HMBOX1 reverses autophagy mediated 5-fluorouracil resistance through promoting HACE1-induced ubiquitination and degradation of ATG5 in colorectal cancer

ABSTRACT

Chemotherapy remains the primary treatment for unresectable or advanced postoperative colorectal cancers. However, its effectiveness is compromised by chemoresistance, which adversely affects patient outcomes. Dysregulated macroautophagy/autophagy is a proposed mechanism behind this resistance, with ubiquitination playing a key regulatory role. In this study, we identify the transcription factor HMBOX1 (homeobox containing 1) as a critical regulator of chemoresistance in colorectal cancer. RNA sequencing revealed that HMBOX1 is downregulated in drug-resistant colorectal cancer cells and tissues, with its low expression linked to poor prognosis. An integrated analysis of genes associated with autophagy and 5-fluorouracil (5-FU) resistance was conducted, verified in the colorectal cancer tissues of patients by single-cell RNA sequencing and immunostaining. Mass-spectrometry-based proteomics and RNA sequencing were used to elucidate the underlying molecular mechanisms. Functionally, upregulation of HMBOX1 enhances the sensitivity of colorectal cancer cells to the first-line treatment with 5-FU by inhibiting autophagy. Mechanistically, HMBOX1 promotes the transcription of the E3 ubiquitin ligase HACE1, which in turn enhances ATG5 K63-ubiquitination and subsequent proteasome-mediated degradation. This results in decreased ATG5 levels, inhibiting autophagy and thus reducing 5-FU resistance in colorectal cancer cells both in vitro and in vivo. Furthermore, we confirm that HMBOX1 expression positively correlates with HACE1 expression and inversely correlates with autophagy levels in clinical colorectal cancer tissues. Our findings suggest that HMBOX1 downregulation drives 5-FU resistance through autophagy enhancement in colorectal cancer, highlighting HMBOX1 as a potential target for improving chemosensitivity and patient prognosis.Abbreviation: 3-MA: 3-methyladenine; 5-FU: 5-fluorouracil; ATG: autophagy related; CASP3: caspase 3; C-CASP3: cleaved caspase 3; C-PARP: cleaved PARP; CCK8: cell counting kit-8; ChIP: chromatin immunoprecipitation; CHX: cycloheximide; CNV: copy number variation; co-IP: co-immunoprecipitation; COAD: colorectal adenocarcinoma; CQ: chloroquine; CRC: colorectal cancer; CR: complete response; FHC: fetal human colon; GEO: Gene Expression Omnibus; HACE1: HECT domain and ankyrin repeat containing E3 ubiquitin protein ligase 1; HMBOX1: homeobox containing 1; IHC: immunohistochemistry; LC-MS/MS: liquid chromatography-tandem mass spectrometry; mIHC: multiplexed immunohistochemistry; MUT: mutant; NC: negative control; OS: overall survival; PBS: phosphate-buffered saline; PD: progressive disease; PFA: paraformaldehyde; PFS: progression-free survival; PR: partial response; qPCR: quantitative polymerase chain reaction; RAPA: rapamycin; SD: stable disease; TCGA: The Cancer Genome Atlas; TEM: transmission electron microscopy; TF: translation factor; USP22: ubiquitin specific peptidase 22; WT: wild type.

Introduction

Colorectal cancer (CRC) ranks as one of the most prevalent malignancies and is a leading cause of cancer-related deaths globally, primarily due to metastasis and relapse [1]. One of the major challenges in treating CRC is the development of resistance to chemotherapy, particularly 5-fluorouracil (5-FU), a first-line chemotherapeutic agent [2,3]. The emergence of chemoresistance to 5-FU significantly limits its therapeutic effectiveness and can result in treatment failure.

Recent studies have extensively explored the role of autophagy in contributing to chemoresistance in tumors [4,5]. Autophagosome formation includes two trimeric ATG16L1 complexes ATG12–ATG5-ATG16L1 and ATG16L1-SQSTM1-LC3-II [6]. It has been observed that autophagy can function as a double-edged sword in cancer: it can suppress tumorigenesis in early stages, but also support tumor cell survival under therapeutic stress, leading to chemoresistance [4,7]. Inhibiting autophagy has shown promise in overcoming or reversing 5-FU resistance in various cancer types, indicating that targeting autophagy could be a viable strategy to enhance chemotherapeutic efficacy [8,9].

Previous research highlights the critical role of E3 ubiquitin ligases in regulating autophagy [10]. For instance, USP22 (ubiquitin specific peptidase 22) enhances ATG5-mediated macroautophagy by stabilizing ATG5 through reduced K27- and K48-linked ubiquitination at the Lys118 site [11]. Similarly, RING-type E3 ligase TRIM7 promotes autophagosome accumulation by ubiquitinating ATG7 at K413 [12]. Autophagy induction leads to MTOR inactivation, which facilitates ULK1 interaction with TRAF6, an E3 ligase that catalyzes K63-linked ubiquitination, thereby stabilizing ULK1 [10]. The E3 ubiquitin ligase HACE1 (HECT domain and ankyrin repeat containing E3 ubiquitin protein ligase 1) is known for its tumor suppressive and autophagy-regulating functions [13,14]. HACE1’s Lys48-linked polyubiquitin chains on OPTN primarily target OPTN for autophagic degradation [15]. Furthermore, HACE1 has been implicated in modulating tumor proliferation and chemoresistance through the ubiquitin-proteasome pathway [16–20]. Although ubiquitination-mediated proteasomal degradation of ATG5 has been reported [11], whether there are other E3s in existence for ATG5 ubiquitination or other consequences of ATG5 ubiquitination aside from degradation is still unknown. Here, we performed an affinity-isolation assay and liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis using a CRC cell line and identified HACE1 as a binding partner of ATG5, but the significance of this interaction is unknown.

Transcription factor (TF) HMBOX1 (homeobox containing 1) belongs to the HNF gene class of the homeobox gene family [21]. Recent studies have revealed that HMBOX1 plays dual roles in tumor progression, acting as both a tumor suppressor and promoter depending on the specific cancer type and its microenvironment [22]. For instance, HMBOX1 expression is notably decreased in liver cancer [23] and ovarian cancer [24], where it inhibits tumor progression by regulating apoptosis and autophagy. Conversely, HMBOX1 is upregulated in renal clear cell carcinoma, lung squamous cell carcinoma [25] and gastric cancer [26], where it may contribute to tumor growth and metastasis. Despite these findings, the function of HMBOX1 in colorectal cancer remains underexplored.

Here, we investigated the role of HMBOX1 in CRC, particularly its influence on autophagy and 5-FU resistance. Our study demonstrated that HMBOX1 is significantly downregulated in 5-FU resistant CRC cells and tissues. Through various in vitro and in vivo assays, we revealed that HMBOX1 upregulation enhances the chemosensitivity of CRC cells to 5-FU by inhibiting autophagy through the promotion of HACE1-mediated ATG5 degradation. These findings suggest that HMBOX1 could be a novel therapeutic target for overcoming 5-FU resistance in CRC.

Results

5-FU activated autophagy in colorectal cancers

5-Fluorouracil is commonly used in chemotherapeutic regimens for colorectal cancer (CRC). However, acquired chemoresistance seriously affects the curative effect in CRC patients, and the mechanism is still unclear. Previous studies have revealed the effects of autophagy on enhancing cancer cell chemoresistance to 5-FU in CRC [2]. In order to further investigate the relationship between 5-FU and autophagy, we first treated CRC cell line HCT116 with 5-FU at different time points (0, 4, 8, 12, 16, 20, 24, and 48 h). The western blot analysis showed that LC3B-II:LC3B-I was increased while SQSTM1 was decreased in HCT116 cells treated with 5-FU over time (Figure 1A). Next, we treated cells with 5-FU combined with chloroquine (CQ), a classic autophagy inhibitor, to inhibit autophagic flux by decreasing the fusion of autophagosomes and lysosomes [27], leading to the accumulation of LC3B-II. The results showed that following 5-FU treatment, a significant accumulation of LC3B-II was observed (Figure 1B). Additionally, the accumulation of the autophagy receptor and substrate SQSTM1/p62 after CQ treatment further indicated that 5-FU activates autophagy, which can be inhibited by CQ (Figure 1B).

Figure 1.

