MND1 protein mediates DNA damage repair and is a promising novel therapeutic target in hepatocellular carcinoma

Abstract

Background and aims

Hepatocellular carcinoma (HCC) remains a leading cause of cancer-related mortality worldwide, highlighting the urgent need to identify novel therapeutic targets. Bioinformatic analysis initially identified meiotic nuclear division 1 (MND1) as a differentially expressed gene in HCC, with significant associations to DNA damage repair pathways, suggesting its potential functional relevance in hepatocarcinogenesis. Based on these findings, this study aims to systematically characterize the expression profile of MND1 in HCC and to further elucidate its biological functions as well as the underlying molecular mechanisms driving HCC progression.

Methods

The expression profile of MND1 was validated using data from public databases along with 22 paired clinical HCC specimens. The functional roles of MND1 were investigated through in vitro assays (CCK-8, Transwell, and flow cytometry) and in vivo subcutaneous/orthotopic xenograft mouse models. Mechanistically, co-immunoprecipitation (Co-IP) coupled with mass spectrometry, immunofluorescence, and rescue experiments were applied to identify interacting proteins and to elucidate the underlying molecular pathways.

Results

Multi-omics analysis and clinical validation revealed that MND1 is overexpressed in HCC tissues and significantly correlates with poor prognosis. Comprehensive in vitro and in vivo functional studies further demonstrated that MND1 drives HCC progression by regulating tumor cell proliferation, migration, and invasion. Mechanistically, we uncovered a close functional association between MND1 and TRIM28, which promotes genomic instability by enhancing the activity of the DNA damage response (DDR) pathway.

Conclusions

These findings suggest that MND1 may serve as both a potential molecular marker and an oncogenic contributor in HCC. Importantly, the newly identified MND1–TRIM28–DDR regulatory axis offers a promising rationale for the development of targeted therapeutic strategies.

1. Introduction

Liver cancer remains one of the most common malignancies worldwide, with its incidence ranking sixth and mortality ranking third among all cancers according to the latest global cancer statistics.¹ Liver cancer development is associated with multiple established risk factors, including chronic viral hepatitis, particularly those caused by hepatitis B virus (HBV) and hepatitis C virus (HCV), excessive alcohol consumption, exposure to dietary carcinogens such as aflatoxin B1, and genetic predisposition.² Systemic therapy has become the standard-of-care for patients with advanced hepatocellular carcinoma (HCC). However, primary or acquired drug resistance frequently emerges during treatment.³ Recent advances in single-cell sequencing and spatial transcriptomics have further revealed the intratumoral heterogeneity of liver tumors.^4,5 Systematic characterization of patient molecular profiles will enable more precise molecular subtyping and facilitate the development of targeted therapeutic strategies for HCC. Therefore, there is an urgent need to elucidate the pathogenesis of liver cancer and identify potential molecular targets for both diagnostic biomarkers and personalized therapeutic interventions.

DNA is the fundamental carrier of genetic information in humans. Exogenous and endogenous agents can act directly or indirectly on DNA, inducing structural and functional alterations that result in DNA damage. To ensure high-fidelity replication and maintain the integrity of genomes, the mechanisms by which cells perceive, respond to, repair, or tolerate DNA damage are collectively referred to as the DNA damage response (DDR).⁶ Cancer cells frequently harbor mutations and dysregulation of DDR-related genes, which drive genomic instability and promote tumor progression.⁷ In related studies, DDR has been found to affect tumor progression in various ways, including the expression imbalance of oncogenes and tumor suppressor genes, tumor immune escape, and oxidative stress.⁸ DDR is not only implicated in cancer progression but also plays an important role in the field of tumor drug resistance.9, 10, 11 Understanding the DNA damage repair mechanism may help us identify possible therapeutic targets for HCC. However, the precise roles of DDR in driving or inhibiting hepatocarcinogenesis still need further investigation.

Meiotic nuclear division 1 (MND1), a meiotic-specific protein, is highly expressed in the nucleus. MND1 plays an essential role in homologous recombination by forming a functional heterodimer with homologous pairing 2 (HOP2), a conserved meiotic protein first identified in Saccharomyces cerevisiae. Deletion of either MND1 or HOP2 results in complete arrest at meiotic prophase. The MND1–HOP2 complex facilitates homologous recombination-mediated DNA repair not only during meiosis but also in mitotic cells.^12,13 MND1 also promotes the repair of DNA double-strand breaks (DSBs) that arise during meiotic homologous recombination.¹⁴ Recent studies have shown that MND1 competitively binds to the tumor suppressor Krüppel-like factor 6 (KLF6), thereby preventing KLF6-mediated transcriptional repression of E2F transcription factor 1 (E2F1), highlighting its oncogenic potential.¹⁵ Emerging evidence links elevated MND1 expression to tumor aggressiveness in several cancer types.16, 17, 18 Despite compelling evidence linking MND1 to the DDR in various cancers, its specific role in HCC remains poorly understood. Elucidating the molecular mechanisms by which MND1 regulates DDR during hepatocarcinogenesis is therefore essential to address this critical gap in liver cancer biology.

Tripartite motif-containing 28 (TRIM28), also known as transcriptional intermediary factor 1β (TIF1β) or KRAB-associated protein 1 (KAP1), is a key member of the TRIM protein family. As a multifunctional scaffold protein, TRIM28 contains several conserved structural domains: an N-terminal RING finger domain that confers E3 ubiquitin ligase activity, followed by tandem B-box zinc-binding domains and a leucine zipper-containing coiled-coil (CC) region that mediates protein oligomerization. Emerging evidence implicates TRIM28 as a central regulator of genomic stability, orchestrating key components of the DDR pathway. Notably, TRIM28 is frequently overexpressed in a wide range of cancers, where it promotes oncogenic transformation and tumor progression through multiple mechanisms.^19,20 In HCC, TRIM28 overexpression has been consistently linked to enhanced tumor cell proliferation, underscoring its potential as a therapeutic target. This study provides evidence for a physical and functional interaction between MND1 and TRIM28 that promotes tumor progression in HCC.

