CUL4B-DDB1-COP1-mediated UTX downregulation promotes colorectal cancer progression

A human TMA was obtained from CRC patients who underwent radical surgery between January 2007 and November 2009 at Fudan University Shanghai Cancer Center (FUSCC), Shanghai, China. Human CRC TMA was carried out as described previously [29]. The clinicopathological characteristics of patients, such as age, gender, tumor location, histological type, tumor-node-metastasis (TNM) stage, grade, venous and nervous invasion, pretreatment serum CEA level, and survival data, were collected from the FUSCC database.

All mice were raised in a specific pathogen-free environment at the Shanghai Institute of Biochemistry and Cell Biology. Utx^f/f and Villin-Cre mice were provided by Dr. Charlie Degui Chen and Dr. Anning Lin (Shanghai Institute of Biochemistry and Cell Biology, CAS), respectively. As described previously [15], the exons 11 to 14 of Utx were deleted using a targeting vector, which disrupts the Utx H3K27 demethylase domain by triggering a frameshift mutation. Cop1^f/f mice were purchased from Laboratory Animal Center, East China Normal University. Utx^−/y or Utx^−/+ mice were generated by crossing Utx^f/f mice with Villin-Cre mice, while Utx^−/− mice were embryonic lethal. Cop1^−/+ mice were generated by crossing Cop1^f/f mice with Villin-Cre mice, while Cop1^−/− mice were embryonic lethal. All mice were maintained in a C57BL/6 background. The primers for genotyping are listed in Additional file 1: Table S1.

We combined the carcinogen azoxymethane (AOM) treatment with repeated administration of dextran sodium sulfate (DSS) in drinking water to induce colorectal tumors. wild-type (WT), Utx^−/y, and Utx^−/+ mice were used for AOM/DSS treatment. 8–10-week-old mice were injected intraperitoneally with AOM (10 mg/kg, Sigma, A5486). After one week, DSS was solubilized into the drinking water to feed mice for 5 days (2%, MP Biomedicals, 160,110). Subsequently, regular drinking water was given for 14 days. The DSS treatment was repeated for another two cycles. Body weights were recorded during the DSS treatment. The mice were sacrificed 90 days after the AOM injection, and colorectums were obtained for further analysis. Gross inspection, total tumor number, tumor burden, and tumor size were recorded. The samples were fixed as ‘‘swiss-rolls,’’ and a histopathological examination was conducted.

For drug treatment, 2 weeks after AOM/DSS treatment, WT and Utx^−/y mice were randomized into two groups. Then, the mice were treated with GSK126 (50 mg/kg, dissolved in SBE-β-CD) or vehicle control daily for additional 2 weeks before being sacrificed for analysis.

Cells were lysed using EBC lysis buffer (50 mM Tris–HCl, pH 8.0, 120 mM NaCl, 0.5% Nonidet P-40) with protease inhibitors (1:100, Selleck) and phosphatase inhibitors (1:100, Selleck). Intestinal/tumor tissues from human or mice were lysed using Minute™ Total Protein Extraction Kit (Invent Biotechnologies, Inc.) with protease inhibitors (1:100, Selleck) and phosphatase inhibitors (1:100, Selleck). The protein concentrations of lysates were determined using the Bio-Rad protein assay kit on a spectrophotometer (Thermo Scientific). An equivalent of 30 μg total lysate from each sample was used for IB. The cell lysates were incubated with anti-FLAG M2 agarose beads or anti-HA agarose beads for 2–4 h for IP. The pellets were washed with NETN buffer (20 mM Tris–HCl, pH 8.0, 100 mM NaCl, 0.5% Nonidet P-40, 1 mM EDTA) before analysis by SDS-PAGE and IB with specific antibodies.

Datasets from The Cancer Genome Atlas (TCGA) (https://​portal.​gdc.​cancer.​gov), Genotype-tissue expression (GTEx) (https://​www.​gtexportal.​org/​home/​index.​html) and the Gene Expression Omnibus (GEO, https://​www.​ncbi.​nlm.​nih.​gov/​sites/​GDSbrowser?​acc=​GDS2947) were utilized to determine COP1 mRNA expression in colon cancer, colorectal adenomas and normal colon tissues.

