The 786-M1A and OS-LM1B human ccRCC cell lines were gifts from Professor Joan Massagué (Memorial Sloan Kettering Cancer Center, NY, USA). These cell lines are metastatic derivatives of 786-O and OS-RC2 cells, respectively (26). The UOK101 human ccRCC cell line was obtained from Dr Marston Linehan (National Cancer Institute, MD, USA). The cell lines were cultured in RPMI-1640 (Nacalai Tesque, Inc.) supplemented with 10% FBS (Tico Europe) and 1% penicillin-streptomycin solution. For lentiviral production, 293T cells (ATCC) were used and cultured in DMEM (Nacalai Tesque, Inc.) supplemented with 10% FBS and 1% penicillin-streptomycin solution. All cells were cultured at 37°C in a humidified atmosphere containing 5% CO₂. Blasticidin (InvivoGen) and hygromycin (Nacalai Tesque, Inc.) were used for antibiotic selection to isolate cells that were successfully transduced with the dCas9-HDAC3 and sgRNA expression plasmid, respectively. JQ1 and A-485 were purchased from Sigma-Aldrich (Merck KGaA). The dCas9-HDAC3 (cat. no. 98591; Addgene, Inc.) (27) and the lentivirus packaging plasmids psPAX2 (cat. no. 12260) and pMD2.G (cat. no. 12259) were obtained from Addgene, Inc. All primers and single guide RNA (sgRNA) constructs (Table I) were purchased from Integrated DNA Technologies, Inc.

The ccRCC cells, 786-M1A, OS-LM1B and UOK101, were seeded at a density of 500–1,000 cells/well in a 96-well. On the following day, the cell culture medium was replaced with fresh medium containing the respective chemical inhibitors, JQ1 or A-485. For JQ1 treatment, concentrations of 1, 10, and 20 µM were tested, whereas A-485 was used at concentrations of 1 and 10 µM. Fresh medium containing an equivalent volume of DMSO was added to control wells, where DMSO served as the vehicle for chemical inhibitor dissolution. The treated cells were incubated at 37°C for 72 h. The medium was replaced with 120 µl fresh medium containing AlamarBlue™ Reagent (Thermo Fisher Scientific, Inc.) at a 1:10 dilution. The cells were incubated for 2 h at 37°C in a humidified atmosphere with 5% CO₂, protected from light. A total of 100 µl medium from each well was transferred to a black 96-well plate and fluorescence was measured using a Varioskan Flash microplate reader (Thermo Fisher Scientific, Inc.) at an excitation wavelength of 560 nm and an emission wavelength of 590 nm. A total of three wells containing medium without cells were included as blank controls. Each treatment group consisted of five replicate wells. To determine the number of plated cells, baseline fluorescence measurements were taken on day 0 (24 h after initial seeding). The mean blank control readings was subtracted from those of each treatment well to eliminate background fluorescence. The fluorescence readings after 72 h incubation at 37°C, were normalized to the day 0 values. Finally, the relative fluorescence of inhibitor-treated cells was calculated against that of the DMSO-treated control group. Images were acquired using an inverted light microscope (Nikon TS100).

The ccRCC cells, 786-M1A, OS-LM1B and UOK101, were seeded at a density of 100–500 cells/well in a 6-well plate. The following day, the medium was replaced with medium supplemented with either JQ1 or A-485 (1 and 10 µM). Control cells were maintained in medium containing DMSO. Following 7 days of treatment at 37°C, the culture medium was aspirated and cells were rinsed twice with 1 ml 1X PBS. The PBS solution was then removed and the cells were fixed with 500 µl 1:1 mixture of 10% methanol and 10% acetic acid at room temperature for 15 min. The 6-well plates were placed on a horizontal shaker for 20 min at room temperature. Colonies, defined as visible cell clusters formed from a single cell, were stained with 500 µl 0.5% crystal violet staining solution (Sigma-Aldrich; Merck KGaA) for 30 min at room temperature. Excess stain was discarded and the wells were washed under running tap water. The plates were then air-dried in an oven until dry. Colony images were captured using the ChemiDoc MP imaging system (Bio-Rad Laboratories, Inc.).

For quantitative analysis of crystal violet staining, 600 µl of 10% acetic acid were added to each well, followed by incubation at room temperature on a shaker for 10 min to elute the bound dye. A 200-µl aliquot of the eluted solution was transferred to a clear 96-well plate and the absorbance was measured at 590 nm using the Varioskan Flash microplate reader (Thermo Fisher Scientific, Inc.). Absorbance values were normalized to the control wells and expressed as a percentage relative to the control group.

Total RNA was extracted from ccRCC cells using TRIzol^® reagent (Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. The yield and purity of total RNA were quantified using the NanoDrop™ 2000c Spectrophotometer (Thermo Fisher Scientific, Inc.). cDNA was synthesized from total RNA using a LunaScript RT SuperMix kit (New England BioLabs, Inc.) as follows: 25°C for 2 min, 55°C for 10 min and 95°C for 1 min. RT-qPCR was performed using TaqMan chemistry with pre-designed TaqMan probes (Thermo Fisher Scientific, Inc.; cat. nos. KLF6 Hs00810569_m1, β-actin Hs01060665_g1) and Luna Universal Probe qPCR Master Mix (New England BioLabs, Inc.). qPCR reactions were run on the 7500 Fast Real-Time System (Applied Biosystems; Thermo Fisher Scientific, Inc.) with the following cycling conditions: 95°C for 1 min for initial denaturation, followed by 40 cycles of 95°C for 15 s and 60°C for 30 s. β-actin served as the reference gene for normalization and gene expression fold-changes were calculated using the 2^−ΔΔCq method (28).