Based on these findings and bioinformatic predictions, this study aims to systematically elucidate the expression pattern of MND1 in HCC and further dissect its biological roles and underlying molecular regulatory mechanisms in hepatocarcinogenesis through gain- and loss-of-function experiments.

2. Materials and methods

2.1. Ethical approval

This study was conducted in compliance with the Declaration of Helsinki and approved by the Ethics Committee of Nanjing Medical University (approval number: NJMU2020-467; approval date: March 21, 2020), and written informed consent was obtained from all participants. All animal procedures were carried out in compliance with the ARRIVE guidelines and adhered to the guidelines for the care and use of laboratory animals issued by the China Animal Research Council. All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Nanjing Medical University (approval number: IACUC-2307012).

2.2. Cell lines, animals, and clinical specimens

The human HCC cell lines MIHA, MHCC97L, HCCLM3, MHCC97H, JHH7, Bel7402, Hep3B, PLC/PRF/5, and SNU-398 were obtained from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China), the American Type Culture Collection (ATCC; Manassas, VA, USA), the Korean Cell Line Bank (KCLB; Seoul, Korea), and the Japanese Collection of Research Bioresource Cell Bank (JCRB Cell Bank; Osaka, Japan). HEK293T cells were sourced from ATCC (Manassas, VA, USA). All cell lines were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% penicillin/streptomycin (Solarbio, Beijing, China), and maintained at 37 °C in a humidified atmosphere containing 5% CO₂. Male nude mice (6 weeks old) were provided by the Animal Core Facility of Nanjing Medical University (Nanjing, China) and acclimated to a 12-h light/dark cycle under specific pathogen-free (SPF) conditions. Twenty-two paired clinical HCC tissue samples and matched adjacent non-tumor liver tissues were collected from patients undergoing surgical resection at Nanjing Drum Tower Hospital, The Affiliated Hospital of Nanjing University Medical School between May 2020 and May 2021.

2.3. Quantitative real-time polymerase chain reaction (qRT-PCR)

Total RNA was extracted from cell lines or tissues using an RNA-Quick Purification Kit (Vazyme Biotech Co., Ltd., Nanjing, China) and used as a template to generate cDNA using a Reverse Transcription Kit (Vazyme Biotech Co., Ltd., Nanjing, China). qPCR was performed with ChamQ SYBR qPCR Master Mix (Vazyme Biotech Co., Ltd., Nanjing, China) on the LightCycler 480 system (Roche, Switzerland). The primer sequences are shown in Table 1.

Table 1.

Sequences of the primers used in quantitative real-time polymerase chain reaction (qRT-PCR).

Gene name	Forward primer sequence (5′-3′)	Reverse primer sequence (5′-3′)
GAPDH	ACCCAGAAGACTGTGGATGG	TTCAGCTCAGGGATGACCTT
MND1	TGTGAGAGGATCGGAACTTCT	CACATCGGCCAATTTTAGCTTTC
CDK2	CAGAGCTTTTGGAGTCCCTGT	AAAGATCCGGAAGAGCTGGT
CDC14B	GTGCCATTGCAGTACATT	AGCAGGCTATCAGAGTG
CDC25A	TGTGCCGGTATGTGAGAGAG	TGCGGAACTTCTTCAGGTCT
CDKN1A	GACTCTCAGGGTCGAAAACG	TAGGGCTTCCTCTTGGAGAA
E2F8	CCAACCCTGCTGTGAATA	TTTCTGGCTCATTACCCT

2.4. Transfection

Small interfering RNA (siRNA), short hairpin RNA (shRNA), MND1 overexpression vector, and lentiviral particles were obtained from Wuhan Miaoling Biology Co., Ltd. (Wuhan, China). Polybrene (Sigma-Aldrich, St. Louis, MO, USA) and Lipo8000™ transfection reagent (Beyotime Biotechnology, Shanghai, China) were used for the transient transfection of siRNAs or shRNAs into cells, in accordance with the manufacturer’s instructions. Lentiviral particles were produced and transfected into the cells to obtain stable clones. The sequences for the siRNAs targeting MND1 (siMND1) were as follows: siMND1#1, 5′-GCUUAGUUGAUGAUGGUAUTT-3′; siMND1#2, 5′-GAAACGGCCAAGCAAAUAATT-3′. The sequences for the siRNAs targeting TRIM28 (siTRIM28) were as follows: 5′-CAGUGCUGCACUAGCUGUGAGGAUA-3′. The sequences of the negative control siRNA (siNC) were 5′-UUCUCCGAACGUGUCACGUTT-3′. The sequences for the shRNAs targeting MND1 (shMND1) were as follows: shMND1#1, 5′-GCTGCTAACAGATGGACTGAT-3′; shMND1#2, 5′-CAAGTAAAGCTCTTCATGCAA-3′. The sequences of the negative control shRNA (shNC) were 5′-TTCTCCGAACGTGTCACGT-3′.

2.5. Western blotting

At the indicated time points, cells were lysed using radioimmunoprecipitation assay (RIPA) buffer supplemented with protease inhibitors. The protein concentrations of the cell lysates were determined using a BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA). Following electrophoresis, protein lysates were transferred onto polyvinylidene fluoride (PVDF) membranes (Merck Millipore, Darmstadt, Germany). Subsequently, the membranes were blocked with 5% non-fat milk, incubated with primary antibodies against β-actin (Proteintech, Wuhan, China), GAPDH (Proteintech, Wuhan, China), MND1 (Proteintech, Wuhan, China), and TRIM28 (Abclonal, Wuhan, China).