HCT116 cells were seeded into 96-well plates at a density of 3000 cells/well. Then, the cells were incubated with CCK-8 reagent (C0039, Beyotime Biotechnology, Shanghai, China) for 2 h, and absorbance was measured at 450 nm. For colony formation assay, HCT116 cells were seeded in the 96-well plates with 500 cells/well and incubated for 10 days. For the soft agar colony formation assay, 6-well plates were covered with lower agar (1.5 mL, 0.8%), and 500 cells were seeded in the upper agar (1.5 mL, 0.4%). After 2 weeks, the colonies were stained with 1% crystal violet (Beyotime).

The distal colorectum samples of 2-month-old WT and Utx^−/y mice (each sample mixed with three animals) were collected for ChIP-seq analysis service by Active Motif Inc. using the antibody against H3K27me3 (Active Motif). The data were deposited in the NCBI’s SRA (Accession: PRJNA752794).

The distal colorectum mRNAs from 2-month-old WT mice and Utx^−/y mice (each RNA sample mixed with three animals) were obtained for transcriptome analysis service by Sangon Biotech. The data were deposited in the NCBI’s SRA (Accession: PRJNA734666).

Total RNA was extracted from cells or tissues using TRIzol (Invitrogen) according to the manufacturer’s instructions. The cDNA was synthesized using ABScript II RT Master Mix (ABclonal Technology). Quantitative PCR was performed using a SYBR.

Kit (Takara) according to the manufacturer’s protocol. Human GAPDH or mouse β-Actin was used as an internal control. The primer sequences are listed in Additional file 1: Table S1.

The ChIP assay was performed using truChIP Chromatin Shearing Kit (Covaris) and EZ-ChIP kit (Millipore). The procedure was according to the kit instruction manual provided by the manufacturer. Primer sequences can be found in Additional file 1: Table S1.

HA-UTX and different mutants were cloned into pcDNA3.1 vector. FLAG-UTX was generated by inserting UTX cDNA into pCMV-FLAG and pRLenti-CMV-MCS-3FLAG-PGK-Puro vectors. The Flag-tagged of DET1, DDB1, RBX1, Cullin1, Cullin2, Cullin3, Cullin4A, Cullin4B, and Cullin5 were generated by inserting cDNAs into pCMV-FLAG vector. The Flag-tagged and HA-tagged coding sequence of human COP1 WT or the relevant isoforms and the mutants were inserted into the lentiviral vector pLex-MCS-CMV-puro (Addgene, USA) to generate COP1 expression plasmids. Myc-Ub was provided by Dr. Cory Ronggui Hu (Shanghai Institute of Biochemistry and Cell Biology, CAS). pLKO.1-TRC cloning vector (Addgene: 10,878) was used to express shRNAs against COP1 or UTX. Briefly, the oligos targeting COP1, UTX, CUL1 and CUL4B were cloned into the pLKO.1-TRC vector to generate shRNA constructs.

HCT116, HT-29, SW620, SW480, LoVo, DLD1, and HEK293T cells were kindly provided by Cell Bank, Chinese Academy of Science and were cultured in DMEM containing 10% FBS (Invitrogen) at 37 °C in 5% CO₂. Plasmids were transiently transfected with polyethylenimine (Sigma) or Lipofectamine 3000 (Invitrogen). Cells were transfected with siRNA using Lipofectamine 3000 (Invitrogen) according to the manufacturer’s protocol.

The siRNAs targeting indicated genes were synthesized by Biotend (Shanghai, China), and the sequences are listed in Additional file 1: Table S2.

