Cell culture was performed in Dulbecco’s Modified Eagle Medium (DMEM; Cat:11965092, Gibco, Grand Island, NY, USA) enriched with 10% heat-inactivated fetal calf serum, under a humidified atmosphere of 5% CO₂ at 37 °C. The human normal colon epithelial cell line (NCM640) and the human CRC cell lines (SW620, HT29, DLD-1) used in this study were all acquired from the American Type Culture Collection (ATCC, Manassas, VA, USA). Transfections of HT29 and SW620 cells were performed using Lipofectamine 2000 (Cat:11668019, Invitrogen, Carlsbad, CA, USA) following the manufacturer’s protocol. All cultures were routinely maintained with mycoplasma scavenger supplementation and regularly monitored for contamination. Cell line authentication was confirmed by short tandem repeat (STR) profiling, and all lines tested negative for mycoplasma.

Stable knockdown cell lines were generated via lentiviral transduction. In brief, cells at ~90% confluence were co-transfected with a mixture of packaging plasmids (pIP1 and pIP2, 5 µg each), the envelope plasmid pIP/VSV-G (5 µg) from the ViraPower™ Lentiviral Packaging Mix (Cat: K497500, Thermo Fisher Scientific, Waltham, MA, USA), and 5 µg of a lentiviral shRNA construct (non-targeting control pLKO.1, Cat: SHC002; shBRD4-L, TRCN0000021424; shBRD4-S, TRCN0000349782; all from Sigma-Aldrich, St. Louis, MO, USA), using 30 µL of Lipofectamine 3000 (Cat: L30000001, Invitrogen, Carlsbad, CA, USA) as per the manufacturer’s protocol. After 16 hours, the medium was refreshed. The resulting viral supernatants were harvested, concentrated, and used to transduce human CRC cell lines (HT29 and SW620) in DMEM containing 8 µg/mL polybrene (Sigma-Aldrich). Transduced cells were then selected with 1 µg/mL puromycin (Sigma-Aldrich) for 4–5 days prior to expansion and subsequent analysis. The shRNA target sequences were as follows:

shBRD4-L: 5^′-CCAACCAAAGTCAGTTCCTTC-3^′ (targeting the CTM-coding region of BRD4-L). shBRD4-S: 5^′-ATTGGACACGGACTCTTAATA-3^′ (targeting the GPA-rich domain of BRD4-S).

To generate the BRD4-S overexpression construct, the full-length coding sequence (CDS) of human BRD4-S was amplified by PCR from cDNA derived from HT29 cells. The PCR product was cloned into the pcDNA3.1(+) vector (Invitrogen) downstream of the CMV promoter, incorporating a C-terminal FLAG tag for detection. The empty pcDNA3.1(+) vector served as the control. All constructs were confirmed by Sanger sequencing.