2.6. Animal experiments

2.6.1. Subcutaneous xenograft tumor model

HCCLM3 cells stably transduced with either a lentiviral shRNA targeting MND1 (shMND1 group) or a non-targeting control shRNA (shCon group; 4 × 10⁶ cells per injection) were bilaterally inoculated into the dorsal subcutaneous flanks of 6-week-old male nude mice (n = 6 per group). Tumor dimensions were measured weekly using digital calipers, and tumor volume was calculated as follows: (length × width²)/2. At day 21 post-inoculation, mice were euthanized, and tumors were excised, photographed, and weighed for comparative analysis.

2.6.2. Orthotopic liver tumor model

HCCLM3 cells stably expressing shMND1 or shCon (2 × 10⁶ cells in 50 μL phosphate-buffered saline) were surgically implanted into the left lobe of the liver in anesthetized nude mice. Five weeks after implantation, mice were euthanized, and body weight and liver weight were recorded. The liver-to-body weight ratio was calculated as an indicator of tumor burden. Tumor burden was further assessed by macroscopic measurement of visible hepatic lesions with vernier calipers (Mitutoyo Corporation, Kanagawa, Japan).

2.7. Immunofluorescent staining

Cells were seeded onto glass-bottom confocal dishes (Thermo Fisher Scientific, Waltham, MA, USA) at a density of 2 × 10⁴ cells per well and allowed to adhere overnight at 37 °C. After fixation with 4% paraformaldehyde (PFA; Thermo Fisher Scientific, Waltham, MA, USA) in phosphate-buffered saline (PBS; Beyotime Biotechnology, Shanghai, China) for 15 min at room temperature, cells were permeabilized with 0.1% Triton X-100 (Beyotime Biotechnology, Shanghai, China) in PBS for 10 min and blocked with 5% bovine serum albumin (BSA; Solarbio Science & Technology Co., Ltd., Beijing, China) in PBS for 30 min at room temperature. Cells were then incubated with primary antibodies against 53BP1 (1:100 dilution; ABclonal, Wuhan, China) and γH2AX (1:100 dilution; ABclonal) diluted in blocking buffer at 4 °C overnight. Following three washes with PBS, cells were incubated with appropriate fluorescently labeled secondary antibodies (Proteintech, Wuhan, China) and counterstained with 4′,6-diamidino-2-phenylindole (DAPI; Beyotime Biotechnology, Shanghai, China) for nuclear visualization.

2.8. Cell migration and invasion assay

Cell migration and invasion were assessed using 24-well Transwell chambers with 8-μm pore polycarbonate membranes (Corning Inc., Corning, NY, USA). For migration assays, cells were serum-starved in serum-free medium (Gibco) for 6 h before seeding. A total of 5 × 10⁴ cells in 200 μL serum-free medium (Gibco) were seeded into the upper chamber, while the lower chamber was filled with 600 μL complete medium supplemented with 20% FBS (Gibco). For invasion assays, the upper chamber was pre-coated with 40 μL of Matrigel (diluted 1:8 in serum-free medium; BD Biosciences, San Jose, CA, USA) and polymerized at 37 °C for 1 h before cell seeding. Following incubation at 37 °C for 48 h, non-migratory or non-invasive cells on the upper surface of the membrane were gently removed with a cotton swab. Cells that had migrated or invaded to the lower surface were fixed with 4% PFA (Thermo Fisher Scientific) for 15 min, stained with 0.1% crystal violet (Beyotime Biotechnology, Shanghai, China) for 20 min, and washed three times with PBS. Images of five random fields per membrane were captured under a light microscope ( × 200 magnification, Olympus, Tokyo, Japan), and the number of stained cells was quantified using ImageJ software (version 1.6.0, the National Institutes of Health, USA). All experiments were performed in triplicate and repeated at least three times independently to ensure reproducibility.

2.9. Cell cycle analysis

MND1-knockdown, MND1-overexpressing, and control cells were seeded in 6-well plates at a density of 1 × 10⁶ cells per well (n = 3 biological replicates). After 48 h, cells were harvested, washed twice with cold PBS, and fixed in ice-cold 75% ethanol (prepared in PBS) at −20 °C overnight. Fixed cells were then washed twice with PBS, resuspended in PBS containing 100 μg/mL RNase A (Yeasen Biotechnology, Shanghai, China), and incubated at 37 °C for 30 min. Subsequently, cells were stained with propidium iodide (PI; Yeasen Biotechnology) for 30 min at room temperature in the dark. Cell cycle distribution was analyzed by flow cytometry (BD FACS Calibur; BD, Franklin Lakes, NJ, USA), and the percentages of cells in G0/G1, S, and G2/M phases were quantified using FlowJo software (version 10.8.1, BD Biosciences).

2.10. Cell proliferation assay

Cell proliferation was evaluated using the Cell Counting Kit-8 (CCK-8; Yeasen Biotechnology, Shanghai, China) following the manufacturer’s instructions. Briefly, cells were seeded into 96-well plates at a density of 2 × 10³ cells per well, with cell-free blank wells and appropriate control (e.g., non-targeting shRNA) and experimental (e.g., MND1-knockdown) groups included. At 0, 24, 48, and 72 h post-seeding, 10 μL of CCK-8 solution was added to each well, and plates were incubated at 37 °C for 2 h. Absorbance was measured at 450 nm using a microplate reader (Thermo Fisher Scientific, Waltham, MA, USA).

2.11. Colony formation assay

For the colony formation assay, 2 × 10³ MND1-knockdown or control cells were seeded into each well of a 6-well plate. The culture medium was replaced every 3 days. After 14 days, colonies were fixed with 4% PFA (Thermo Fisher Scientific) in PBS for 30 min at room temperature, stained with 0.5% crystal violet (Beyotime Biotechnology) for 30 min, and rinsed gently with PBS. Colonies containing more than 50 cells were counted, and representative images were acquired using a digital imaging system (Leica Microsystems, Wetzlar, Germany). All experiments were performed with at least three independent biological replicates to ensure reproducibility.