Organoids were cultured in Advanced DMEM/F12 medium (GIBCO, 12,634–010), containing 500 ng/mL R-spondin 1 (Sino Biological, 11,083-HNAS), 100 ng/mL Noggin (Sino Biological, 50,688-M02H), 50 ng/mL EGF (Sino Biological, 50,482-MNCH), 100 ng/mL FGF-10 (Sino Biological, 10,573-HNAE), 1X HEPES (GIBCO, 15,630,080), 1X Glutamax (GIBCO, 35,050,061), 100 μg/mL Normocin (InvivoGen, ant-nr-1), 1X Gentamicin/amphotericin B (GIBCO, R01510), 1X B27 (Invitrogen, 17,504–044), 1X N2 (Invitrogen, 17,504–048), 1 mM n-Acetylcysteine (Sigma-Aldrich, A9165), 10 mM nicotinamide (Sigma-Aldrich, N0636), 0.5 μM A-83–01 (Tocris, 2939), 10 μM Y-27632 (Sigma-Aldrich, Y0503), 10 nM Gastrin, 1 μM prostaglandin E2 (Sigma-Aldrich, P6532), and 3 μM SB202190 (Sigma-Aldrich, S7067). The culture medium was refreshed every 3 days.

Organoids were harvested, passaged, and seeded in a 48-well cell culture plate. Each well contained about 100 ± 50 organoids in 20 μL Matrigel with 300 μL culture medium. Once the organoids grew to 100 μm in diameter, the organoids culture medium was replaced with 300 μL drug-containing culture medium with GSK126. The drug-containing culture medium was refreshed every 3 days. Organoids were photographed every 3 days during the 9-day drug treatment period. The size of live organoids was measured using ImageJ 1.53a (National Institutes of Health, USA).

TUNEL assay was performed with a colorimetric TUNEL apoptosis assay kit according to the manufacturer’s instructions (C1091, Beyotime).

EdU assay was conducted using BeyoClick™ EdU Cell Proliferation Kit with DAB (C0085S, Beyotime).

HCT116 cells expressing ectopic COP1, ectopic COP1 plus UTX mutant (V607A/P608A/V1205A/P1206A), and empty vector were injected into the flank of each nude mouse (1 × 10⁷ cells each mouse) to evaluate the tumorigenesis. Tumor volumes were measured and recorded every 3 days.

Primary antibodies used in this study are as follows: UTX (33510S, Cell Signaling Technology; GTX121246, GeneTex), COP1 (A300-894A, Bethyl), CUL1 (SC-17775, Santa Cruz Biotechnology), CUL2 (A5308, Abclonal), CUL3 (2759, Cell Signaling Technology), CUL4A (AB60215a, Abgent), CUL4B (A6198, Abclonal), CUL5 (SC-373822, Santa Cruz Biotechnology), EMP1 (ab230445 and ab202975, Abcam), AUTS2 (25,001–1-AP, Proteintech), Ki67 (ab15580, Abcam), EZH2 (ab191080, Abcam), VEGFC (A2556, Abclonal), Caspase-9 (A0281, Abclonal), Bcl2 (15,071, Cell Signaling Technology), DDB2 (ab181136, Abcam), DCAF7 (ab138490, Abcam), DCAF13 (ab195121, Abcam), H3K27me3 (ab192985, Abcam; A2363, Abclonal), H3 (ab1791, Abcam), P27 (610,241, BD), HA-Tag (51,064–2-AP, Proteintech; SC-7392, Santa Cruz Biotechnology), Flag-Tag (F7425-0.2MG, Sigma; F3165-1MG, Sigma), GFP (66,002–1-Ig, Proteintech), Myc-Tag (16,286–1-AP, Proteintech), α-tubulin (SC-23948, Santa Cruz Biotechnology), and Vinculin (V4505, Sigma). The chemical inhibitors MG132 (S2619), MG101 (S7386), MLN4924 (S7109), Baf-A1 (S1413), and GSK126 (S7061) were purchased from Selleck.

Statistical evaluation was conducted using SPSS 25.0 (SPSS Inc., Chicago, IL, USA) and GraphPad Prism v.8 (La Jolla, CA, USA). The experimental data were analyzed using a two-tailed Student’s t-test, one-way analysis of variance (ANOVA) with Dunnett’s multiple comparisons test, Pearson’s correlation coefficients test, and log-rank test. P < 0.05 was considered significant.