5-FU treatment activated autophagy in colorectal cancer cells. (A) Western blotting performed on the HCT116 cells to evaluate the expression levels of LC3B-I, LC3B-II and SQSTM1 after being treated with 5-FU (20 µm) over time. (B) Western blotting performed on the HCT116 cells to evaluate the expression levels of LC3B-I and LC3B-II after being treated with 5-FU (20 µm) combined with or without CQ (20 µm). (C) cell viability of the HCT116 and HCT116/R cells 24 h after treatment with 5-FU combined with or without CQ (20 µm) detected by CCK-8 assay. (D) Representative TEM images of the HCT116, HCT116/R cells. The red arrows indicated the autophagosomes in the cytoplasm. The graph on the right summarized the numbers of autophagosomes in the cytoplasm of different cells. (E) Western blotting performed on the HCT116 and HCT116/R cells 24 h after treatment with 5-FU combined with or without CQ (20 µm) against LC3B-I, LC3B-II and SQSTM1. (F) HCT116/R cells exhibit a greater accumulation of GFP-LC3B puncta compared to normal HCT116 cells. Immunofluorescence (IF) staining was used to analyze GFP-LC3B puncta in HCT116 cells. Quantitative analysis of GFP-LC3B puncta is presented in the right panel. Scale bars: 20 µm (n = 3 independent experiments). (G) clustered heatmap of top 20 differential expressed genes (fold change > 2) in the HCT116 and HCT116/R cells. (H) volcano plot showed the differential expression of RNAs in HCT116 and FHC cells (log2 fold change > 1, adjusted p-value <0.05). Differentially expressed RNAs are highlighted in red (upregulated in FHC cells) or blue (downregulated in HCT116 cells). (I) venn diagram summarized differentially expressed RNAs in the FHC upregulation cohort and HCT116/R downregulation cohort. Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, vs. The relative control.

To explore the link between autophagy and chemoresistance, we developed HCT116 cells with secondary resistance to 5-FU, designated HCT116/R. CCK-8 assays revealed that HCT116/R cells had higher viability compared to parental HCT116 cells when treated with increasing concentrations of 5-FU (Figure 1C), confirming successful 5-FU resistance development. Additionally, HCT116/R cells treated with CQ showed lower viability than untreated HCT116/R cells (Figure 1C). Transmission electron microscopy (TEM) images showed an increased number of autophagosomes in HCT116/R cells, indicating active autophagy (Figure 1D). Western blotting confirmed higher LC3B-II:LC3B-I ratios and decreased SQSTM1 in HCT116/R cells, further demonstrating activated autophagy, which was inhibited by CQ treatment (Figure 1E). Further results show that HCT116/R cells exhibit a significantly higher accumulation of GFP-LC3B puncta compared to normal HCT116 cells, as determined by immunofluorescence staining and quantitative analysis (Figure 1F). Fluorescence microscopy of GFPmCherry-LC3-transfected CRC cells showed more yellow (GFP⁺ RFP⁺) and red (GFP^– RFP⁺) puncta in HCT116/R cells compared to parental HCT116 cells, indicating increased autophagy activity (Figure S1A). These results support that autophagy activation in colorectal cancer cells is closely associated with 5-FU resistance.

HMBOX1 was highly downregulated in 5-FU resistant colorectal cancer cells and low expression of HMBOX1 was associated with the poor prognosis of colorectal cancer

We performed RNA sequencing on HCT116/R and normal HCT116 cells (Figure 1G), and paired RNA sequencing on normal colonic epithelial FHC cells and HCT116 cells (Figure 1H). To identify genes associated with 5-FU resistance, we intersected genes overexpressed in FHC cells and downregulated in HCT116/R cells, focusing on those with a fold change cutoff of 2.0. This led us to investigate CCDC194 and HMBOX1. qPCR results showed a significant decrease in both genes’ expression with extended 5-FU treatment (0, 12, 24, and 48 h) (Figure S2A) and in HCT116/R cells compared to parental HCT116 cells (Figure S2B), suggesting their potential role in 5-FU resistance. Further analysis using TCGA data revealed HMBOX1 was downregulated in COAD (Figure S2C), while CCDC194 lacked sufficient data for differential expression. Evaluation of fresh tissue samples from 26 CRC tumors showed no significant change in CCDC194 but a notable decrease in HMBOX1 in CRC tissues compared to adjacent tissues (Figure S2D and E). Expanded validation confirmed low HMBOX1 expression in CRC tissues via PCR and western blot (Figure 2A,B). Immunohistochemistry (IHC) of 30 colorectal cancer samples revealed higher HMBOX1 expression in tumor tissues from sensitive patients versus resistant patients (Figure 2C). Kaplan – Meier survival analyses indicated that high HMBOX1 expression correlated with favorable prognosis in CRC patients (Figure 2D,E).

Figure 2.

HMBOX1 downregulation is associated with the poor prognosis of colorectal cancer. (A) qRT-PCR analyses showing the expression level of HMBOX1 in CRC tissues and paired adjacent tissues (n=70). The p value was determined by a two-tailed paired Student t test. (B) Immunoblots of HMBOX1 in 12 pairs of colorectal cancer samples and matched adjacent normal tissues. GAPDH served as the loading control. (C) IHC staining of HMBOX1 expression in 25 paired human colorectal cancer tissues, compared with their adjacent normal ones. (D and E) Kaplan–Meier analysis of overall survival (D) and progression-free survival (E) of 70 colorectal cancer patients with low versus high HMBOX1 expression. (F and G) The relative mRNA expression (F) and protein expression (G) levels of HMBOX1 in FHC and 7 cultured colorectal cancer cell lines. (H and I) Seven CRC cell lines were exposed to increasing concentrations of 5-FU from 0 µM −60 µM for 48 h to determine the IC50 values by CCK-8 assay. (J) Correlation analysis between HMBOX1 expression levels and the IC50 values of 5-FU in CRC cell lines. Data are presented as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.001, vs. the relative control.

We also evaluated the relative mRNA and protein expression levels of HMBOX1 in normal colonic epithelial cells (FHC) and seven cultured colorectal cancer cell lines. The results showed that HMBOX1 expression was significantly downregulated in all seven CRC cell lines compared to FHC, indicating that HMBOX1 May play a role in the development or progression of CRC (Figure 2F,G). The biological role of HMBOX1 in CRC was explored using a CCK-8 assay to compare the viability of HMBOX1-overexpressing and vector control HCT116 cells. Both HCT116 and 5-FU-resistant HCT116/R cells were treated with 5-FU for 24 h. The results demonstrated that HMBOX1 overexpression significantly reduced cell viability in response to 5-FU treatment, suggesting that HMBOX1 May enhance chemosensitivity in CRC cells (Figure S2F). To further explore the relationship between HMBOX1 expression and 5-FU sensitivity, seven CRC cell lines were exposed to increasing concentrations of 5-FU (0–60 µM) for 48 h, and the IC50 values were determined by CCK-8 assay (Figure 2H,I). A subsequent correlation analysis revealed a significant negative correlation between HMBOX1 expression levels and the IC50 values of 5-FU, suggesting that higher HMBOX1 expression is associated with increased 5-FU sensitivity (Figure 2J). These findings collectively highlight HMBOX1 as a potential key regulator of 5-FU sensitivity in colorectal cancer, with its expression inversely correlated with 5-FU resistance and clinical prognosis.

HMBOX1 confers resistance to 5-FU-induced apoptosis through autophagy

To further clarify the role of HMBOX1 in cell death under both basal conditions and 5-FU treatment, we conducted CCK-8 assays and flow cytometry analysis. The results showed that HMBOX1 overexpression significantly reduced cell viability in HCT116 cells under basal conditions, and this effect was further amplified upon 5-FU treatment. Conversely, HMBOX1 knockdown in HT29 cells increased cell viability under both conditions (Figure S2G). Flow cytometry further revealed that apoptosis rates were markedly elevated in HMBOX1-overexpressing HCT116 cells following 5-FU treatment compared to basal conditions. In contrast, HT29 cells with HMBOX1 knockdown exhibited reduced apoptosis rates under both conditions (Figure S2H). These findings indicate that HMBOX1 enhances apoptosis in response to 5-FU while promoting basal autophagy-induced cell death, underscoring its dual role in regulating cell death in colorectal cancer cells.

Given the critical role of autophagy in counteracting chemotherapy-induced apoptosis in colorectal cancer, we hypothesized that HMBOX1 might mediate 5-FU resistance via autophagy. To investigate this, we overexpressed HMBOX1 in HCT116 cells with low endogenous HMBOX1 levels and knocked it down in HT29 cells with high endogenous HMBOX1 expression (Figure 3A and Figure S3A). Cytotoxicity analysis showed that HMBOX1 knockdown increased IC 50 of cells to 5-FU, while HMBOX1 overexpression decreased it, and inhibition of cell autophagy with CQ efficiently neutralized the effects (Figure 3B and Figure S3B). Flow cytometry analyses confirmed that HMBOX1 knockdown conferred resistance to 5-FU treatment, while HMBOX1 overexpression increased sensitivity to 5-FU-induced apoptosis. Inhibition of autophagy almost eliminated these differences (Figure 3C). Immunoblotting demonstrated that HMBOX1 overexpression enhanced 5-FU-induced cleavage of apoptosis-related markers, including PARP and CASP3 (caspase 3), indicating increased apoptotic activity. Conversely, HMBOX1 knockdown reduced the cleavage of these markers. Notably, autophagy inhibition with CQ abolished these effects, suggesting that HMBOX1 modulates apoptosis primarily by influencing autophagy (Figure 3D).