Male BALB/c (nu/nu) nude mice, aged 6–8 weeks and weighing around 25 grams, were obtained from the Shanghai Laboratory Animal Center and housed in a pathogen-free environment under controlled temperature and humidity. HT-29 cells transduced with either control or BRD4-S shRNA were cultured in McCoy’s 5A medium at 37 °C with 5% CO₂. Cells in the logarithmic growth phase were harvested, washed, and resuspended in a serum-free medium/Matrigel mixture (1:1) at a concentration of 1 $\times$ 10⁷ cells/mL with $>$95% viability. Each mouse received a subcutaneous injection of 1 $\times$ 10⁶ cells in 100 µL into the right flank. Tumor growth was monitored three times per week by two independent investigators using digital calipers, and tumor volume was calculated as (major axis $\times$ minor axis²)/2. Mice were euthanized when tumors reached 1500 mm³ or 28 days after inoculation. Tumors were excised, weighed, bisected, and either fixed in formalin for histological examination or snap-frozen for molecular analysis. All surgical procedures were performed under general anesthesia to minimize discomfort. Anesthesia was induced with isoflurane (5% v/v, inhalation) via a precision vaporizer and maintained with ketamine (90 mg/kg, intraperitoneal injection). Animals were fasted for 12 hours prior to anesthesia to reduce aspiration risk and metabolic variability. During procedures, anesthetic depth was assessed by corneal reflexes, and body temperature was maintained at 37 °C using a heating pad. Animals were placed in a dedicated, unrestrained euthanasia chamber and exposed to a gradually increasing concentration of compressed CO₂, delivered at a flow rate of 30% of the chamber volume per minute to minimize distress. Exposure was maintained for at least one minute after the cessation of vital signs. All animal procedures received approval from the Experimental Animal Ethics Committee of The Second Affiliated Hospital of Nantong University (Approval No. P20250303-039) and adhered to the standards of the Institutional Animal Care and Use Committee (IACUC). Euthanasia was performed following a two-step protocol, which was designed to minimize animal anxiety in compliance with the American Veterinary Medical Association (AVMA) Panel on Euthanasia guidelines. Death was subsequently verified by cervical dislocation as a secondary measure.

Total RNA was isolated from target cells with Trizol (Ambion) following the manufacturer’s protocol. Subsequently, 1 µg of the extracted RNA was subjected to reverse transcription using the RevertAid First Strand cDNA Synthesis Kit (Cat: K1622, Thermo Fisher Scientific, Waltham, MA, USA). Quantitative PCR was then performed on the CFX96 Connect™ Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA) using the SYBR Green PCR Kit (Cat: 204145, Qiagen, Hilden, Germany) in 20 µL reactions. The thermal cycling conditions comprised an initial denaturation at 95 °C for 2 minutes, followed by 40 cycles of 95 °C for 10 seconds and 60 °C for 10 seconds. Primers specific for DDX27, GAPDH, BRD4-L, BRD4-S, ZFP64, RBL1, ZNF217, HNF4A, GNL3L, and POFUT1 were designed using Primer3 software (version 1.0; https://primer3.org/). The sequences were as follows: Human DDX27 F (forward): 5^′-CTCACTAAAGGCACCGAAG-3^′, R (reverse): 5^′-GGCAGAGAAGTTGCTTGTGG-3^′; Human GAPDH F (forward): 5^′-ACGGCAAGTTCAACGGCACAG-3^′, R (reverse): 5^′-GACGCCAGTAGACTCCACGACA-3^′; Human BRD4-L F (forward): 5^′-CTCCTCCTAAAAAGACGAAGA-3^′, R (reverse): 5^′-TTCGGAGTCTTCGCTGTCAGAGGAG-3^′; Human BRD4-S F (forward): 5^′-TTTCTCTCTCCCTCTACGT-3^′, R (reverse): 5^′-TTAGGCAGGACCTACGTAG-3^′; Human ZFP64 F (forward): 5^′-ATGGCTGCAGTTCTGTGTACT-3^′, R (reverse): 5^′-CTGCAGGATCTGCAGATGT-GGTA-3^′; Human RBL1 F (forward): 5^′-CGTGATGTCCGTGTACTTTGCAG-3^′, R (reverse): 5^′-GCTG-CAGTTGC-AAGATGTGCGATG-3^′; Human ZNF217 F (forward): 5^′-CAGACCTACAGCAACAGCAG-3^′, R (reverse): 5^′-GCT-GTACTTGGCCTTCATCC-3^′; Human HNF4A F (forward): 5^′-CGGACTGGGTCTGATGTGCAGTT-3^′, R (reverse): 5^′-CAGGCATTGCAGTGCCACGTAGAT-3^′; Human GNL3L F (forward): 5^′-AGCAAGATGGACGAGACCCA-3^′, R (reverse): 5^′-TCACCAGGTACTTGTAGCCG-3^′; Human POFUT1 F (forward): 5^′-CGGTTCTGCTGTGGAGATG-TGT-3^′, R (reverse): 5^′-GCATTGGTGCCAGCTGTGATGTGC-3^′; Human STAT5B F (forward): 5^′-CAGCCTACCATTGACAGCGT-3^′, R (reverse): 5^′-TGTCCAGCATCCTTGAACCA-3^′.