2.12. EdU proliferation assay

HCC cells were seeded into fluorescence culture dishes at a density of 4 × 10⁵ cells per dish and cultured for 24 h. Following the manufacturer’s instructions, the cells were stained with EdU and DAPI using an EdU proliferation kit (Yeasen Biotechnology, Shanghai, China). After staining, cells were maintained in PBS to prevent drying. The stained cells were observed under the fluorescence microscope (Olympus, Tokyo, Japan), and images of EdU-positive cells were captured immediately to prevent fluorescence quenching.

2.13. TUNEL assay

Apoptosis was detected using the TUNEL Apoptosis Detection Kit (Alexa Fluor 488; Cat. No. 40307ES20; Yeasen Biotechnology). Harvested cells were resuspended in PBS at a density of approximately 2 × 10⁷ cells/mL. Aliquots of 50–100 μL of the cell suspension were transferred into fluorescence-compatible dishes and fixed with paraformaldehyde for 1 h. After fixation, cells were washed twice with PBS for 5 min each. Samples were then incubated with Proteinase K solution (20 μg/mL, diluted 1:100), followed by two rinses with PBS. Subsequently, cells were treated with DNase I (10 U/mL, diluted) and washed three to four times with deionized water. Finally, samples were stained with the Alexa Fluor-488-12dUTP Labeling Mix and analyzed under a fluorescence microscope (Olympus, Tokyo, Japan).

2.14. Co-immunoprecipitation (Co-IP) and mass spectrometry analysis

Whole-cell lysates were prepared in NP-40 lysis buffer (Beyotime Biotechnology) supplemented with protease inhibitors (Beyotime Biotechnology). For Co-IP, 500 μg of total protein was incubated with either 2 μg of anti-MND1 antibody (Proteintech, Wuhan, China) or an equal amount of normal rabbit IgG (BioWorld Technology, St. Louis Park, MN, USA) as a negative control, at 4 °C for 2 h with gentle rotation. Subsequently, 30 μL of Protein A agarose beads (Yeasen Biotechnology, 50% slurry in PBS, pre-washed and equilibrated) was added to each sample, followed by overnight incubation at 4 °C with rotation to capture the antibody–antigen complexes.

The bead complexes were then pelleted by centrifugation at 2500×g for 1 min at 4 °C. After three washes with cold PBS, the immunoprecipitated proteins were eluted by resuspending the beads in 60 μL of 2 × SDS-PAGE loading buffer and boiling at 100 °C for 5 min. The samples were centrifuged again (2500×g, 1 min), and the supernatants were collected as the final IP products.

Aliquots of the IP eluates were resolved by SDS-PAGE and stained with Coomassie Brilliant Blue R-250 (Yeasen Biotechnology) to visualize co-precipitated proteins. The protein band corresponding to the expected molecular weight of the MND1-associated complex was excised and subjected to in-gel trypsin digestion, followed by nano-liquid chromatography-tandem mass spectrometry (nanoLC-MS/MS) analysis at Cosmos Wisdom Biotechnology (Hangzhou, Zhejiang, China).

2.15. Immunohistochemistry (IHC)

The preparation of liver and tumor tissue sections, along with subsequent immunohistochemical staining, was performed by Servicebio (Wuhan, China). Formalin-fixed, paraffin-embedded (FFPE) tissue blocks were sectioned, and the slides were processed according to standard immunohistochemical protocols to assess protein expression and localization within the tissue architecture.

2.16. RNA extraction and sequencing

Total RNA was extracted from MND1-overexpressing and control Bel7402 cells using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) and submitted to Novogene Co., Ltd. (Beijing, China) for RNA sequencing (RNA-seq). mRNA was enriched using magnetic beads to construct cDNA libraries, which were then sequenced on the Illumina NovaSeq 6000 platform to generate paired-end reads. Subsequently, raw data were quality-filtered, and high-quality reads were aligned to the human reference genome (GRCh38/hg38). Differentially expressed genes (DEGs) were identified using the DESeq2 algorithm with the thresholds of FDR <0.05 and |log₂FC| > 1.

2.17. Bioinformatics analysis

Gene expression data from clinical liver tissue samples were normalized and subjected to bioinformatic analysis. Differential expression analysis was carried out using R software (version 4.5.2, https://www.r-project.org/). For raw RNA-seq data, differential expression analysis was performed using the DESeq2 package. For microarray data or log-transformed expression matrices (e.g., from RNA-seq after voom transformation), the limma package was applied. DEGs were defined as those satisfying |log₂(fold change)| > 1 and an adjusted P-value <0.05 using the Benjamini–Hochberg procedure to control the false discovery rate. Survival analysis was performed using the publicly available online platform GEPIA2 (Gene Expression Profiling Interactive Analysis 2; http://gepia2.cancer-pku.cn), which integrates transcriptomic and clinical data from The Cancer Genome Atlas (TCGA) and the Genotype-Tissue Expression (GTEx) projects. Kaplan–Meier overall survival (OS) curves were generated by stratifying patients into high- and low-expression groups based on the median expression level of the gene of interest. Statistical significance between survival curves was assessed using the log-rank test, with P < 0.05 considered significant.

2.18. Statistical analysis

Statistical analysis was performed using GraphPad Prism (version 8.0.2, GraphPad Software, San Diego, CA, USA). Continuous data are expressed as the mean ± standard deviation (SD) to represent variability among sample means. Statistical significance was determined using two-tailed Student’s t-tests for pairwise comparisons or one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test for multiple group comparisons. A P-value <0.05 was considered statistically significant.