Figure 3.

HMBOX1 confers resistance to 5-fu-induced apoptosis through autophagy. (A) qRT-pcr of HMBOX1 expression in HCT116 with stable transfection of HMBOX1 or in HT29 cells with HMBOX1 knockdown. (B) IC 50 of 5-FU in the indicated cells under combined treatment with CQ or without CQ. (C) flow cytometry analyses of HCT116 cells with HMBOX1 overexpression and HT29 cells with HMBOX1 knockdown, compared to their respective parental control cells, following treatment with indicated concentrations of 5-FU and time with or without CQ. Statistical apoptosis ratio shown in the right. Each experiment was independently repeated three times. (D) immunoblots of cleaved PARP and cleaved CASP3 expression with or without the treatment combined with CQ (10 µm). 18 µm 5-FU was used to treat HCT116 series cells and 7 µm 5-FU for HT29 series cells. GAPDH was used as an internal loading control. Statistical analyses of C-CASP3 expression and C-PARP in indicated cells. (E and F) xenograft model of HCT116 and HT29 tumor cells for tumor progression in nude mice. The mean tumor weight of the dissected tumors after five weeks of tumor growth, and the growth curves of each tumor progression were measured by tumor volume (E) and statistical analysis of their weight (F). Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.001, vs. The relative control.

The blockade of autophagy using the known inhibitor CQ or inhibition of autophagy initiation with 3-methyladenine (3-MA), a class III phosphatidylinositol 3-kinase inhibitor, showed that while these inhibitors had marginal effects on apoptosis in both HMBOX1-overexpressing and control cells (Figure S3C), they significantly sensitized HMBOX1-knockdown cells to 5-FU treatment. Specifically, the blockade of autophagy in HMBOX1 knockdown cells led to apoptosis levels comparable to those observed in their parental counterparts treated with 5-FU. Notably, in the HMBOX1-overexpressing group, autophagy inhibition further enhanced the cleavage of apoptotic markers such as cleaved caspase 3 (C-CASP3) and C-PARP, suggesting a synergistic effect of 5-FU and autophagy blockade in promoting cell death. These results indicate that HMBOX1 mediates chemoresistance by promoting autophagy, and inhibiting autophagy can partially reverse this resistance, enhancing 5-FU-induced apoptosis.

Building on our findings regarding HMBOX1’s modulation of cell death under both basal and 5-FU-treated conditions, we next evaluated its role in 5-FU sensitivity using a subcutaneous xenograft tumor model. HMBOX1 knockdown in HCT116 cells resulted in enhanced tumor growth despite 5-FU therapy, while HMBOX1 overexpression in HT29 cells sensitized tumors to 5-FU, leading to reduced tumor growth and increased apoptosis (Figure 3E,F and Figure S3D). Immunohistochemical staining confirmed these findings, showing that HMBOX1 overexpression increased markers of apoptosis (C-CASP3) and decreased markers of autophagy (LC3B), whereas HMBOX1 knockdown showed the opposite trends (Figure S3E). These results demonstrate that HMBOX1 enhances 5-FU-induced apoptosis through its modulation of autophagy, highlighting the dual role of HMBOX1 in regulating the interplay between autophagy and apoptosis, and its contribution to chemoresistance in colorectal cancer.

HMBOX1 inhibited autophagy in colorectal cancer cells

To investigate the possible relationship between HMBOX1 and macroautophagy, we examined the two typical autophagy markers, LC3B and SQSTM1, and showed that HMBOX1 knockdown decreased SQSTM1 level with 5-FU-induced stress, whereas HMBOX1 overexpression increased it in colorectal cancer cells along with the relevant LC3B-I to LC3B-II transition for autophagy progression (Figure 4A). These results suggested that HMBOX1 expression inhibits autophagy in colorectal cancer cells. To further explore this, we treated the cells with the autophagy agonist rapamycin (RAPA) and the autophagy inhibitor 3-methyladenine (3-MA). Western blotting showed that RAPA enhanced autophagy in Vector cells. While in HMBOX1-overexpression cells, RAPA had a reduced effect, indicating that HMBOX1 expression inhibits autophagy activation (Figure 4B). Similarly, 3-MA inhibited autophagy in HMBOX1-knockdown cells (Figure 4C). Transmission electron microscopy revealed that HMBOX1 overexpression significantly reduced the number of autophagosomes compared to control cells following 5-FU treatment, whereas HMBOX1 knockdown increased the number of autophagosomes, further supporting the inhibitory effect of HMBOX1 on autophagy progression (Figure 4D,E). Additionally, HMBOX1 overexpression resulted in fewer autophagosome-related puncta (GFP⁺ RFP⁺ and GFP^– RFP⁺) following 5-FU treatment, while HMBOX1 knockdown increased these puncta, reinforcing the role of HMBOX1 in modulating autophagy (Figure 4F,G). Collectively, these findings indicate that HMBOX1 inhibits autophagy progression in colorectal cancer cells.

Figure 4.

HMBOX1 inhibits autophagy in the colorectal cancer cells. (A) Expression levels of SQSTM1, LC3B-I and LC3B-II in indicated cells with or without 5-FU treatment by western blotting, with corresponding statistical analysis. (B) Expression levels of LC3B-I, LC3B-II and SQSTM1 in the NC and shHMBOX1 HT29 cells treated with RAPA (20 µm) detected by western blotting, with corresponding statistical analysis. (C) Expression levels of LC3B-I, LC3B-II and SQSTM1 in the vector and overexpression HMBOX1 hCT116 cells treated with 3-MA (2 mm) detected by western blotting, with corresponding statistical analysis. (D and E) TEM performed on the indicated HCT116 and HT29 cells treated with 5-FU. The red arrows indicated the autophagosomes in the cytoplasm. 2500X scale bar: 2 µm. 10000X scale bar: 500 nm. The graph summarized the numbers of autophagosomes in different groups (E). (F and G) Representative immunofluorescence images of the mRFP-GFP-LC3 transfected HCT116 (F) and HT29 (G) Cells 24 h after treatment with 5-FU. Scale bar: 20 µm. Numbers of the GFP⁺ RFP⁺ and GFP⁻ RFP⁺ fluorescent puncta in different treated groups were calculated in (F and G). GFP and mRFP double-positive dots (Green⁺ Red⁺; yellow in the merged images) represent autophagosomes, where LC3 is associated with the phagophore before fusion with lysosomes. mRFP single-positive dots (Green⁻ Red⁺) indicate autolysosomes, where GFP fluorescence is quenched in the acidic lysosomal environment, but mRFP fluorescence remains stable. Data are presented as mean ± SEM. **p < 0.01, ***p < 0.001, vs. The relative control.

HMBOX1 reversed the 5-FU resistance induced by autophagy through promoting degradation of ATG5

The specific mechanism that HMBOX1 regulated autophagy in colorectal cancer cells required further investigation. Combined analysis of the TCGA Provisional database and qPCR validation revealed no significant positive correlation between HMBOX1 expression and the mRNA levels of autophagy-related genes (Figure 5A). However, a negative correlation was observed between HMBOX1 and ATG5 at the protein level (Figure 5B). Consistent results were obtained in HCT116 cells subjected to a time-gradient treatment with 5-FU (Figure S4A) and in HCT116/R cells (Figure S4B and C). Immunofluorescence staining showed that HMBOX1 downregulation increased the colocalization of endogenous LC3B and ATG5 in the cytoplasm, whereas HMBOX1 overexpression produced the opposite effect (Figure 5C). Considering that HMBOX1 effectively suppresses ATG5 at the protein level in colorectal cancer cells rather than at the mRNA level or translation events, we conducted protein stability assays. Cycloheximide (CHX) chase experiments demonstrated that HMBOX1 significantly accelerated the degradation rate of ATG5 in HCT116 cells, while HMBOX1 silencing slowed the degradation rate of ATG5 in HT29 cells (Figure 5D,E). To further investigate the relationship between HMBOX1 and the proteasomal degradation of ATG5, cells were treated with the proteasome inhibitor MG132, and the ubiquitination levels of ATG5 were assessed. Ubiquitination assays confirmed increased ubiquitin-conjugated ATG5 in HMBOX1-overexpressing cells and decreased ubiquitin-conjugated ATG5 in HMBOX1-knockdown cells (Figure 5F). Immunoblotting revealed that, upon proteasomal inhibition, ATG5 expression levels in HMBOX1-overexpressing HCT116 cells and HMBOX1-knockdown HT29 cells became similar to those in the control group (Figure 5F,G). This suggests that proteasome inhibition mimics the effect of HMBOX1 knockdown. Interestingly, reciprocal co-immunoprecipitation (co-IP) indicated that HMBOX1 does not interact with ATG5 (Figure S4D). These results indicate that HMBOX1 inhibits autophagy in colorectal cancer cells by promoting the degradation of ATG5 through an indirect, ubiquitin-proteasome-mediated mechanism, rather than via direct interaction.