Cellular proteins were extracted using RIPA buffer supplemented with 1 mM PMSF. The extracts were then subjected to SDS–PAGE on 8% gels and electrophoretically transferred onto PVDF membranes (Cat: IPVH00010, Millipore, Billerica, MA, USA). Membrane blocking was performed with 5% non-fat dry milk in TBST. Primary antibody incubations were carried out overnight at 4 °C, followed by a 2-hour incubation with corresponding secondary antibodies at room temperature. Signal detection was achieved with an ECL substrate (Cat: 180-5001, Tanon, Shanghai, China), and imaging was performed using a ChemiDoc system (Bio-Rad). The following primary antibodies were used in Western blot analyses: anti-BRD4 (Cat: ab128874, Abcam, Cambridge, UK; 1:1000), anti-DDX27 (Cat: ab177950, Abcam; 1:1000), anti-SRSF6 (Cat: ab38017, Abcam; 1:1000), anti-pSRSF6 (Cat: MABE50, EMD Millipore, Billerica, MA, USA; 1:1000), anti-$\beta$-actin (Cat: ab8227, Abcam; 1:2000), and anti-GAPDH (Cat: sc-47724, Santa Cruz Biotechnology, Dallas, TX, USA; 1:1000).

Cell proliferation was evaluated by seeding transfected cells (2500 cells/well) into 96-well clear plates containing complete medium. Proliferation was quantified at designated time points using the Cell Counting Kit-8 (CCK-8; Cat: WH1199, Dojindo Laboratories, Kumamoto, Japan), and the absorbance at 450 nm was measured with a microplate reader.

For immunofluorescence staining, cells were first fixed with 4% paraformaldehyde at low temperature for 20 min and permeabilized with 0.2% Triton X-100 for 10 min. After three Phosphate-Buffered Saline (PBS) washes, non-specific sites were blocked with 1% BSA in PBS for 30 min. The samples were then incubated overnight at 4 °C with primary antibodies against DDX27 (Cat: ab177950, Abcam, 1:200) or SRSF6 (Abcam, ab38017, 1:200), followed by a 1-hour incubation at room temperature with secondary antibodies (Alexa Fluor 488 goat anti-rabbit, Invitrogen, A-11034, 1:40; or Alexa Fluor 594 goat anti-mouse, Invitrogen, A-11012, 1:40). Following additional PBS washes, nuclei were stained with DAPI for 10 min, and the samples were mounted in glycerol for visualization.

The RNA co-immunoprecipitation (RIP) assay was conducted with the Magna RIP Kit (Millipore, MA, USA) following the manufacturer’s protocol. Briefly, cells were lysed in ice-cold RIPA buffer containing protease inhibitors. For each immunoprecipitation, 500 µg of protein lysate was incubated with 40 µL of A/G beads (sc-2003, Santa Cruz Biotechnology, Dallas, TX, USA). Subsequently, 2 µg of either control IgG (Santa Cruz Biotechnology) or specific antibody against DDX27/SRSF6 (Abcam) was added, and the mixture was subjected to overnight incubation at 4 °C. An additional 40 µL of A/G beads were then introduced for a further 6-hour incubation. The beads were then washed with PBS containing 0.2% NP-40, and the bound proteins were eluted by boiling in 2 $\times$ SDS-PAGE sample buffer for 5 minutes, followed by Western blot analysis. After immunoprecipitation, reverse cross-linking was carried out, and total RNA was extracted using Trizol (5596026, Invitrogen, Carlsbad, CA, USA). The extracted RNA was treated with DNase I (AM2222, Invitrogen) to remove contaminating genomic DNA. BRD4 expression levels were quantified by qPCR on an ABI 7500 Real-Time PCR system (Applied Biosystems, Foster City, CA, USA).