3. Results

3.1. MND1 is upregulated in HCC and associated with poor prognosis

To systematically investigate the role of MND1 in HCC, we performed an integrated bioinformatics analysis combined with experimental validation. Differential expression analysis revealed that MND1 was significantly upregulated in tumor tissues (Fig. 1A and B) using our in-house database.^21,22 Further validation in the HCCDB database confirmed that MND1 levels were significantly elevated in HCC compared with adjacent non-tumor tissues (Fig. 1C). Consistent overexpression of MND1 in HCC was also observed in the analysis of the TCGA-LIHC and ICGC HCC datasets (Fig. 1D). In clinical correlation studies, Kaplan-Meier survival analysis demonstrated that HCC patients with high MND1 expression had significantly worse OS (Fig. 1E and F). Genomic analysis from TCGA revealed amplifications, mutations, and copy number alterations (CNAs) of MND1 (Fig. 1G), suggesting its potential oncogenic role.

Fig. 1.

MND1 is upregulated in HCC and associated with poor prognosis. (A, B) Expression levels of MND1 mRNA in HCC and matched normal (MN) tissues from our databases (P < 0.0001). (C) Comparison of MND1 expression levels in tumor and adjacent tissues in the HCCDB database (P < 0.01). (D) MND1 mRNA expression levels in HCC and adjacent normal tissues from the TCGA and ICGC databases (P < 0.0001). (E, F) Survival analysis of HCC patients with high or low MND1 expression. (G) MND1 showed amplifications, mutations, and copy number alterations (CNAs) in the TCGA database. (H) Relative MND1 mRNA levels in tumor and normal tissues from clinical samples, and IHC staining examining MND1 expression in normal and HCC tissues. Scale bar, 100 μm. ∗P < 0.05.

To experimentally validate these findings, we performed qRT-PCR analysis on 22 pairs of HCC and normal tissues, which confirmed the significant upregulation of MND1 mRNA in tumor tissues (Fig. 1H). IHC analysis with anti-MND1 antibody and H-score quantification showed that MND1 protein levels were significantly higher in HCC tissues than in normal liver tissues (Fig. 1H). Our integrated bioinformatics and experimental results collectively establish MND1 as a consistently overexpressed gene in HCC, closely associated with genomic instability, tumor progression, and poor prognosis. These findings highlight the potential value of MND1 as a diagnostic biomarker and therapeutic target in the clinical management of HCC.

3.2. MND1 promotes the proliferation and growth of HCC cells and inhibits apoptosis

MND1 may be involved in the malignant progression of HCC. To investigate the regulatory role of MND1 in HCC cell proliferation, we first examined its expression levels across various HCC cell lines using Western blotting and qRT-PCR (Fig. 2A). Given the high basal expression of MND1 in HCCLM3 and MHCC97H cells, these two lines were selected for subsequent knockdown experiments, with HCCLM3 cells further chosen for the construction of a stable MND1-knockout cell line. Validation results confirmed that MND1 was effectively suppressed at both the protein and mRNA levels in the stable HCCLM3 knockout cells (Fig. 2B), as well as at the protein level in the MHCC97H and HCCLM3 knockdown cells (Fig. 2B). CCK-8 assays showed that MND1 knockdown significantly reduced the proliferative capacity of both cell lines (Fig. 2C). Consistently, TUNEL staining revealed a marked increase in apoptotic cells following MND1 depletion (Fig. 2D). In addition, colony formation assays demonstrated a substantial decrease in the number of colonies formed after MND1 knockdown (Fig. 2E), indicating that MND1 promotes HCC cell proliferation. This result was further supported by EdU incorporation assays, which showed a significant reduction in EdU-positive cells upon MND1 silencing (Fig. 3A). The tumor-promoting role of MND1 was further validated in vivo. To assess the impact of MND1 knockdown on tumor growth, we established a subcutaneous xenograft tumor model by injecting HCCLM3 cells stably expressing shMND1 or shNC into the dorsal flanks of nude mice (Fig. 3B). Additionally, an orthotopic liver tumor model was generated by direct intrahepatic implantation of the same cell lines (Fig. 3C). In both models, MND1 knockdown resulted in a significant reduction in tumor volume and burden compared to the control group (Fig. 3B and C). Mice bearing shMND1-expressing tumors exhibited a significantly lower liver index (the weight ratio of the mouse liver to body) than those in the control group (Fig. 3C). IHC analysis of excised tumors revealed markedly decreased Ki-67 protein expression in the MND1 knockdown group compared to the control group (Fig. 3D), indicating reduced proliferative activity. Consistent with these findings, correlation analysis across tumor samples demonstrated a strong positive association between MND1 and Ki-67 protein levels (Fig. 3E).

Fig. 2.

MND1 promotes proliferation and inhibits apoptosis in HCC cells. (A) The relative expression of MND1 in several HCC cell lines was determined by Western blotting and qRT-PCR. (B) MND1 protein and mRNA levels were determined by Western blotting and qRT-PCR in HCCLM3 and MHCC97H cells transfected with shRNA or siRNA. (C) CCK-8 assay was used to detect cell proliferation following MND1 knockdown by siRNA or shRNA. (D) The apoptosis was detected by TUNEL assay. (E) Colony formation assays were performed to assess the cell proliferation of MND1 knockdown MHCC97H and HCCLM3 cells by siRNA. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.0001.

Fig. 3.

MND1 promotes HCC cell proliferation and tumor growth in vivo. (A) Images and statistics of EdU proliferation assay in MND1 knockdown of MHCC97H and HCCLM3 cells. Scale bar, 100 μm. (B) Representative photographs of subcutaneous xenograft tumors, with tumor volumes measured weekly (n = 6). (C) Representative photographs of orthotopic liver tumors, with the weight ratio of the mouse liver to body (liver index) calculated (n = 6). (D) Representative IHC staining images of MND1 and Ki-67 in subcutaneous tumors. Scale bar, 100 μm. (E) Correlation analysis of MND1 and Ki-67 protein expression. ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.0001.