To investigate the putative role of UTX in CRC, a TMA containing 260 CRC samples was generated to examine UTX expression. Immunochemistry (IHC) staining results indicated significantly lower UTX expression in stages II and III CRC patients than in stage I (Fig. 1A and B). Specifically, low UTX expression was commonly observed in male patients (P = 0.003) and patients with advanced T stage disease (P = 0.045) (Table 1). Importantly, low UTX expression was significantly associated with poor overall survival (OS) (P = 0.048, Fig. 1C) and disease-free survival (DFS) (P = 0.019, Fig. 1D) in the current CRC cohort. Multivariable Cox regression analysis further revealed that UTX expression is an independent predictor for risk stratification of DFS in CRC patients (P = 0.037, Tables 2 and 3). Therefore, low UTX expression is associated with advanced CRC stages and unfavorable clinical outcomes, which is consistent with the tumor suppressor role of UTX reported in other cancer types [16, 17].

Next, we generated mice with specifically depleted Utx alleles in intestinal epithelial cells by crossing mice expressing Villin-Cre and Floxed Utx (Utx^f/f or Utx^f/y) mice (Additional file 2: Fig. S1A). When we crossed Utx^−/+ and Utx^−/y mice, among a total of 327 little mid-born expressing Villin-Cre, only mice with Utx^+/y; Villin-Cre (n = 103), Utx^−/+; Villin-Cre (n = 115) and Utx^−/y; Villin-Cre (n = 109) genotypes were born. This could be due to the fact that the deletion of both copies of Utx in intestinal epithelial cells results in the embryonic lethality in female mice, while the homologous Uty gene may partially compensate for Utx loss in male mice. Western blotting confirmed the efficiency of UTX depletion in male mice’s large intestines (Additional file 2: Fig. S1B). Over a 16 months period, no abnormalities were detected in the appearance and structure of the large intestine in the Utx^−/y genotype mice, where Utx was depleted in the intestinal epithelial cells (Additional file 2: Fig. S1C and D).

Subsequently, AOM was combined with DSS to induce colitis and carcinoma (Fig. 2A). IHC was applied to determine the levels of UTX and H3K27me3. Consistent with a previous finding [15], Utx deletion led to increased H3K27me3 levels in intestinal tumors (Additional file 2: Fig. S1E).

During AOM/DSS treatment, four Utx^−/y mice and one Utx^−/+ mouse while no deaths were observed in WT mice. As expected, the mice with different genotypes developed tumors located mainly in the distal to middle colorectum following AOM/DSS treatment. Utx^−/+ and Utx^−/y mice showed marginal evidence of greater body weight loss than WT mice during AOM/DSS treatment (Fig. 2B), suggesting that the loss of Utx facilitates colitis and intestinal damage. We also observed that Utx^−/+ and Utx^−/y mice had more macroscopic tumors and a higher average tumor load than WT mice (Fig. 2C–E). Moreover, Utx^−/y mice showed a more pronounced tumor load than Utx^−/+ mice. Importantly, Utx^−/y mice formed larger tumors than WT mice (P = 0.049) (Fig. 2F). And the large intestines of Utx^−/y mice exhibited higher proliferative rates as compared with WT mice, as evidenced by the Ki67 signals (Fig. 2G). These results demonstrated that loss of Utx promoted tumorigenesis in the intestinal epithelial cells, and the tumor suppressive role of Utx is dose-dependent.