Chromatin immunoprecipitation (ChIP) was carried out with the EZ-ChIP™ Kit (17-295, Millipore, Burlington, MA, USA) per the manufacturer’s protocol. In brief, HT-29 cells were fixed with 1% formaldehyde for 10 min at room temperature, and the cross-linking was terminated by the addition of glycine (0.125 M final concentration). After washing with cold PBS, the cells were harvested and lysed in SDS lysis buffer containing protease inhibitors. The chromatin was then fragmented by sonication using a Bioruptor® Pico (B01060001, Diagenode, Denville, NJ, USA) to achieve fragments ranging from 200 to 500 bp. The sheared lysates were subjected to immunoprecipitation overnight at 4 °C with 2 µg of either anti-BRD4 antibody (Abcam, ab128874) or control rabbit IgG. Protein–DNA complexes were captured with protein A/G magnetic beads and sequentially washed with low-salt, high-salt, LiCl, and TE buffers. Following cross-link reversal (65 °C for 4 h with 200 mM NaCl) and proteinase K digestion, the immunoprecipitated DNA was purified using spin columns. Quantitative PCR analysis was performed on a CFX96 Real-Time PCR System (Bio-Rad) using SYBR Green, with enrichment expressed as the percentage of input and normalized to the IgG control. The following primers were used to amplify promoter regions of ERK target genes:

CCND1 promoter

Forward: 5^′-CCTCTCGCTCCGTAACCATC-3^′

Reverse: 5^′-GCTGGACTTGACCACCTTCC-3^′

DUSP6 promoter

Forward: 5^′-GAGGCGAGAGAGAGGAAGGA-3^′

Reverse: 5^′-CCTCCTCCTCCACCTCTTTC-3^′

To assess the prognostic significance of BRD4 isoforms in CRC, mRNA expression matrices of CRC tissues and matched adjacent normal tissues were retrieved from The Cancer Genome Atlas (TCGA) database. The mRNA expression values of BRD4-L (long isoform) and BRD4-S (short isoform) were then extracted for comparative analysis. Patients were stratified into high- and low-expression groups according to the median BRD4-S expression value (or predefined clinical cutoffs where applicable). This stratification facilitated systematic evaluation of the association between BRD4-S expression levels and patient survival outcomes.

Transcriptomic data from TCGA were analyzed to identify BRD4-S-related transcriptional alterations in CRC by comparing tumor and normal tissues. Differential expression analysis, performed with DESeq2 or edgeR, identified significant genes using a $|$log₂ fold change$|$ $>$1 and adjusted p-value 0.05 cutoff. A ggplot2-generated volcano plot visualized these results, depicting log₂ fold change versus –log₁₀ (adjusted p-value), with gene expression changes color-coded (red for up, blue for down). Key BRD4-S–associated genes, including potential interactors or targets, were highlighted for subsequent investigation.

To characterize BRD4-L, BRD4-S, and DDX27 -associated genes in CRC, candidate genes were identified by integrating differentially expressed genes (DEGs) with BRD4-S targets from the GEPIA2 database. Functional enrichment analysis was performed using ClusterProfiler (Open-source R package, Bioconductor, Seattle, WA, USA) or DAVID, with Gene Ontology (GO) terms classified into biological processes, molecular functions, and cellular components. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis was also conducted, focusing on BRD4-S–related pathways such as MAPK/ERK signaling. Significant terms were defined as those with a false discovery rate (FDR) 0.05 and ranked by gene ratio. Enriched pathways and GO terms were visualized using bar and dot plots.

