these mechanisms.

From a therapeutic perspective, targeting copper metabolism or its downstream signaling pathways may represent a promising strategy for ccRCC treatment. In this study, copper chelation (TETA), antioxidant treatment (NAC), and NF-κB inhibition (PDTC) all effectively suppressed tumor-associated phenotypes in vitro, highlighting their potential translational value (7,17,22).

However, this study had several limitations. First, in vivo validation using appropriate animal models is warranted to further confirm the role of the copper-ROS-NF-κB/AID axis in ccRCC progression. Second, although our findings support an important role for ROS in copper-associated NF-κB/AID signaling, mitochondrial ROS, oxidative stress markers, and potential ROS-independent effects of copper were not independently assessed, warranting further investigation.

Conclusions

In conclusion, this study identified a previously unrecognized copper-driven ROS/NF-κB/AID signaling axis that promotes EMT and malignant transformation in ccRCC. These findings provide novel insights into the role of copper metabolism in renal carcinogenesis and suggest potential therapeutic targets for the early diagnosis and treatment of ccRCC.

Acknowledgments

The authors would like to thank our senior colleagues and fellow students at Hainan Medical University, whose support was instrumental in the completion of the majority of this work.

Footnote

Reporting Checklist: The authors have completed the MDAR reporting checklist. Available at https://tau.amegroups.com/article/view/10.21037/tau-2026-0795/rc

Data Sharing Statement: Available at https://tau.amegroups.com/article/view/10.21037/tau-2026-0795/dss

Peer Review File: Available at https://tau.amegroups.com/article/view/10.21037/tau-2026-0795/prf

Funding: This work was supported by the Hainan Provincial Natural Science Foundation of China (Nos. 822RC839, 825RC897, and 326MS0410), the Academic Enhancement Support Program of Hainan Medical University (No. XSTS2025005), the National Natural Science Foundation of China (No. 82560531), the College Students’ Research Innovation Training Program of Hainan Medical University (No. RZ2500002143), the Hainan Provincial Joint Project for Health Science and Technology Innovation (No. WSJK2026QN082), and Zhumadian Science and Technology Key-research Project of China (No. ZMDSKJGG2025004).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://tau.amegroups.com/article/view/10.21037/tau-2026-0795/coif). All authors report that this work was supported by the Hainan Provincial Natural Science Foundation of China (Nos. 822RC839, 825RC897, and 326MS0410), the Academic Enhancement Support Program of Hainan Medical University (No. XSTS2025005), the National Natural Science Foundation of China (No. 82560531), the College Students’ Research Innovation Training Program of Hainan Medical University (No. RZ2500002143), the Hainan Provincial Joint Project for Health Science and Technology Innovation (No. WSJK2026QN082), and Zhumadian Science and Technology Key-research Project of China (No. ZMDSKJGG2025004). The authors have no other conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. This study was conducted in accordance with the Declaration of Helsinki and its subsequent amendments. The study was approved by the Ethics Committee of The First Affiliated Hospital, Hainan Medical University (No. 2025-KYL-153). All participants provided informed consent for the use of their samples and data.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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