In order to investigate the mechanism of UTX function in CRC, UTX-knockdown HCT116 cells were generated using two independent shRNA constructs (Fig. 3A). As expected, the depletion of UTX facilitated the proliferation of HCT116 cells, as evidenced by Cell Counting Kit-8 (CCK8) assay and anchorage-independent soft agar assay (Fig. 3B and C). Since UTX is an epigenetic enzyme regulating H3K27me3, its function is expected to involve gene-specific H3K27me3 levels and subsequent changes in transcription. Thus, we performed ChIP-seq and transcriptome analysis of normal colorectal tissues from WT and Utx knockout mice (8-weeks-old, male) to elucidate how UTX deficiency drives an oncogenic program. This analysis identified 92 genes with > 1.5-fold increment of H3K27me3 modifications and > 33.3% decrease in transcripts upon Utx depletion. Among these genes, 55 genes were further selected for a siRNA-based screening in human CRC cell line HCT116 to identify conserved target genes mediating the growth inhibitory function of human UTX based on their gene function and literature evidence. Out of the 55 genes tested in the screening, the depletion of 9 genes (comprising FRMPD1, HPSE2, EMP1, SELL, IFIT2, SLC8A1, SLC44A5, LIMCH1, and AUTS2) significantly enhanced cell proliferation. Next, we employed qPCR to verify the expression changes in these 9 genes in mice intestinal tissues and HCT116 cells upon UTX depletion or knockdown, and we validated their function in suppressing HCT116 cell proliferation. Eventually, AUTS2 and EMP1 were identified as the significant UTX target genes as their expression showed dependence on UTX depletion in both mice tissue and HCT116 cells (Fig. 3D–F). The increased H3K27me3 modifications in Emp1 and Auts2 gene locus upon Utx depletion were displayed by genomic snapshots (Additional file 3: Fig. S2A). To investigate whether Utx directly regulates Emp1 and Auts2, we performed chromatin immunoprecipitation followed by quantitative PCR (ChIP-qPCR). Consistently, Utx depletion caused significantly an increased H3K27me3 around the promoter of Emp1 and Auts2 (Additional file 3: Fig. S2B). Typically, transcriptome sequencing revealed 401 significantly downregulated and 211 significantly upregulated genes upon Utx depletion (Additional file 3: Fig. S2C). Surprisingly, gene ontology (GO) analysis reflected that most biological processes affected were associated with immune processes (Additional file 3: Fig. S2D). Knockdown of EMP1 or AUTS2 significantly enhanced HCT116 cell proliferation in vitro (Fig. S3A-E). To further strengthen the potential functional role of EMP1 and AUTS2 as UTX’s major downstream genes to regulate cell growth, re-expression of EMP1 and AUTS2 was effectuated in human colorectal cancer cell line HCT116 post-shUTX treatment. Interestingly, the overexpression of EMP1 or AUTS2 largely rescued the increased growth phenotype caused by UTX depletion in the anchorage-independent soft agar assay (Additional file 4: Fig. S3F and G). Additionally, we analyzed UTX, EMP1, and AUTS2 mRNA expression in CRC patients derived from the TCGA database, and found a positive correlation between UTX-EMP1 and UTX-AUTS2 expression levels (Additional file 4: Fig. S3H). It is worth noting that AUTS2 is a component of the PRC1 complex, which regulates the development of neuronal gene expression [30]. Although several studies reported its possible function in lymphoma or leukemia, the mechanistic role of AUTS2 in cancer remains largely unclear.

EMP1 (Epithelial Membrane Protein1) has been implicated in various cancer types and is regarded to promote cancer progression by modulating cell growth and metastasis [31‐34]. Unlike other cancer types, EMP1 has been implicated in CRC as a putative tumor suppressor, which may inhibit cell migration and promote apoptosis [35]. To explore the underlying mechanisms by which EMP1 inhibited HCT116 cell proliferation, the western blotting analysis demonstrated downregulation of EMP1 upregulated VEGFC, Bcl-2 expression and downregulated Caspase-9 expression (Additional file 4: Fig S3A), consistent with the previous studies [35, 36]. Moreover, overexpression of EMP1 partially counteracted the suppressive role of UTX knockdown on apoptosis in HCT116 cells, as indicated by TUNEL assay and western blot (Fig. 3G–I). Taken together, these results suggested that EMP1 and AUTS2 play roles in mediating colorectal tumorigenesis driven by the loss of UTX.