3.3. MND1 mediates the migration and invasion of HCC cells

MND1 plays a critical role in promoting tumor proliferation and growth while suppressing apoptosis. To determine whether MND1 also regulates the migratory and invasive capacities of HCC cells, we performed functional assays following both knockdown and overexpression of MND1. Wound healing (scratch) assays showed that MND1 knockdown significantly impaired HCC cell migration (Fig. 4A). To further validate this phenotype, we established MND1-overexpressing HCC cells and confirmed overexpression efficiency by Western blotting and qRT-PCR (Fig. 4B). Transwell migration and invasion assays demonstrated that MND1-overexpressing cells exhibited markedly enhanced migratory and invasive abilities after 48 h of incubation (Fig. 4C). Consistently, scratch assays revealed accelerated wound closure in MND1-overexpressing cells (Fig. 4D), further supporting the pro-migratory role of MND1.

Fig. 4.

MND1 mediates the migration and invasion of HCC cells. (A) Cell scratch assays were performed to assess the migratory capacity of MND1 knockdown HCCLM3 cells at 24 and 48 h. (B) MND1 protein and mRNA levels were determined by Western blotting and qRT-PCR in MND1 overexpressing Bel7402 cells. (C) The Transwell assay assessed the cell migration and invasion of the Bel7402 cells following 48 h of MND1 overexpression. (D) Cell scratch assays were performed to assess the cell migratory capacity of MND1 overexpression in Bel7402 cells at 24 and 48 h. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.0001.

3.4. MND1 is associated with DNA damage in HCC cells

To investigate the molecular mechanisms underlying MND1 function in HCC, we performed RNA-seq on MND1-overexpressing HCC cells. Differential expression analysis revealed that several genes involved in DDR and cell cycle regulation, including CDK2, CDC14B, CDC25A, CDKN1A, and E2F8 (Fig. 5A), were significantly correlated with MND1 overexpression. These RNA-seq findings were validated by qRT-PCR in both MND1-overexpressing and control cells (Fig. 5B), showing consistent expression trends. To assess the functional link between MND1 and DNA damage, we examined the formation of DSB markers by immunofluorescence. MND1 knockdown led to a marked increase in nuclear foci of γH2AX and 53BP1 (Fig. 5C), indicating elevated DNA damage accumulation. Given the close interplay between DNA damage and cell cycle progression, we further analyzed cell cycle distribution in MHCC97H and Bel7402 cells. Flow cytometry revealed a significant increase in the proportion of cells in G1 phase upon MND1 depletion (Fig. 5D). In contrast, MND1 overexpression led to an opposite alteration in the cell cycle (Fig. 5D), suggesting that MND1 deficiency induces G1 arrest in HCC cells. Collectively, these results implicate MND1 in the maintenance of genomic stability through regulation of DDR and cell cycle progression.

Fig. 5.

MND1 is associated with DNA damage in HCC cells. (A) RNA-seq analysis of DNA damage repair-related genes in Bel7402 cells overexpressing MND1. Expression values are size-factor normalized counts from DESeq2. The control group is set as the reference (baseline = 1). Statistical significance was determined by DESeq2 Wald test. (B) qRT-PCR validation of DNA damage repair-related genes in MND1 overexpression and control Bel7402 cells. (C) Immunofluorescence staining of γH2AX (red) and 53BP1 (red) in MND1 knockdown cells, with DAPI counterstaining (blue). Magnification, 200×; scale bar, 100 μm. (D) Cell cycle of the MND1 knockdown, overexpression, and corresponding control groups was examined by flow cytometry, and statistical analysis was performed to determine the percentage of G1, S, and G2/M cells in each group. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.0001.

3.5. MND1 mediates DNA damage in HCC by binding to TRIM28 protein

The aforementioned studies have demonstrated that MND1 is upregulated in HCC and strongly associated with poor prognosis. Functionally, MND1 promotes HCC cell proliferation and migration, suppresses apoptosis, and facilitates cell cycle progression—its depletion leading to G1 phase arrest and increased DNA damage. To elucidate the molecular mechanisms by which MND1 drives HCC progression, we performed Co-IP using an anti-MND1 antibody, followed by mass spectrometry to identify MND1-interacting proteins. This approach revealed that MND1 physically associates with multiple cellular proteins, among which TRIM28, a well-established regulator of DDR, showed a robust interaction with MND1 (Fig. 6A). TRIM28 is a multidomain protein belonging to the tripartite motif (TRIM) family, which comprises more than 70 members in humans. It is also known as KAP1, reflecting its role as a corepressor for KRAB domain-containing zinc finger proteins. TRIM28 is frequently overexpressed in various cancers and is generally associated with poor prognosis. Upon DNA damage, phosphorylation of TRIM28 at Ser473 by ATM/ATR kinases disrupts its interaction with chromatin-modifying complexes such as NuRD and SETDB1, leading to heterochromatin relaxation and facilitating DNA repair factor access to damaged sites,²³ thereby promoting efficient DNA repair and cell survival.²⁴ In response to DNA damage, TRIM28 promotes the formation of the E2F1–histone deacetylase 1 complex and inhibits the acetylation of E2F1, preventing the expression of pro-apoptotic target genes (e.g., Apaf1, p73, and Bim) induced by E2F1.^25,26 Additionally, TRIM28 has been reported to play an important role in cancer cell metastasis through the epithelial–mesenchymal transition (EMT) pathway.27, 28, 29 Bioinformatic analysis using TCGA and our in-house database revealed that TRIM28 expression was significantly upregulated in HCC tumor tissues compared to non-tumor controls (Fig. 6B). Co-expression analysis further demonstrated a positive correlation between MND1 and TRIM28 mRNA levels (Fig. 6C). Consistently, survival data from the HCCDB database indicated that high TRIM28 expression was associated with poorer prognosis in HCC patients (Fig. 6D).