Figure 5.

HMBOX1 revers the chemotherapy resistance induced by autophagy through promoting degradation of ATG5. (A) relative expression of autophagy-related genes in indicated cells were verified by qRT-pcr. (B) Immunoblots of ULK1, ATG7, BECN1, ATG12, ATG5 and ATG3 in the indicated cells. (C) Representative immunofluorescence staining of LC3B puncta and colocalization of ATG5 and LC3B in HCT116 and cells by confocal analysis (left). Scale bar: 5 µm. The increasing fold of ATG5 are shown in right. (D) immunoblot analysis showing the turnover rate of ATG5 in HCT116 cells upon HMBOX1 overexpression or knockdown. Cells were treated with cycloheximide (CHX; 60 mg/mL) for indicated time. The relative abundance of remaining ATG5 protein was normalized to GAPDH and then normalized to baseline (t = 0 h) controls (E). (F and G) ubiquitination analysis of ATG5. Cells were treated with or without MG132 for 12 h. Lysates from indicated cells were immunoprecipitated with ATG5 antibody, followed by immunoblotting analysis with antibody against ubiquitin. Statistical analysis of ATG5 expression was performed with three independent experiments (G). Data are presented as mean ± SEM. ***p < 0.001, vs. The relative control.

The interaction between HMBOX1 and HACE1 promotes ATG5 ubiquitination and degradation

To explore how HMBOX1 influences ATG5 ubiquitination and degradation, we conducted transcriptome sequencing on control and HMBOX1-overexpressing HCT116 cells (Figure 6A) and an MS-based interactome study of ATG5, confirming its expression and enrichment via IP (Figure 6B ; Table S4). Our analysis revealed a positive correlation between HMBOX1 expression and both the negative regulation of the autophagy gene signature (GO_0010507) (Figure S5A) and the ubiquitin protein ligase binding gene signature (GO_0031625) (Figure S5B). Intersection of upregulated genes in HMBOX1-overexpressing cells with those identified in the ATG5-IP study highlighted MCM6, HACE1, and RPL3 as common genes (Figure 6C). Validation in HMBOX1-overexpressing and knockdown cell lines showed that only HACE1 expression decreased with HMBOX1 knockdown (Figure 6D). Further investigation revealed that HMBOX1 promotes HACE1 expression at both mRNA and protein levels (Figure 6D,E) and negatively correlates with the expression of the autophagy marker protein LC3B(Figure 6F).

Figure 6.

The interaction between HMBOX1 and HACE1 promotes ATG5 ubiquitination and degradation. (A) Hierarchical clustered heatmap displaying the differentially expressed genes between HMBOX1 overexpression HCT116 cells compare to vector cells. (B) Coomassie brilliant blue following IP assay showing the interaction between ATG5 and HACE1. The positions of ATG5 and HACE1are indicated by arrows. (C) Summary of differentially expressed candidate genes in HMBOX1 overexpression HCT116 cells upregulation cohort and ATG5-IP cohort in a venn diagram. (D) Relative expression of candidate genes from (C) In indicated cells were verified by qRT-pcr. (E) Immunoblots of HACE1 in the indicated cells. (F) Representative immunofluorescence staining of LC3B puncta and colocalization of HACE1 and LC3B in HCT116 and HT29 cells by confocal analysis (left). Scale bar: 10 µm. The increasing fold of HACE1 are shown in right. (G) Schematic representation of the HACE1 promoter. The sequences of wild type/mutant HMBOX1 binding sites are indicated. (H) Luciferase activity assays were performed in the indicated cells transfected with wild-type (HACE1-WT) or mutant-type (HACE1-mut) HACE1 promoter-reporter plasmids. (I) ChIP-qPCR analysis displays HACE1 enrichment on the promoter of HACE1. IgG indicates the negative control. Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.001, vs. The relative control.

We hypothesized that HMBOX1 regulates ATG5 stability by modulating E3 ligase expression, specifically focusing on HACE1. To investigate this, we first analyzed the HACE1 promoter region and identified potential HMBOX1 binding sites. A schematic representation of the HACE1 promoter, including wild-type and mutant HMBOX1 binding sequences, is shown (Figure 6G). Subsequently, luciferase reporter assays (Figure 6H) and ChIP (Figure 6I) confirmed HMBOX1’s transcriptional regulation of HACE1, with HMBOX1 overexpression enhancing luciferase activity of the wild-type HACE1 promoter and binding directly to the HACE1 promoter region. Furthermore, single-cell RNA-sequencing of CRC patient samples identified 18,338 cells, which were categorized into 23 clusters (Figure S5C). To further refine the analysis, key marker genes were used for cell type annotation, as illustrated by a dot plot (Figure S5D). Re-clustering of epithelial cells extracted from the initial analysis revealed 12 distinct clusters, as shown in the UMAP plot (Figure S5E). Analysis showed that all epithelial cells, even with low CNV scores, were malignant (Figure S5F and G). We used kNN imputation for differential analysis and found that HMBOX1 is positively correlated with autophagy-related genes (Figure S5I). Stratifying cancer cells by HMBOX1 expression levels and comparing HACE1 expression using the Wilcoxon test revealed that HACE1 is downregulated in colorectal adenocarcinoma (Figure S5H and J). These findings indicate that HMBOX1 directly regulates HACE1 transcription, thereby influencing ATG5 regulation in colorectal cancer.

The E3 ubiquitin ligase HACE1 promotes the ubiquitination and degradation of ATG5

The mechanism by which HACE1 mediates the ubiquitination and degradation of ATG5 remains to be fully understood.To gain deeper insight into this mechanism, we examined the effect of HACE1 overexpression on ATG5 protein stability and its degradation pathway. Overexpression of HACE1 significantly reduced ATG5 protein levels, reversible by MG132 treatment (Figure 7A), indicating HACE1 regulates ATG5 via proteasomal degradation. CHX treatment revealed that HACE1 overexpression shortens ATG5’s half-life (Figure 7B). Co-IP confirmed the interaction between HACE1 and ATG5 (Figure 7C), and immunofluorescence showed colocalization of HACE1 and ATG5 in colorectal cancer cells (Figure 7D). HACE1 has been reported to promote the ubiquitination and degradation of RAC1 and CCNC (cyclin C) [18,28–30]. Consequently, we investigated the effect of HACE1 on ATG5 ubiquitination. Results showed that overexpression of HACE1 increased ATG5 ubiquitination in HCT116 cells (Figure 7E), and in vitro assays demonstrated enhanced ATG5 ubiquitination by HACE1, which was further augmented by HMBOX1 overexpression (Figure 7F) but reduced by HMBOX1 downregulation (Figure 7G).

Figure 7.

E3 ubiquitin ligase HACE1 promotes the ubiquitination and degradation of ATG5. (A) protein expression of ATG5 in HCT116 cells expressing either the vector or FLAG-HACE1, with or without MG132 treatment. (B) immunoblot analysis illustrating the turnover rate of ATG5 in HCT116 cells upon HACE1 overexpression (left). Cells were treated with cycloheximide (CHX; 60 mg/mL) for the indicated times. The relative abundance of remaining ATG5 protein was normalized to GAPDH and then to the baseline (t = 0) controls (right). (C) Co-immunoprecipitation (co-ip) of HACE1 and ATG5 in the indicated cells. (D) colocalization of HACE1 and ATG5 in HCT116 and HT29 cells analyzed by confocal microscopy. Cells were stained with HACE1 (red) and ATG5 (green) antibodies. Merged images are shown in yellow. Scale bar: 5 µm. (E) HCT116 cells transfected with the indicated plasmids were treated with 20 µm MG132 for 12 h before immunoprecipitation (IP). (F and G) HCT116 (F) and HT29 (G) cells were transfected with the indicated plasmids and treated with 20 µm MG132 for 12 h. HA-Ub was used to co-expressed for 72 h followed by in vivo ubiquitination analyses.

It is widely accepted that K48 or K63 ubiquitination can be recognized by the proteasome. To confirm the ubiquitin linkages on the substrate protein, we transfected cells with ubiquitin single lysine mutants (Figure S6A). Notably, transfection with the Lys63-only ubiquitin mutant (UbK63R) failed to induce ATG5 ubiquitination (Figure S6B and C), indicating a specific Lys63-linkage in ATG5 ubiquitination. Collectively, these findings elucidate that HACE1 promotes the ubiquitination and degradation of ATG5 through a Lys63-linked ubiquitination pathway, with HMBOX1 modulating this process.

The HMBOX1-HACE1-ATG5 ubiquitination axis related to tumorigenicity and 5-FU resistance in vitro and in vivo

To further assess the role of HACE1 in 5-FU-induced cell death, we performed knockdown and overexpression experiments in colorectal cancer cells. Results from CCK-8 assays (Figure S6D) and flow cytometry (Figure S6E) revealed that HACE1 knockdown significantly decreased apoptosis and reduced sensitivity to 5-FU, while HACE1 overexpression enhanced both apoptosis and the response to 5-FU treatment. These findings highlight the critical role of HACE1 in modulating 5-FU-induced cell death and further confirm its involvement in the HMBOX1-HACE1-ATG5 axis.