Statistical significance was defined as a two-sided p-value 0.05. All analyses were carried out with SPSS software (version 23.0; IBM Corporation, Armonk, NY, USA). mRNA expression differences between groups were assessed by Student’s t-test, while survival analysis was performed using the Kaplan-Meier method. Quantitative data are expressed as mean $\pm$ standard deviation (SD).

To examine the expression levels of BRD4-L (200 kDa) and BRD4-S (140 kDa) (Fig. 1A) in CRC, we first obtained mRNA expression matrices for CRC tissues and paired adjacent non-tumor tissues from the Gene Expression Profiling Interactive Analysis (GEPIA2) database. From these matrices, we extracted the mRNA expression values of BRD4-L and BRD4-S. Comparative analysis revealed that overall BRD4 expression was significantly higher in CRC tissues than in adjacent normal tissues (p 0.05). However, BRD4-L expression did not differ significantly between the two groups (p $>$ 0.05) (Fig. 1B). Notably, patients with high BRD4-S expression had significantly shorter survival times compared to those with low BRD4-S expression (p = 0.017), indicating a strong association between elevated BRD4-S levels and poor prognosis. In contrast, BRD4-L expression showed no significant correlation with survival outcomes (p = 0.24) (Fig. 1C). We next employed RT-PCR to quantify BRD4-S expression in CRC tissues and adjacent normal tissues, which confirmed that BRD4-S was markedly upregulated in tumor tissues (Fig. 1D). To further investigate its functional role in CRC cells, we analyzed BRD4-S expression using RT-qPCR and Western blotting. Our results demonstrated that BRD4-S was elevated in multiple CRC cell lines, including SW620, HT29, and DLD-1, compared with the normal control cell line NCM640 (Fig. 1E,F).

Fig. 1.

BRD4-S is highly expressed in CRC cells and tissues and related to poor prognosis. (A) Schematic representation of the domain structures of the long (BRD4-L) and short (BRD4-S) isoforms of BRD4. (B) Differential expression analysis of BRD4-L and BRD4-S transcripts in normal and malignant tissues based on the TCGA database. (C) Kaplan–Meier survival curves comparing overall survival in patients with high BRD4-L versus BRD4-S expression using TCGA data. (D) qRT-PCR analysis of BRD4-L and BRD4-S expression in normal colon tissue and CRC tissue. (E,F) qRT-PCR and western blot analyses of BRD4-L and BRD4-S expression in CRC cell lines, including NCM640, SW620, HT29, and DLD. Data are presented as mean $\pm$ SEM. No significant difference (ns) is indicated when p $>$ 0.05; *, p 0.05; **, p 0.01. BRD4-S, short isoform of bromodomain-containing protein 4; CRC, colorectal cancer; TCGA, the Cancer Genome Atlas; qRT-PCR, quantitative real-time PCR; COAD, colon adenocarcinoma.

We designed isoform-specific shRNA primers targeting the CTM domain unique to BRD4-L and the GPA domain exclusive to BRD4-S, successfully generating CRC cell lines with stable knockdown of BRD4-S or BRD4-L (Fig. 2A). To investigate isoform-specific functions of BRD4 in CRC, we performed CCK-8 and colony formation assays to evaluate the impact of BRD4-S or BRD4-L silencing on cell proliferation. Notably, BRD4-S knockdown significantly reduced both the size and number of colonies (p 0.01), whereas BRD4-L knockdown did not exert comparable inhibitory effects on clonogenicity (Fig. 2B). Consistent with these findings, BRD4-S depletion markedly suppressed the proliferation of HT29 and SW620 cells relative to the control group (sh-NC), while BRD4-L silencing produced no significant changes in proliferation (Fig. 2C). Furthermore, scratch assays revealed that BRD4-S silencing significantly impaired the migratory capacity of CRC cells, whereas BRD4-L depletion only slightly reduced migration in HT29 cells and showed no significant effect in SW620 cells compared with controls (Fig. 2D). To further validate the oncogenic role of BRD4-S, we established BRD4-S overexpression models in HT29 cells via plasmid transfection. Overexpression efficiency was confirmed by Western blotting (Supplementary Fig. 1A). In contrast to the knockdown experiments, BRD4-S overexpression significantly enhanced colony formation in HT29 cells (p 0.01) (Supplementary Fig. 1B). Similarly, CCK-8 assays demonstrated that BRD4-S overexpression markedly promoted proliferation compared with the empty vector control (p 0.01) (Supplementary Fig. 1C). Scratch assays further indicated that BRD4-S overexpression accelerated wound closure, reflecting increased migratory potential (Supplementary Fig. 1D). Our results critically support a model wherein BRD4-S, in contrast to BRD4-L, is the primary driver of CRC cell proliferation and migration.