Since the tumor suppressor role of UTX is related to H3K27me3 demethylation activity, tumors with low UTX activity may be driven by EZH2-related gene inhibition and are therefore sensitive to EZH2 inhibitors. Previous studies indicated that loss of UTX sensitizes the cells or tissues to EZH2 inhibition in multiple cancer settings [16, 18]. To explore whether UTX loss would sensitize CRC to EZH2 inhibition, spontaneous CRC was induced in WT and Utx^−/y mice through AOM/DSS treatment, followed by administration of EZH2 inhibitor GSK126 or vehicle control for additional two weeks (Fig. 4A). Compared to vehicle control, GSK126 treatment did not affect tumor number and tumor load in the WT mice. In contrast, treatment with GSK126 resulted in significant tumor regression in Utx^−/y mice (Fig. 4B–D) This observation is consistent with the finding that GSK126 treatment decreases the levels of H3K27me3 in both WT and Utx^−/y mice (Fig. 4E). To further explore the correlation between UTX expression and sensitivity to EZH2 inhibitor, three human CRC organoids were treated with different doses of GSK126 and their growth was examined. As indicated in Fig. 4F and G, organoid #Pv24 was the most sensitive organoid to GSK126 treatment with the lowest UTX expression, while organoid #Pv11 was the most resistant organoid to GSK126 treatment with the highest UTX expression, indicating an intrinsic reverse correlation between UTX expression and sensitivity to EZH2 inhibitor in human CRC. The expression of UTX target genes EMP1 or AUTS2 at protein and mRNA levels was increased post-GSK126 treatment (Additional file 4: Fig. S3I and J). These results suggested that UTX expression determines the sensitivity of CRC to EZH2 inhibitor, and low UTX expression may serve as a biomarker to identify CRC patients who can benefit from EZH2 inhibitor therapy.

Since the downregulation of UTX promoted CRC occurrence and progression, it is crucial to understand how UTX expression is regulated in CRC cells. In order to investigate the pathway governing UTX protein stability, we observed that treatment with the proteasome inhibitor MG132 or MG101 and the neddylation inhibitor MLN4924, but not lysosomal inhibitor BafA1, resulted in a marked increase in endogenous UTX abundance (Fig. 5A). This finding indicated that UTX is degraded via the ubiquitination-proteasome system, and the Cullin-RING E3 ligase (CRL) complexes are likely involved in mediating UTX ubiquitination, as MLN4924 inhibits CRL activity, which is well-documented. We then assessed how UTX interacts with and responds to various Cullin proteins. Although UTX interacted with CUL1 and CUL4B (Additional file 5: Fig. S4A), their expression levels were elevated only upon CUL4B knockdown (Additional file 5: Fig. S4B). Subsequently, we conducted a screening by transfecting siRNAs targeting all six Cullins in HCT116 cells and found that only CUL4B knockdown upregulated UTX protein levels (Additional file 5: Fig. S4C). Furthermore, we found that endogenous UTX protein levels were increased upon depletion of CUL4B in CRC cell line SW620 (Fig. 5B) Additionally, ectopic expression of CUL4B, DDB1 and DET1, the components of the CRL4 complex, reduced UTX expression in a dose-dependent manner (Additional file 5: Fig. S4D-F). These results strongly indicate that the CUL4B-based CRL4 complex is responsible for governing UTX protein stability.

Then, we tested siRNAs targeting several substrate recognition subunits of CUL4B E3 ligases (DDB2, COP1, DCAF7, and DCAF13) in HCT116 cells and found that COP1 deficiency increased UTX protein levels (Additional file 5: Fig. S4G). Knockdown of COP1 upregulated the ectopic UTX protein levels, while overexpression of COP1 decreased ectopic UTX protein levels (Fig. 5C and D). We also discovered that COP1 splice variants interacted with UTX (Fig. 5E). Furthermore, UTX interacted with each component of the complex, including COP1, DET1, Cul4B, DDB1 and RBX1 (Additional file 5: Fig. S4H). Consistently, ectopic expression of a mutation or deletion of the COP1 RING domain (C136A/C139A), which abolishes its intrinsic E3 ubiquitin ligase activity, decreased the abundance of UTX to the same extent (Additional file 5: Fig. S4I). Importantly, the depletion of endogenous COP1 diminished the interaction between exogenous CUL4B and UTX (Fig. 5F), further confirming that COP1 is the substrate recognition subunit of the CRL4 complex that modulates UTX protein stability. Moreover, an inverse correlation was observed between UTX and COP1 proteins in CRC cell lines (Fig. 5G). Importantly, depletion of COP1 in HCT116 and LoVo cells caused a distinct increase in endogenous UTX expression and reduced H3K27me3 expression (Fig. 5H). In contrast, ectopic expression of COP1 reduced the endogenous UTX expression in SW620, HCT116, and LoVo cells without affecting the transcription of UTX gene (Fig. 5I, J and Additional file 5: Fig S4J-M). Consistently, COP1 also promoted ubiquitination of UTX transfected in HEK293T cells (Fig. 5K).