HACE1 and ATG5 were identified as key regulators in the HMBOX1-mediated autophagy and chemoresistance pathway. To validate the autophagy-mediated 5-FU resistance involving the HMBOX1-HACE1-ATG5 axis, we performed knockdown of HACE1 or overexpression of ATG5 in HMBOX1-overexpressing cells, and conversely, knockdown of ATG5 or overexpression of HACE1 in HMBOX1-knockdown cells (Figure S7A). Silencing HACE1 or forcing ATG5 expression in HMBOX1-overexpressing HCT116 cells significantly increased the IC50 to 5-FU (Figure S7B and C), reduced apoptosis (Figure S7D), and elevated apoptosis markers like cleaved PARP and CASP3, along with increased autophagy upon 5-FU treatment (Figure S8A-D). Conversely, HACE1 overexpression or ATG5 knockdown in HMBOX1-knockdown HT29 cells promoted apoptosis, reduced autophagy, and decreased 5-FU resistance (Figure S7B-D and Figure S8A-D), highlighting the role of autophagy in HMBOX1-induced chemoresistance through HACE1 upregulation and ATG5 downregulation.

In a subcutaneous xenograft tumor model, HMBOX1 overexpression in colorectal cancer cells sensitized tumors to 5-FU, reducing tumor burden, while silencing HACE1 or overexpressing ATG5 negated this effect (Figure 8A,B and Figure S8E). HMBOX1 overexpression inhibited autophagy and promoted apoptosis in untreated tumors (Figure 8C,D). In the 5-FU-treated group, HMBOX1 overexpression with HACE1 silencing or ATG5 overexpression led to enhanced autophagy and reduced apoptosis, while combined 5-FU and HMBOX1 overexpression increased tumor cell apoptosis (Figure 8C,D). These results confirm that the HMBOX1-HACE1-ATG5 autophagy axis is critical for 5-FU resistance in colorectal cancer.

Figure 8.

The HMBOX1-HACE1-ATG5 axis related to tumorigenicity and 5-FU resistance. (A and B) chemotherapy xenograft model in nude mice. Six nude mice per group were subcutaneously injected with the indicated cells for one week, followed by treatment with 5-FU (intraperitoneal injection, 40 mg/kg) for two weeks. Mean tumor weights were quantified approximately three weeks later (B). (C) Immunoblots of HMBOX1, HACE1, ATG5, SQSTM1, LC3B-I, PARP, cleaved-parp, CASP3, and cleaved-CASP3 in the indicated tumor tissues from mice. (D) immunohistochemistry staining for autophagy marker LC3B-I and apoptosis marker cleaved-CASP3 in subcutaneous tumors from mice in (A). Scale bars: 50 µm. Data are presented as mean ± SEM. ****p < 0.001, vs. The relative control.

HMBOX1-HACE1-ATG5 ubiquitination axis related to the prognosis of colorectal cancer patients with 5-FU based treatment

Our experiments confirmed that chemotherapy resistance in colorectal cancer cells correlates with low HACE1 and high ATG5 expression, leading to autophagy activation. qPCR assays showed elevated ATG5 mRNA and reduced HACE1 mRNA in tumor tissues compared to normal tissues (Figure 9A,D). Kaplan-Meier analysis revealed that patients with low ATG5 expression (70 patients) had better overall survival (Figure 9B) and progression-free survival (Figure 9C) compared to other groups. Conversely, low HACE1 expression was associated with poorer survival outcomes (Figure 9E,F). We categorized 70 CRC patients into sensitive (44 patients) and resistant (26 patients) groups based on treatment response. qPCR results indicated elevated HMBOX1 and HACE1 mRNA in chemotherapy-sensitive tumors (Figure S9A and B), whereas ATG5 was higher in resistant tumors (Figure S9C). There was no significant correlation between HMBOX1 and ATG5 mRNA (Figure S9D, R² = 0.0427, p = 0.0859), but a strong correlation between HMBOX1 and HACE1 mRNA (Figure S9E, R² = 0.5297, p < 0.001), suggesting HMBOX1 mainly affects ATG5 at the protein level.

Figure 9.

HMBOX1-HACE1-ATG5 ubiquitylation axis related to the prognosis of colorectal cancer patients with 5-FU based treatment. (A) expression of ATG5 in fresh tumor tissues collected from colorectal cancer patients (n = 70) detected by qPCR. The p value was determined by a two-tailed paired Student t test. (B, C) Kaplan – Meier analysis of overall survival (B) or progress-free survival (C) of 70 colorectal cancer patients with low versus high ATG5 expression. (D) expression of HACE1 in fresh tumor tissues in (A) (n = 70) detected by qPCR. The p value was determined by a two-tailed paired Student t test. (E and F) Kaplan – Meier analysis of overall survival (E) or progress-free survival (F) of 70 colorectal cancer patients with low versus high HACE1 expression. (G). Representative mIHC staining of HMBOX1, HACE1, ATG5 and LC3B expressions in 30 primary colorectal cancer specimens along with the matched adjacent normal tissues. Scale bar: 100 µm. (H) Fisher’s exact test analysis of HMBOX1 protein level to that of HACE1 (p = 0.0187) or ATG5 (p = 0.0065) in colorectal cancer tissues in (G). (I) Model of HMBOX1-HACE1 axis-induced ATG5 destabilization in preventing autophagy-mediated chemoresistance, progression, and apoptosis in colorectal cancer (by figdraw). Data are presented as mean ± SEM. ****p < 0.001, vs. The relative control.

Moreover, we performed a combined analysis based on the expression levels of HMBOX1, HACE1, and ATG5, categorizing patients into five groups: HMBOX1 (High) – HACE1 (High) – ATG5 (High), HMBOX1 (High) – HACE1 (High) – ATG5 (Low), HMBOX1 (Low) – HACE1 (High) – ATG5 (High), HMBOX1 (Low) – HACE1 (Low) – ATG5 (High), and HMBOX1 (Low) – HACE1 (Low) – ATG5 (Low). Kaplan-Meier survival analysis demonstrated that patients in the HMBOX1 (High) – HACE1 (High) – ATG5 (Low) group exhibited significantly better OS and PFS compared to the other groups (Figure S9F and G). These findings suggest that high HMBOX1 and HACE1 expression, coupled with low ATG5 expression, is strongly associated with improved clinical outcomes. Multiplexed immunohistochemistry (mIHC) on 30 paraffin-embedded tumor samples showed higher ATG5 and LC3B expression and lower HMBOX1 and HACE1 levels in tumors (Figure 9G). mIHC staining intensity revealed a significant negative correlation between ATG5 and HMBOX1 (Figure 9H), and a positive correlation between HACE1 and HMBOX1 (Figure 9H). These findings suggest HMBOX1 suppresses ATG5 through HACE1-mediated ubiquitination and degradation, affecting CRC proliferation, autophagy, and 5-FU resistance (Figure 9I).

Discussion

5-FU remains widely used as the first-line treatment of colorectal cancer [2,31]. However, acquired resistance to 5-FU treatment is often with disease progression, relapse and metastasis [32], posing a significant challenge in colorectal cancer therapy. In this study, we disclose the transcription factor HMBOX1 as a novel regulator of chemoresistance in colorectal cancer through its modulation of autophagy.

As a translation factor (TF), it is rational to predict that HMBOX1 could play important roles in genetic transcription and signal transduction to regulate multiple important cellular processes. Recent studies have shown that HMBOX1 is involved in the autophagy regulation of cancer [23]. However, there are still no studies focused on its effect on chemoresistance by autophagy in colorectal cancer. In the present study, we performed RNA sequencing in 5-FU-resistant HCT116 cells and their parental cells. Combined with qPCR validation, results showed HMBOX1 to be significantly downregulated in HCT116/R cells. Importantly, we show that HMBOX1 upregulation sensitizes CRC cells to 5-FU by inhibiting autophagy. This is consistent with previous studies suggesting that HMBOX1 can act as a tumor suppressor in various cancers by regulating apoptosis and autophagy. Our findings suggest that HMBOX1 may contribute to 5-FU resistance in CRC cell lines. Specifically, the significant correlation between HMBOX1 expression levels and 5-FU IC50 values implies that HMBOX1 plays a role in modulating chemoresistance. A potential mechanism could involve HMBOX1 regulating autophagy, as autophagy is a known mediator of drug resistance in cancer cells. High levels of HMBOX1 expression might enhance autophagy flux, thereby promoting cell survival under chemotherapeutic stress. Furthermore, the role of HMBOX1 in resistance to other chemotherapeutic agents warrants further investigation. Exploring whether HMBOX1 exhibits similar regulatory effects under different treatment conditions could provide valuable insights into its broader role in chemoresistance. These studies would deepen our understanding of HMBOX1-mediated mechanisms and potentially guide the development of novel therapeutic strategies targeting HMBOX1 to overcome drug resistance in CRC.