Fig. 2.

BRD4-S promotes CRC proliferation and migration in vitro. (A) BRD4-S and BRD4-L knockdown cell lines were successfully established in HT29 and SW620 cells. (B) Colony formation assays were performed to evaluate the sphere-forming ability of BRD4-S and BRD4-L knockdown in HT29 and SW620 cells. (C,D) CCK-8 and scratch assays were conducted to assess the proliferative capacity of BRD4-S and BRD4-L knockdown cell lines. Data are presented as mean $\pm$ SEM. No significant difference (ns) is indicated when p $>$ 0.05; *p 0.05; **p 0.01. CCK-8, cell counting kit-8. Scale bar = 100 µm.

To further evaluate the oncogenic role of BRD4-S in vivo, we performed subcutaneous injections of BRD4-L-shRNA cells, BRD4-S-shRNA cells, and control cells into nude mice. In the xenograft model, tumor formation was significantly reduced in the BRD4-S-shRNA group compared with both the control and BRD4-L-shRNA groups (Fig. 3A). Tumor growth curve analysis further demonstrated that BRD4-S knockdown markedly suppressed tumor progression (Fig. 3B). Consistently, both tumor weight and volume were significantly lower in the BRD4-S-shRNA xenografts compared with the other groups (Fig. 3C,D). These results indicate that BRD4-S possesses oncogenic properties that drive CRC tumor growth.

Fig. 3.

BRD4-S promotes CRC tumorigenesis in vivo. (A) Representative images of tumor formation in nude mice at 8 weeks following ectopic transplantation of HT29 cells with BRD4-S knockdown or empty vector (n = 5). (B) Tumor volume measurements of the two groups (BRD4-S knockdown vs. empty vector–expressing HT29 cells) at 7, 14, 21, and 28 days after ectopic transplantation. (C,D) Comparison of tumor weights and volume between BRD4-S knockdown and empty vector–expressing HT29 cells in nude mice at 4 weeks post-ectopic transplantation. Data are presented as mean $\pm$ SEM. **p 0.01.

Using the GEPIA2 database, we systematically screened the top 100 genes associated with the BRD4-S and BRD4-L isoforms. After excluding genes co-associated with both isoforms, we identified 63 genes that were exclusively correlated with BRD4-S (Fig. 4A). Subsequent differential expression analysis of CRC transcriptomes from the TCGA cohort revealed 5331 significantly dysregulated genes ($|$log₂ fold change$|$ $>$1, adjusted p 0.05), as illustrated by volcano plot analysis (Fig. 4B). Among these, only eight DEGs demonstrated functional associations with BRD4-S (Fig. 4C). Of these candidates, ZFP64, RBL1, ZNF217, HNF4A, GNL3L, POFUT1, and DDX27 were consistently upregulated in CRC tissues compared with normal controls, whereas STAT5B was downregulated (Fig. 4D). To validate these findings, RT-PCR analysis was performed in HT-29 CRC cells. The results confirmed significant overexpression of most identified DEGs, with the exception of ZFP64, HNF4A, and GNL3L, which did not show statistically significant changes (Fig. 4E).

Fig. 4.