Typically, COP1 interacts with “VP” motifs of its substrates, which are termed destruction “degron” sequences [25]. Thus, we mutated each of the eight conserved “VP” motifs (VP to AA) to identify the putative COP1-degron in UTX (Fig. 5L). Strikingly, only mutation in degron 3 (V607A/P608A) or degron 7 (V1205A/P1206A) impaired COP1-mediated UTX degradation (Fig. 5M and Additional file 6: Fig S5A). Taken together, these results indicate that the CRL4-COP1 complex governs UTX stability and H3K27me3 levels in CRC cells, with COP1 recognizing specific degron sequences in UTX to facilitate its degradation.

Since COP1 negatively regulates UTX expression in intestinal tissues, we speculated that COP1 might act as an oncoprotein in CRC. Analysis of TCGA, GTEx, and GEO databases revealed that COP1 mRNA expression was upregulated in colon cancer or adenomas compared to normal colon tissues (Additional file 6: Fig. S5B and C). Immunoblotting of six pairs of CRC tumors and adjacent normal tissues confirmed that COP1 expression was upregulated in CRC tumor tissues (Additional file 6: Fig. S5D). Next, we detected the COP1 expression in paired CRC and normal (N = 104) specimens in TMA using IHC staining. The results demonstrated that COP1 expression was upregulated in CRC tissues compared to normal tissues (Fig. 6A and B). Moreover, siRNA-mediated COP1 deficiency dramatically impaired the colony formation capacity of HCT116 and LoVo cells (Fig. 6C and D). Therefore, COP1 is a putative oncoprotein in CRC tumor cells. Next, we generated mice with specifically deleted Cop1 alleles in intestinal epithelial cells by crossing flox Cop1 (Cop1^f/f) mice with Villin-Cre mice (Additional file 6: Fig. S5E). When we crossed Villin-Cre;Cop1^−/+ and Villin-Cre;Cop1^−/+ mice, Villin-Cre;Cop1^−/− mice were unavailable over the 2-year period, which could be ascribed to embryonic lethality due to the deletion of both copies of Cop1 in intestinal epithelial cells. Next, we utilized WT and Cop1^−/+ mice to generate a spontaneous CRC model following AOM/DSS treatment. Initially, eight male mice were used in each group, but one mouse in the WT group died during AOM/DSS treatment. Subsequently, we observed that Cop1^−/+ mice displayed fewer macroscopic tumors and a low average tumor load than WT mice (Fig. 6E–G). These observations further support the notion of an oncogenic role of COP1 in spontaneous CRC, consistent with the findings of elevated COP1 levels in clinical CRC cases.

To investigate the physiological role of COP1 in regulating UTX, we detected the protein expression of UTX in large intestines of WT and Cop1^−/+ mice. Reduced COP1 expression increased the protein levels of UTX in mouse intestinal tissues, as detected by both immunoblotting and IHC staining (Fig. 7A and B). Then, we stained 260 CRC tumor tissues by the IHC method to determine the protein expression of COP1 and UTX. As indicated, the COP1 signal was high in most CRC tumors and significantly correlated with lower UTX expression (r = -0.257, P < 0.001, Fig. 7C and D). These findings suggest COP1 may facilitate CRC growth via UTX degradation. To investigate this hypothesis, we performed xenograft tumorigenesis experiment using HCT116 cells that ectopically expressed HA-COP1 and Flag-UTX mutant (V607A/P608A/V1205A/P1206A) resistant to COP1-mediated degradation (Additional file 6: Fig. S5F). The ectopic expression of COP1 dramatically promoted tumorigenesis compared to parental HCT116 cells, while expression of non-degradable UTX essentially reversed the tumor-promoting effect derived from COP1 overexpression (Fig. 7E–G). Importantly, this reversal was associated with increased H3K27me3 levels upon COP1 overexpression and reduced H3K27me3 levels after overexpression of non-degradable UTX in the tumors from xenograft experiment (Fig. 7H). These results suggested that COP1-mediated UTX degradation is crucial for COP1-driven CRC tumorigenesis.