Tumors enhance autophagy activity to survive microenvironmental stress and to facilitate proliferation and aggressiveness [7]. Recently, dysregulations in autophagy function have been proven to be a potential mechanism of developing chemoresistance [33]. Our results confirmed that autophagy was activated in the HCT116/R cells compared with their parental cells. In this study, overexpression of HMBOX1 decreased autophagy and chemotherapy resistance in colorectal cancer cells. According to bioinformatics analysis and qPCR validation, the expression of ATG5 was not significantly related to the expression of HMBOX1 in mRNA level but positively in protein level. ATG5 is one of the known ATGs, acting as an effective activator for autophagy [34–37]. Our results demonstrated that HMBOX1 interacts with ATG5 to promote its degradation through ubiquitin-mediated pathways. However, the detailed mechanisms underlying this process require further investigation. We speculate that HMBOX1 binding to ATG5 may interfere with the interaction between ATG5 and its E3 ligase or, alternatively, that HMBOX1 may interact directly with the E3 ligase, thereby preventing ATG5 from binding to it. Moreover, our study did not comprehensively assess the intracellular distribution of LC3B and ATG5 under varying HMBOX1 expression levels.Previous studies have reported nuclear localization of LC3B under specific conditions, suggesting that HMBOX1 might influence the subcellular localization of LC3B and ATG5. For example, LC3B-II and ATG5 colocalization has been observed in the nuclei of CD4-positive T cells from primary SS salivary glands under specific pathological contexts [38,39]. These findings open an intriguing avenue for exploring whether HMBOX1 regulates nuclear translocation of autophagosome-related proteins. Future experiments, such as subcellular fractionation and advanced imaging techniques, are required to confirm the precise intracellular localization of these proteins. This limitation highlights the need for further investigation to fully elucidate the role of HMBOX1 in regulating the spatial dynamics of autophagy-related proteins.

To elucidate the intricate role of ATG5 in autophagy, we utilized MS to analyze the ATG5 interactome and investigated the regulatory mechanisms through its interacting proteins. Transcriptome sequencing of HMBOX1-overexpressing cells, compared to normal cells, revealed an upregulation of the E3 ubiquitin ligase HACE1 in the HMBOX1-overexpressing group, which was found to bind to the ATG5 protein. Utilizing the JASPAR database, we identified 14 binding sites between HMBOX1 and the HACE1 promoter (Table S3). Our findings identified HACE1 as an E3 ubiquitin ligase for ATG5, uncovering the mechanisms of ATG5 ubiquitination by HACE1 and its effects on autophagy. Much of our current understanding of ubiquitination is based on studies of homotypic polyubiquitin linkages (K6, K11, K27, K29, K33, K48, K63, and Met1) [40,41]. For example, K48-linked polyubiquitin chains are involved in proteasomal degradation [42,43], whereas K63 and K11 chains serve as degradation signals for autophagy in the lysosome [44,45]. In this study, we showed that in K63 ubiquitin chain was involved in the insoluble aggregation of ATG5 (Figures S5). This is consistent with the result that MG132 treatment cannot rescue HACE1-reduced expression of ATG5 (Figure 7E,F). Our study reveals that HMBOX1 promotes the transcription of HACE1, an E3 ubiquitin ligase that has been implicated in tumor suppression and autophagy regulation. We show that HACE1 promotes the K63-ubiquitination and proteasome-mediated degradation of ATG5, thereby inhibiting autophagy. Unfortunately, we were not able to purify the aggregated ATG5 due to technical limitations. In addition, the pathologic or physiological conditions under which HACE1 exerts its effects on ATG5 ubiquitination remain to be explored.

The role of HACE1 in mediating ATG5 degradation is consistent with its known regulatory functions for many other targets. HACE1 acts as a tumor suppressor protein by recognizing and degrading several oncoproteins, such as CCNC [18], RAC1 [16,28–30], RAC2 [19], and RAC-family [19]. Thus, it is plausible that the tumor-suppressive E3 ligase HACE1 would target the potential oncogenic protein ATG5 for degradation. Our study identifies ATG5 as a new target of HACE1 E3 ligase. Although the regulation of HACE1 in colorectal cancer is not well understood, our research into the HMBOX1-HACE1-ATG5 pathway sheds light on the significant roles of this emerging tumor suppressor.

In summary, our findings reveal a connection between HMBOX1 expression, HACE1 binding, ATG5 stability, autophagy, and 5-FU resistance. The role of HMBOX1 in reducing ATG5 stability through its interaction with HACE1 provides a nuanced regulatory mechanism during cellular autophagy and contributes to 5-FU resistance. Therapeutic strategies that enhance HACE1-mediated ATG5 degradation via HMBOX1 or inhibit autophagy activity using CQ could be further developed as rational treatments for colorectal cancer patients with HMBOX1 deficiency.

Materials and methods

Human specimens

Seventy pairs of colorectal cancer tissues with the corresponding adjacent counterparts were collected from 70 pairs of CRC patients who underwent surgery (no chemotherapy or radiotherapy before surgery) at Xiangya Hospital, Central South University, Changsha, China, with approval from the Ethics Committee (2022020043). The detailed clinical features are provided in Table 1. The clinical samples were rapidly frozen in liquid nitrogen and stored at −80°C for subsequent experiments.

Table 1.

Clinical characteristics of colorectal cancer patients with low and high HMBOX1 expression.

	Group	Cases	HMBOX1 Low N = 49	HMBOX1 high N = 21	P value
Age	≤53	24	17	7	>0.9999
	>53	46	32	14
Gender	Male	34	27	7	0.1210
	Female	36	22	14
T grade	T1 + T2	16	7	9	0.0138
	T3 + T4	54	42	12
Tumor size	≤5 cm	38	27	11	>0.9999
	>5 cm	32	22	10
Stage	I-II	26	18	8	>0.9999
	III-IV	44	31	13
Distant metastasis	M0	45	29	16	0.2761
	M1	25	20	5
Lymphatic invasion	N0	31	22	9	0.9938
	N1-N3	39	27	11

Cell culture and reagents

Human colorectal cancer cell lines (LoVo, Caco2, HCT8, HCT116, SW480, SW620, HT29) and healthy fetal human colon (FHC) cells were procured from the Institutes of Biomedical Sciences/IBS (TCHu 82, TCHu 146, TCHu 18, TCHu 99, TCHu172, TCHu 101, TCHu 103, respectively) and the American Type Culture Collection/ATCC (CRL-1831). and handled according to the supplier’s recommendations. LoVo, Caco2, and FHC cells were cultured in MEM Medium (Gibco 11,095,080), while SW480 and SW620 were maintained in L-15 Medium (Gibco 11,415,064). HCT116 and HCT116 were cultured in RPMI‐1640 Medium (Gibco 12,633,020). HT29 was cultured in McCoy’s 5A Medium (Gibco 16,600,082). The culture media were supplemented with 10% fetal bovine serum (Gibco, A5670701) and 1% penicillin and streptomycin (Gibco 15,140,122). All cell lines were maintained at 37°C in a humidified atmosphere containing 5% CO₂. All used cell lines were also authenticated using the STR method and tested for mycoplasma contamination. Chloroquine diphosphate salts (CQ, 20 µM; MCE, 17589A) were dissolved in DMSO (Sigma, D8418) at 10 µM as a stock to be used, while the equal volume of DMSO was used as control. Likewise, cycloheximide (CHX, 30 µg/ml; MCE 12,320), 3-methyladenine (3-MA, 5 mm; MCE 19,312), rapamycin (RAPA, 20 nM; MCE 10,219) and MG132 (20 µM; Sigma, C2211) was also used at the indicated time respectively.

RNA sequencing

Total RNA was isolated using Trizol Reagent (Invitrogen Life Technologies 15,596,018), and its concentration, quality, and integrity were determined using a NanoDrop spectrophotometer (Thermo Fisher Scientific). Three micrograms of RNA were used for RNA sample preparation, with mRNA purified using poly-T oligo-attached magnetic beads (New England Biolabs, S1419S) and fragmented with divalent cations at elevated temperature. First-strand cDNA synthesis was performed using random oligonucleotides and SuperScript II (Invitrogen Life Technologies 18,064,014), followed by second-strand synthesis with DNA polymerase I (Thermo Fisher Scientific, EP0401) and RNase H (Thermo Fisher Scientific, EN0201). After adenylation of the 3– ends, Illumina PE adapter oligonucleotides were ligated, and cDNA fragments of 400–500 bp were selected using the AMPure XP system (Beckman Coulter). The library was enriched with a 15-cycle PCR, purified, quantified using the Agilent High Sensitivity DNA assay on a Bioanalyzer 2100 system (Agilent), and sequenced on the NovaSeq 6000 platform (Illumina) at Shanghai Personal Biotechnology Co. Ltd.