Identification of BRD4-S-specific functionally associated candidates in CRC. (A) Venn diagram analysis of the top 100 genes most strongly correlated with BRD4-L and BRD4-S in the GEPIA2 database identified genes exclusively associated with BRD4-S. (B) Volcano plot analysis of DEGs in CRC. (C) Venn diagram showing the overlap between DEGs in CRC and those correlated with BRD4-S expression. (D) mRNA expression of ZFP64, RBL1, ZNF217, HNF4A, GNL3L, POFUT1, and DDX27 in paired CRC tissues according to the GEPIA2 database. (E) qRT-PCR analysis of ZFP64, RBL1, ZNF217, HNF4A, GNL3L, POFUT1, and DDX27 in HT-29 cells. Data are presented as mean $\pm$ SEM. No significant difference (ns) is indicated when p $>$ 0.05; *p 0.05; **p 0.01.

To further investigate the regulatory role of candidate factors in BRD4 splicing, we performed GO and KEGG enrichment analyses on the previously identified genes. This analysis revealed a significant enrichment of DDX27 in spliceosome -related pathways (Fig. 5A). To evaluate the clinical relevance of DDX27 in CRC, we examined its expression in the TCGA CRC cohort. DDX27 mRNA levels were markedly elevated in CRC tissues compared with adjacent normal colon tissues (Fig. 5B), supporting its potential role as an oncogenic factor in CRC. Functional validation through DDX27 knockdown in HT-29 CRC cells (Fig. 5C) demonstrated reduced mRNA and protein levels of the BRD4-S isoform, accompanied by increased BRD4-L protein expression (Fig. 5D,E). Furthermore, correlation analysis using the GEPIA2 database identified SRSF6 as the splice factor most strongly associated with DDX27 (Fig. 5F).

Fig. 5.

DDX27-mediated BRD4 splicing regulation modulates isoform switching through SRSF6 interaction in HT-29 cells. (A) GO and KEGG enrichment analysis of DEGs in samples with high DDX27 expression. The red box indicates the association between DDX27 and the spliceosomal pathway investigated in subsequent studies. (B) Analysis of DDX27 mRNA Expression in Paired Colorectal Cancer Tissues from TCGA Cohort. (C) DDX27 knockdown cell lines were successfully established in HT29 and SW620 cells. (D,E) qRT‒PCR and Western blotting analyses of BRD4-L and BRD4-S expression in DDX27 knockdown HT-29 cells. (F) Correlation analysis between the splicing factor SRSF6 and DDX27. Data are presented as mean $\pm$ SEM. *p 0.05; **p 0.01; ***p 0.001.

Our study identified DDX27 as a regulator of BRD4-S expression in experimental models. To elucidate the molecular mechanisms underlying this regulation, we examined its interactions with splicing factors. RNA immunoprecipitation (RIP) assays confirmed the association of DDX27 with BRD4 pre-mRNA in HT29 cell extracts. The RIP assays also demonstrated that SRSF6 interacts with BRD4 pre-mRNA (Fig. 6A). To further investigate the specificity of the DDX27–SRSF6 interaction, co-immunoprecipitation (Co-IP) and immunofluorescence analyses were performed in HT29 cells using anti-DDX27 and anti-SRSF6 antibodies. These experiments suggested that DDX27 and SRSF6 likely form a complex (Fig. 6B,C). Moreover, knockdown of DDX27 resulted in a marked reduction of phosphorylated SRSF6 levels (Fig. 6D), indicating that DDX27 promotes SRSF6 phosphorylation. To assess whether SRSF6 phosphorylation is required for DDX27-mediated splicing activity, HT29 cells were treated with the SRSF6 phosphorylation inhibitor SPHINX31 at concentrations of 0, 1, and 2 µM for 24 hours. Inhibition of SRSF6 phosphorylation effectively suppressed the alternative splicing of BRD4 toward the BRD4-S isoform (Fig. 6E).

Fig. 6.