Fig. 6.

MND1 mediates DNA damage in HCC by binding to TRIM28. (A) MND1-interacting proteins were identified by mass spectrometry, with TRIM28 validated by Co-IP. (B) TRIM28 mRNA expression in TCGA, ICGC, and our databases. (C) Correlation analysis of TRIM28 and MND1 expression in in-house and public databases. (D) Survival analysis of HCC patients with high or low expression of TRIM28 in the HCCDB database. (E) TRIM28 protein levels are reduced upon siTRIM28 transfection. Representative Western blot images showing the expression of TRIM28 in siNC and siTRIM28 groups. GAPDH was used as a loading control. (F) The mRNA expression levels of DNA damage repair-related molecules in MND1 overexpressing Bel7402 cells following TRIM28 knockdown. (G) Representative pictures of MND1 overexpressing Bel7402 cells following TRIM28 knockdown by cellular immunofluorescence assay. Scale bar, 100 μm. (H) Immunohistochemical staining of TRIM28 in subcutaneous tumors. Scale bar, 100 μm. (I) After transfection with siTRIM28, the migration of MND1 overexpressing cells and control cells was detected by Transwell assay. Scale bar, 100 μm. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.0001.

To dissect the functional interplay between MND1 and TRIM28, we generated HCC cells with MND1 overexpression or in combination with TRIM28 knockdown. qRT-PCR analysis showed that the induction of DNA damage-related genes (CDC14B, CDC25A, CDKN1A, and E2F8) by MND1 overexpression was markedly elevated upon concurrent TRIM28 depletion (Fig. 6E and F).

In immunofluorescence assays, MND1-overexpressing Bel7402 cells exhibited more pronounced 53BP1 foci formation upon TRIM28 knockdown, suggesting enhanced DNA repair activity. This indicates that TRIM28 depletion exerts a functional effect similar to that of MND1 depletion (Fig. 6G). Furthermore, immunohistochemical staining of subcutaneous xenografts from previous animal experiments confirmed lower TRIM28 protein levels in the MND1-knockdown group (Fig. 6H). To further assess the role of the MND1–TRIM28 axis in tumor progression, we performed Transwell assays. Transwell migration assays revealed that TRIM28 knockdown partially reversed the pro-migratory effect of MND1 overexpression (Fig. 6I), indicating that TRIM28 is required for MND1-mediated migration. Collectively, these findings demonstrate that MND1 physically interacts with TRIM28 and relies on TRIM28 to exert its oncogenic functions in HCC, including promotion of cell proliferation, migration, and modulation of the DDR. The positive correlation between MND1 and TRIM28 expression in clinical samples, together with their functional interdependence in vitro and in vivo, positions the MND1–TRIM28 axis as a potential therapeutic target in HCC.

4. Discussion

HCC remains one of the most prevalent and lethal malignancies worldwide.³⁰ It arises from a complex interplay of multiple etiological factors, including chronic HBV or HCV infection, excessive alcohol consumption, exposure to dietary aflatoxins and parasitic infections, and inherited metabolic disorders.³⁰ In recent decades, the epidemiological landscape of HCC has shifted alongside rising living standards and changing lifestyles. Emerging risk factors, such as metabolic dysfunction-associated steatotic liver disease (MASLD), alcohol-related cirrhosis, tobacco use, obesity, type 2 diabetes mellitus, hereditary hemochromatosis, and diverse dietary exposures, have contributed to a growing global burden of HCC. Given its high incidence, frequent diagnosis at advanced stages, and poor prognosis, elucidating the underlying pathophysiological mechanisms of HCC is imperative for developing effective diagnostic biomarkers and novel therapeutic strategies.³¹

This study found that MND1 is significantly upregulated in HCC tumor tissues compared to adjacent non-tumor tissues, as demonstrated by qRT-PCR and immunohistochemical analyses. Consistently, high MND1 expression was associated with poorer OS in HCC patients. Integrated analysis of public datasets, including TCGA and ICGC, along with our own clinical samples, revealed a concordant pattern of MND1 overexpression in HCC. Subsequent in vitro and in vivo functional experiments confirmed that MND1 promotes HCC cell proliferation, migration, and invasion.

While emerging evidence has linked MND1 to tumorigenesis in various cancers, its roles in regulating the cell cycle, DDR, and tumor immune microenvironment are increasingly recognized. Our findings further establish MND1 as a critical oncogenic driver in HCC progression.^15,16,32 Collectively, these findings indicate that MND1 drives HCC progression by promoting tumor growth, migration, and invasion, thereby contributing to the poor prognosis observed in HCC patients.

DNA damage repair plays a dual role in tumorigenesis: while it safeguards genomic integrity under physiological conditions, its dysregulation in cancer cells can promote tumor survival, stemness, and resistance to genotoxic therapies such as chemotherapy and radiotherapy.^33,34 In HCC, our integrated approach, including bioinformatic analysis, RNA-seq, qRT-PCR, and immunofluorescence, demonstrates that MND1 is functionally linked to the DDR. RNA-seq data revealed significant alterations between MND1 and key DDR-associated genes, including E2F8,³⁵ CDK2,³⁶ CDC14B,³⁷ CDC25A,³⁸ and CDKN1A.³⁹