Construction of the 5-FU-resistant HCT116 cells

HCT116 cells were exposed to an initial concentration of 1 µM 5-fluorouracil (5-FU; MCE, HY-90006). After 72 h, the medium was replaced with fresh drug-free medium to allow recovery. This cycle of drug exposure followed by a recovery period was repeated, gradually increasing the concentration of 5-FU in increments of 1 µM. Each concentration level was maintained until the cells reached a stable proliferation rate comparable to untreated cells. The process continued until a final concentration of 20 µM 5-FU was reached. The resulting 5-FU-resistant HCT116 cells (HCT116/5-FU) were then maintained in medium containing 20 µM 5-FU for subsequent experiments. Resistance was confirmed by performing cell viability assays and comparing the half-maximal inhibitory concentration (IC50) of 5-FU between resistant and parental cell lines (Figure 1C).

The stable cell establishment

A lentiviral vector pHBLV-CMV-MCS-3flag-EF1-puro containing the gene encoding human HMBOX1 or ATG5 was constructed by Hanbio Biotechnology (Shanghai, China). ShRNA targeting HMBOX1, HACE1 or random sequence was synthesized and inserted into lentiviral vector pHBLV-U6-MCS-PGK-PURO of Hanbio Biotechnology. CRC cells are then transduced with appropriate lentiviruses. To screen stable transduction cells, puromycin (2 µg/mL) was used in cells resuspended for 2 weeks. The level of HMBOX1, HACE1 or ATG5 was detected by quantitative reverse transcription polymerase chain reaction (qRT-PCR) and western blot.

Plasmids construction and transfection

The siRNAs targeting ATG5 or HACE1 were constructed by RiboBio Biotech (Guangzhou, China) (Table S2). The negative control RNA duplex (NC) for siRNAs was nonhomologous to any human genome sequence. Then, CRC cells were transfected with specified plasmids and siRNAs, respectively, using Lipofectamine 3000 reagent (Invitrogen, L3000015) according to the manufacturer’s protocol.

Immunohistochemistry (IHC)

Tumor tissues were fixed in 4% paraformaldehyde and embedded in paraffin for immunohistochemical (IHC) analysis. Paraffin-embedded xenograft tumors were initially processed at 60°C for 2 h, followed by dewaxing with 100% xylene and rehydration with varying concentrations of ethanol. Subsequently, the slides were subjected to EDTA treatment at 121°C for antigen retrieval, and endogenous peroxidase activity was quenched with 3% hydrogen peroxide at room temperature. The sections were then incubated overnight at 4°C with primary antibodies, followed by incubation with corresponding secondary antibodies at room temperature for 2 h the next day. Slices were developed with Polink-1 hRP DAB Detection System (ZSGB-BIO, ZLI-9036) and Images were captured and confirmed by two independent professional pathologists by microscopy (Nikon, Tokyo, Japan). To quantify protein expression levels, color intensity was scored on a four-point scale (0, no staining; 1, weak staining; 2, moderate staining; 3, strong staining) for statistical analysis. To ensure data robustness, multiple replicates and independent tissue sections were analyzed. This approach minimizes selection bias while maintaining consistency in observations. Although we acknowledge the potential benefit of using comprehensive serial sections for all analyses, current resource constraints limited their application. Nonetheless, our methodology adheres to widely accepted standards and is consistent with previous studies in the field [46–49].

Immunofluorescence microscopy

After the required treatments, cells were washed with 1× PBS and fixed with 4% paraformaldehyde at room temperature for 15 min. Fixed cells were blocked with PBS (Gibco 10,010,023) containing 1% BSA (Biosharp, BS114) for 30 min, followed by permeabilization with 0.1% Triton X-100 (Sigma, 9036-19-5) in PBS for 10 min. Cells were then incubated with primary antibodies against LC3 or ATG5, diluted in blocking solution, at 4°C overnight. After three washes with 1× PBS, cells were stained with Alexa Fluor 488 (Abcam, ab150077)- or Alexa Fluor 594 (Abcam, ab150120)-conjugated secondary antibodies at room temperature for 1 hour in the dark. DAPI was used to counterstain nuclei for 3 min. Finally, cells were washed with PBS and mounted with an anti-fade reagent. High-resolution images were captured using a TCS SP8 confocal microscope (Leica, Wetzlar, Germany) with consistent imaging parameters, including laser intensity and exposure time, across all groups. To ensure reproducibility and transparency, raw, unprocessed images were uploaded as supplementary materials. All experiments were repeated at least three times under identical conditions to validate results and prevent stress-related artifacts in control vector cells.

Antibodies and immunoblotting

The total protein was separated by SDS-PAGE gel and transferred onto polyvinylidene fluoride membrane (PVDF, Millipore, ISEQ0010). Membranes were incubated with the following primary antibodies at 4°C overnight: anti-HA (Proteintech 51,064–2-AP), anti-His (Proteintech 66,005–1-Ig), anti-FLAG (Proteintech 66,008–4-Ig), anti-ATG12 (Cell Signaling Technology [CST], 4180), anti-ATG7 (CST, 8558), anti-ATG5 (Proteintech 10,181–2-AP), anti-ATG3 (CST, 3415), anti-BECN1 (CST, 3495), anti-HACE1 (Proteintech 24,104–1-AP), anti-ubiquitin (CST 58,395), anti-PARP (Proteintech 13,371–1-AP), anti-C-PARP (CST, 9541), anti-CASP3/caspase 3 (CST, 9662) and anti-C-CASP3 (CST, 9661), anti-SQSTM1/p62 (CST, 5114), anti-LC3B (CST, 3868), anti-LC3B (Proteintech 18,725–1-AP), anti-GAPDH (Proteintech 60,004–1-Ig), anti-HMBOX1 (Thermo Fisher Scientific, PA5–21558), anti-ULK1 (CST, 4773), normal IgG (CST, 2729). Subsequently, the PVDF membrane was incubated with the corresponding HRP labeled goat anti-mouse (Abcam, ab6728) or goat anti-rabbit IgG (Proteintech, SA00001–2) at room temperature for 1 h. ChemiDocXRS+ System (Bio-Rad) was used to visualize the protein bands. Protein band intensity was quantified and analyzed with densitometry and ImageJ software.

Autophagy assays

mRFP-GFP-LC3 (Hanbio Biotechnology, HB-AP210 0001) transfected HCT116 and HT29 cells (1 × 10⁴ cells) were seeded in confocal dishes and incubated overnight. The next day, 5-FU was added to each dish for 24 h stimulation. After treatment, the culture media was discarded, and cells were washed three times with PBS. Cells were then fixed with 1 mL of 4% PFA for 20 min and washed again three times with PBS. To assess autophagy flux, both GFP and Red LC3 dots were analyzed to distinguish autophagosomes (GFP⁺ Red⁺) from autolysosomes (Red only). Images were acquired using a TCS SP8 confocal microscope (Leica, Wetzlar, Germany). The accumulation of yellow puncta (GFP⁺ Red⁺) indicates autophagosome formation without degradation, suggesting impaired autophagic flux, while an increase in red puncta (Red only) reflects successful autolysosome degradation, indicative of active autophagic flux. The ratio of yellow to red puncta was quantified to evaluate autophagy activity under different conditions. This approach was applied to both sensitive and resistant cell lines to analyze differences in autophagy flux [50].

Transmission electron microscopy (TEM)

Cells were fixed with 2.5% glutaraldehyde at 4°C for 2 h, washed three times with 0.1 M Na-cacodylate buffer (pH 7.4), and post-fixed with 1% osmium tetroxide in the same buffer for 2 h at room temperature. Dehydration was performed using a gradient of ethanol (50, 70, 80, 90, 95, and 100%) and 100% acetone for 15 min each. Cells were then permeabilized with SPI-Pon 812 epoxy resin monomer (SPI Supplies 25,068-38-6) and acetone (1:1 for 4 h, 1:2 overnight), followed by fixation with SPI-Pon 812 for 5–8 h. Embedding was done in SPI-Pon 812, incubated at 37°C overnight and at 68°C for 48 h. Samples were stained with uranyl acetate and lead citrate for 15 min, dried overnight at room temperature, and imaged using a TEM HT7700 (Hitachi, Japan). Autophagosomes were identified based on their characteristic double-membrane structure containing electron-dense material, as described in established literature [51–54]. Quantification of autophagosomes was performed by counting the number of these structures per field of view under high magnification, ensuring consistency with previously published methods. To minimize bias and potential misidentification, ambiguous structures, such as those resembling lysosomes or multivesicular bodies, were excluded from the analysis. The arrows in the TEM images indicate representative autophagosomes identified according to these criteria.