DDX27 Mediates BRD4-S alternative splicing through phosphorylation dependent interaction with SRSF6. (A) Immunoprecipitation from HT-29 cells using DDX27 and SRSF6 antibodies followed by RT-PCR or qPCR with BRD4-specific pre-mRNA primers. BRD4 pre-mRNA levels were quantified as fold enrichment relative to the IgG control. (B) Co-immunoprecipitation (Co-IP) of HT-29 cells with SRSF6, DDX27, or IgG antibodies, followed by western blot detection of SRSF6 and DDX27 in both precipitates and cell lysates. (C) Immunofluorescence analysis showing co-localization of SRSF6 (green) and DDX27 (red) with DAPI-stained nuclei (blue). Scale bar = 50 µm. (D) Western blot analysis of SRSF6 phosphorylation in HT-29 cells following DDX27 knockdown. (E) Western blot analysis of BRD4-S, BRD4-L, and p-SRSF6 levels in HT-29 cells treated with the SRSF6 phosphorylation inhibitor SPHINX31.

To elucidate the downstream pathways regulated by BRD4-S in CRC, we performed GO-KEGG enrichment analyses on both BRD4-S and BRD4-L. To avoid confounding effects from overlapping signals, pathways commonly enriched in both isoforms were excluded. The analysis revealed a significant enrichment of BRD4-S in the MAPK signaling pathway (Fig. 7A). To validate this finding, we examined MAPK pathway alterations in HT-29 cells following BRD4-S knockdown. Western blot analysis showed a reduction in ERK1/2 phosphorylation, whereas phosphorylation of JUN and P38, as well as total protein levels of ERK1/2, JUN, and P38, remained unchanged (Fig. 7B). Similarly, treatment of HT-29 cells with the SRSF6 phosphorylation inhibitor SPHINX31 suppressed ERK1/2 phosphorylation (Fig. 7C). To further establish a causal link between BRD4-S and MAPK/ERK pathway activation, we overexpressed BRD4-S in HT-29 cells and treated them with the clinically approved MEK inhibitor trametinib. Notably, while trametinib effectively reduced baseline ERK1/2 phosphorylation in control cells, BRD4-S overexpression not only enhanced p-ERK1/2 levels but also substantially reversed trametinib-mediated suppression of ERK phosphorylation (Fig. 7D). Finally, to assess whether BRD4-S directly regulates the transcription of ERK target genes, we performed chromatin immunoprecipitation followed by quantitative PCR (ChIP-qPCR) in HT-29 cells using an anti-BRD4 antibody. The results demonstrated significant enrichment of BRD4-S at the promoter regions of CCND1 and DUSP6 compared with the IgG control (Fig. 7E). These findings suggest that BRD4-S is directly recruited to the chromatin of key ERK target genes, thereby facilitating their transcription downstream of ERK activation.

Fig. 7.

BRD4-S drives CRC progression via MAPK/ERK signaling activation. (A) GO and KEGG enrichment analyses of highly expressed DEGs in BRD4-L and BRD4-S samples from the TCGA database. (B) Western blot analysis of ERK1/2, p-ERK1/2, p38, p-p38, c-Jun, and p-c-Jun in HT29 cells with BRD4-S knockdown. (C) Western blot analysis of ERK1/2, p-ERK1/2, p38, p-p38, c-Jun, and p-c-Jun in HT29 cells treated with the SRSF6 phosphorylation inhibitor SPHINX31 (0, 1, 2 µM). (D) Western blot analysis of ERK1/2 and p-ERK1/2 in NC–overexpressing and BRD4-S–overexpressing HT29 cells treated with or without the MEK inhibitor trametinib (5 µM). (E) ChIP-qPCR analysis showing enrichment of BRD4 at the promoter regions of ERK target genes CCND1 and DUSP6 in HT-29 cells. Data are presented as mean $\pm$ SEM. **p 0.01.