Accumulating evidence indicates that persistent DNA damage or impaired repair capacity can drive oncogenesis through multiple mechanisms, such as genomic instability due to imbalanced oncogene/tumor suppressor activity, mitochondrial dysfunction, and immune evasion. Paradoxically, once tumors are established, enhanced DNA repair pathways may be co-opted by cancer cells to maintain their malignant phenotype and withstand therapeutic stress. Our findings position MND1 as a potential modulator of this adaptive DDR in HCC. MND1 is a meiosis-specific protein that forms a stable heterodimer with HOP2. This HOP2–MND1 complex acts as a critical cofactor for the recombinases DMC1 and RAD51, facilitating their assembly on single-stranded DNA and promoting efficient homologous recombination to repair programmed double-strand breaks during meiosis.⁴⁰ Although MND1 is predominantly expressed in meiotic cells as part of the HOP2–MND1 complex that supports DMC1/RAD51-mediated homologous recombination, emerging evidence suggests that aberrant expression of MND1 in somatic cancer cells may contribute to DDR pathways.⁴¹ In HCC, we observed ectopic MND1 expression and found it associated with altered repair of DSBs. Notably, MND1 overexpression correlated with changes in the expression of key homologous recombination factors, and TRIM28 depletion enhanced 53BP1 focus formation following genotoxic stress, implying a potential role in modulating DSB repair, possibly through interactions with core recombination machinery such as RAD51. However, whether MND1 functions independently or requires HOP2 in this context remains to be determined. Further studies are needed to clarify the molecular mechanisms by which tumor-associated MND1 influences genome stability and therapeutic response.

This study employed mass spectrometry, Co-IP, and Western blotting to demonstrate that MND1 physically interacts with TRIM28 (also known as KAP1), a universal co-repressor for KRAB domain-containing zinc finger proteins (KRAB-ZFPs). TRIM28 is a well-established regulator of DDR, chromatin remodeling, and transcriptional repression, and its dysregulation has been implicated in multiple cancers, including HCC. Our data suggest that the MND1–TRIM28 interaction may modulate DNA damage repair pathways in HCC cells, thereby contributing to tumor cell proliferation and survival. However, the precise molecular mechanism by which MND1 influences TRIM28-mediated DNA repair remains to be elucidated.

Upon DSBs, TRIM28 is rapidly phosphorylated at Serine 473 by the ATM kinase. This phosphorylation triggers a conformational change that enables TRIM28 to localize to DNA damage foci and recruit chromatin-modifying complexes such as the NuRD deacetylase complex and the SETDB1 histone methyltransferase. Through these interactions, TRIM28 facilitates local chromatin relaxation and promotes efficient DNA repair.^42,43 Furthermore, phosphorylation of TRIM28 at Ser473 enhances its interaction with the transcription factor E2F1. This association represses E2F1-mediated transcriptional activation of pro-apoptotic target genes (e.g., APAF1, PUMA, and BAX), thereby attenuating apoptosis in response to DNA damage. By transiently suppressing E2F1-driven cell death, the pS473-TRIM28–E2F1 axis may facilitate DNA repair and promote short-term cell survival under genotoxic stress.^26,44 Collectively, previous studies have established that TRIM28 plays a critical role in the DDR, and emerging evidence—including our findings—demonstrates that the MND1–TRIM28 axis is strongly associated with poor prognosis in HCC. Although this interaction may enhance the survival of cancer cells under genotoxic stress, thereby supporting tumor progression, the precise mechanisms by which the MND1–TRIM28-mediated DNA repair pathway drives HCC proliferation and aggressiveness remain to be fully elucidated. Further investigation is warranted to delineate its functional contributions and therapeutic implications.

Despite these significant advances, several limitations in our study should be recognized. The prognostic value of MND1 across diverse HCC populations requires validation through large-scale, multi-center clinical cohorts with rigorous statistical analysis. The precise molecular mechanisms by which the MND1–TRIM28 complex drives tumor progression remain incompletely understood. Future studies should employ CRISPR-Cas9-mediated knockout models to minimize off-target effects and compensatory adaptations inherent in RNAi-based approaches, thereby enabling more definitive functional interrogation. The development of small-molecule inhibitors targeting the MND1–TRIM28 interaction represents a promising avenue for translational research. Progress in these areas will accelerate the transition of MND1 from a mechanistic insight to a clinically actionable target.

5. Conclusions

This study elucidates the critical role of MND1 in the progression of HCC. Integrating multi-omics analyses with functional experiments, we demonstrate that MND1 directly binds to TRIM28 to regulate the DNA damage repair pathway, thereby driving the malignant progression of HCC. Clinical analyses reveal that high MND1 expression is significantly associated with poor patient prognosis (P < 0.05), indicating its potential as a prognostic marker and a candidate molecule for risk stratification. Notably, the identification of the MND1–TRIM28 complex provides novel insights for precision therapies targeting DNA repair pathways in HCC.

In conclusion, the MND1–TRIM28 interaction represents a crucial mechanism driving HCC progression, offering a new molecular pathway and potential therapeutic targets for the treatment of liver cancer.

Authors' contributions

Cheng Wang: Investigation, Formal analysis, Writing – original draft, Data curation, Writing – review & editing. Minghua Hu: Data curation, Writing – original draft, Writing – review & editing, Investigation. Wanyu Xia: Investigation, Validation, Writing – review & editing, Data curation, Writing – original draft. Qian Zheng: Validation, Writing – review & editing. Chengfei Zhang: Methodology, Validation, Funding acquisition. Ruoyu Shi: Investigation, Writing – review & editing. Chengsong Cao: Writing – review & editing, Supervision, Project administration. Hongping Xia: Writing – review & editing, Supervision, Funding acquisition, Conceptualization. Chengyun Yao: Resources, Funding acquisition, Writing – review & editing, Supervision.

Data availability statement

All data generated or analyzed in this study are included in this article and available from the corresponding authors upon reasonable request.

AI declaration

Kimi (Moonshot AI, Kimi k1.5, https://kimi.moonshot.cn) was used solely to improve grammar and readability. All research ideas, experimental designs, data analyses, and conclusions were independently developed and critically reviewed by the authors, who assume full responsibility for the accuracy, originality, and integrity of the work.

Declaration of competing interest

The authors declare that there is no conflicts of interest.