Co-immunoprecipitation assay and MS analysis

Cell lysates were prepared and incubated overnight at 4°C with 30 µL of antibody or IgG conjugated to protein G magnetic beads (Thermo Fisher Scientific, 10003D). The beads were then washed six times with washing buffer (25 mm HEPES, pH 7.4, 150 mm NaCl, 0.5% NP-40 [Thermo Fisher Scientific 28,324], 1 mm EDTA, 1 mm PMSF [Thermo Fisher Scientific 36,978]). After washing, the beads were resuspended in 30 µL of 1 M glycine (pH 3.0) to elute the bound proteins. The eluted proteins were subjected to electrophoresis and subsequent immunoblot analysis. Separated protein bands were stained using Coomassie Brilliant Blue, and specific bands were excised and submitted for mass spectrometry (MS) analysis, performed by a commercial service provider.

Chromatin immunoprecipitation (ChIP)

Chromatin immunoprecipitation (ChIP) assays were conducted using a ChIP Assay Kit (Merck, 17–10085) according to the manufacturer’s instructions. Briefly, cells were crosslinked with 1% formaldehyde at room temperature for 10 min to preserve protein-DNA interactions, and the crosslinking reaction was quenched with 125 mm glycine for 5 min. Cells were then lysed, and the chromatin was sheared to an average size of 200–1000 bp using sonication. The sheared chromatin was incubated overnight at 4°C with anti-HMBOX1 antibody and protein G beads at 4°C overnight. After extensive washing to remove nonspecifically bound chromatin, the immunoprecipitated complexes were eluted from the beads, and crosslinks were reversed by heating at 65°C for 4 hours. The DNA was then purified using spin columns provided in the kit. The recovered DNA was analyzed by quantitative PCR using specific primers to amplify regions of interest. Results were normalized to input DNA and expressed as fold enrichment over the IgG control. The primer sequences for the HACE1 promoter are listed in Supplementary Table S1.

Dual-luciferase reporter assay

The binding sites of HMBOX1 with HACE1 were separately cloned into the luciferase reporter vector (OBiO Technology, HY-PL-000104). To validate the interaction between HMBOX1 and HACE1, co-transfections of dual luciferase reporter plasmid were performed. After co-incubation for 2 days, the fluorescence enzyme activity of luciferase was measured using the Dual-Luciferase Reporter Assay System (Promega, E1910).

Ubiquitination and Half-life analysis of endogenous ATG5

To detect the ubiquitination of endogenous ATG5, CRC cells were treated with 20 µM MG132 for 9 h to inhibit proteasome-mediated degradation. Following treatment, ATG5 was immunoprecipitated using a specific antibody and subsequently subjected to immunoblotting with an antibody against ubiquitin. For the analysis of ATG5 protein half-life, cells were treated with 30 µg/mL cycloheximide for 0, 3, 6, and 9 h. At each time point, cells were harvested, and ATG5 protein levels were assessed by immunoblotting. The relative protein levels were quantified by densitometry analysis to determine the degradation rate over time.

RNA extraction, cDNA synthesis, and real-time quantitative PCR

Total RNA was isolated using the TRIzol Reagent (Invitrogen, 15596018CN). For cDNA synthesis, 1 µg of total RNA per sample was reverse transcribed using the Prime Script Kit (Accurate Biology, AG11705) following the manufacturer’s protocol. Real-time PCR was performed in triplicate using a SYBR Green fluorescent-based assay (Accurate Biology, AG11701) on a ViiA™ 7 RT-PCR system (Applied Biosystems, Carlsbad, CA, USA). The specific primers used for real-time PCR are detailed in Table S1. Relative mRNA expression levels were calculated using the 2^−(ΔΔCt) method, normalized to GAPDH as the internal control. The calculations were as follows: ΔCt = Ct (target gene) – Ct (GAPDH) and ΔΔCt = ΔCt (case) – ΔCt (control).

Cell counting kit-8 (CCK8) assay

To assess cell viability, CRC cells were seeded into 96-well plates in triplicate (6000 cells per well) and cultured overnight. After incubation, 10 µL of CCK-8 solution was added to each well, followed by a 2-h incubation according to the manufacturer’s instructions (FUDE, FD3788). Absorbance was measured at 450 nm using a microplate reader. Each assay was performed in triplicate to ensure reproducibility and accuracy of the results.

Apoptosis analysis

Cell apoptosis was assessed using the Annexin V-FITC Apoptosis Detection Kit I (BD Pharmingen 556,547) following the manufacturer’s instructions. Briefly, cells were resuspended in 1× Binding Buffer and incubated with 5 µL ANXA5/annexin V-FITC reagent and 5 µL propidium iodide reagent in the dark at room temperature for 15 min. Apoptosis analysis was performed using the FACS Canto II flow cytometer (BD Biosciences) within 1 hour of staining.

Mice

Subcutaneous xenograft models were established in accordance with ethical guidelines. CRC cells (5 × 10⁶ per mouse) were subcutaneously implanted into the right flank regions of female BALB/c nude mice (4–5 weeks old, 6 mice per group). Tumor volumes were measured every 3 days using the formula: V (mm³) = (L × W²) × 0.5, where L is tumor length and W is tumor width. After three weeks, the mice were euthanized, and the tumors were excised and quantified. All animal experimental protocols were approved by the Medical Laboratory Animal Care Committee of Central South University (2022020043) and conducted in compliance with the guidelines authorized by the Institutional Animal Care and Use Committee (IACUC) of Central South University.

Gene expression profiling and gene set enrichment analysis (GSEA)

Analyses of human colorectal cancer data from The Cancer Genome Atlas (TCGA) data (https://portal.gdc.cancer.gov/projects/TCGA-BRCA) and Gene Expression Omnibus (GEO) datasets. HMBOX1 expression was considered as a numeric variable. A continuous-type CLS file of the HMBOX1 expression to phenotype labels in Gene Set Enrichment Analysis (GSEA) was applied. The metric for ranking genes in GSEA was set as “Pearson”, and the other parameters were set to their default values. GSEA was performed using GSEA 2.0.9 software (http://www.broadinstitute.org/gsea/).

Multiplexed immunohistochemistry (mIHC)

The mIHC kit was purchased from AiFang Biological, and the experimental protocol was according to the manufacturer’s instructions (AiFang, AFIHC025). Staining for HMBOX1, ATG5, HACE1, and LC3B was performed using the sequence of specific primary antibody dyes to detect expression in colorectal cancer tissues, and nuclei were highlighted using DAPI. The fluorescent dyes used were TYR-520 (green) for HMBOX1 detection, TYR-570 (yellow) for LC3 detection, TYR-620 (purple) for HACE1 detection and TYR-690 (red) for ATG5 detection. Finally, the slides were observed under a multi-channel fluorescence scanner and images were collected.

Statistical analysis

Statistical analyses were conducted using GraphPad Prism 7 software. Student’s t-test was employed to compare two groups, while the ANOVA test was utilized for comparisons involving more than two groups. The relationship between HMBOX1, HACE1, or ATG5 expression and clinicopathological characteristics was assessed using the v2 test. Survival analyses were performed by generating Kaplan – Meier survival curves, which were subsequently compared using the log-rank test. Additionally, survival data were evaluated through univariate and multivariate Cox regression analyses. Bivariate correlations between variables were determined using Spearman’s rank correlation coefficients. A significance level of p < 0.05 was considered statistically significant for all analyses.

Funding Statement

This study was supported by grants from the National Natural Science Foundation of China [No. 82373275, 81974384, 82173342 & 82203015], the China Postdoctoral Science Foundation [No.2023JJ40942], three projects from the Nature Science Foundation of Hunan Province [No.2021JJ3109, 2021JJ31048, 2023JJ40942], Nature Science Foundation of Changsha [No.73201], CSCO Cancer Research Foundation [No.Y-HR2019-0182 & Y-2019Genecast-043], Natural Science Foundation [Youth Funding] of Hunan Province of China [2022JJ40458], Hunan Provincial Natural Science Foundation of China [2024JJ6662], The Youth Science Foundation of Xiangya Hospital [2023Q01] and Scientific Research Program of Hunan Provincial Health Commission [202203105261]. National Natural Science Foundation of China [No. 82403920], Nature Science Foundation of Hunan Province [No. 2024JJ6662 and 2025JJ20077], The Science and Technology Innovation Program of Hunan Province [No. 2024RC3042], the Youth Science Foundation of Xiangya Hospital [No. 2023Q01], the Postdoctoral Fellowship Program of the CPSF under grant number GZC20242044, the China Postdoctoral Science Foundation under grant number 2024M753679, and the Nature Science Foundation of Changsha [No. kq2403008].

Disclosure statement

No potential conflict of interest was reported by the author(s).

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/15548627.2025.2477443

Ethics approval and consent to participate

Human CRC tissues were collected from Xiangya Hospital of Central South University. This study was reviewed and approved by the Xiangya Hospital Medical Ethics Committee of Central South University (No. 2,022,020,043) and got the consent from all participates. All experiments involving mice were performed in compliance with the animal protocol approved by 2022020043.

Data availability statement

All other data supporting the findings of this study are present in the paper and Supplementary Information.