Site-directed mutagenesis of the original dCas9-HDAC3 plasmid was performed using a PCR-based whole-plasmid amplification approach to substitute the stop codon TAA with AAA. Complementary forward and reverse primers were designed to anneal to the target region of the dCas9-hHDAC3 plasmid. This primer pair carries the nucleotide to be substituted, located in the middle of the primers (Table I).

PCR amplification was performed using Accuprime™ Pfx Supermix (Thermo Fisher Scientific, Inc.) on the Thermal Cycler T100 (Bio-Rad Laboratories, Inc.). The thermocycling conditions were as follows: Initial denaturation at 95°C for 2 min, followed by 40 cycles of denaturation at 95°C for 15 sec, annealing at 50–65°C for 30 sec and extension at 68°C for 15 min. During amplification, the forward and reverse primers anneal to opposite strands of the plasmid in their respective 5′-3′ orientations and extend in opposite directions, resulting in amplification of the entire plasmid while incorporating the intended nucleotide substitution.

Following PCR amplification, the original and amplified plasmids were treated with DpnI restriction enzyme (New England BioLabs, Inc.), which selectively digests the methylated original plasmid template, for 1 h at 37°C. The undigested plasmids were transformed into Escherichia coli (New England BioLabs, Inc.) and selected on LB agar plates containing ampicillin. The plasmids were extracted from the transformed bacteria using the Monarch Plasmid Miniprep kit (New England BioLabs, Inc.) and subjected to Sanger sequencing (Apical Scientific, Inc.) to confirm the successful site-directed mutagenesis.

SE constituent regions at the KLF6 locus were identified based on our previously published H3K27ac ChIP-seq dataset (accession number GSE115749) (25). Regions with the highest H3K27ac signal intensity and closest proximity to the KLF6 locus were annotated as SE_1-3. The sgRNAs targeting these regions, iSE_1, iSE_2 and iSE_3, were described in our previous study using the webtool GPP sgRNA Designer (portals.broadinstitute.org/gppx/crispick/public) (25). The sgRNA sequences are listed in Table I. The dCas9-HDAC3 plasmid used for CRISPR-mediated deacetylation was obtained from Addgene, Inc (cat. no. 98591).

Plasmids were introduced into chemically competent E. coli cells by adding the plasmid DNA directly to the competent cells, followed by incubation on ice for 30 min. Heat shock was then performed at 42°C for 1 min, after which the tubes were immediately returned to ice for 5 min. Subsequently, 300 µl fresh Luria Bertani (LB; Sigma-Aldrich; Merck KGaA) medium was added to each tube and the cultures were incubated at 37°C with shaking at 200 rpm for 1 h. The transformed cells were spread onto LB agar plates supplemented with ampicillin (1:1,000) for selection of successfully transformed cells and incubated overnight at 37°C. Single colonies from LB + ampicillin plates were inoculated into 3 ml LB medium containing ampicillin (1:1,000) and cultured overnight at 37°C in a shaking incubator at 200 rpm. Plasmids were extracted using the Monarch Plasmid Miniprep kit (New England BioLabs, Inc.) according to the manufacturer's protocol. The concentration and purity of the extracted plasmids were assessed using a NanoDrop 2000c Spectrophotometer (Thermo Fisher Scientific, Inc.).

Lentivirus was produced by co-transfecting 293T cells (ATCC) with second generation lentivirus packaging plasmids psPAX2 and envelope plasmid pMD2.G together with the plasmid of interest using the Attractene transfection reagent (Qiagen GmbH). The lentivirus plasmids were obtained from Addgene, Inc. Briefly, 1.3 µg psPAX2, 0.5 µg pMD2.G and 1.5 µg plasmid of interest were mixed, followed by the addition of 10 µl Attractene transfection reagent. The mixture was incubated at room temperature for 30 min and added to the 293T cells. Transfected cells were incubated 37°C overnight, after which the media was replaced with fresh complete media. Lentivirus-containing media was collected 72 h post-transfection and filtered through a 0.45 µm Minisart NML syringe filter (Sartorius AG).

Cells at 60–70% confluency were transduced with filtered lentivirus-containing media in the presence of 8 µg/ml Polybrene (MilliporeSigma) and incubated at 37°C overnight. Cells transduced with dCas9-HDAC3 plasmid were selected with 30 µg/ml blasticidin for approximately 5 days, while cells transduced with the sgRNA expression vector were selected with 1,000 µg/ml hygromycin for approximately 7 days. For validation of dCas9-HDAC3 expression by western blotting, non-transduced parental cells were maintained in parallel and used as a negative control. In addition, untransduced cells were included during antibiotic selection to monitor selection efficiency. Selection was considered complete once all untransduced cells had died, after which the antibiotic-containing medium was replaced with standard culture medium.

The transduced ccRCC cells were lysed on ice in 1X RIPA buffer supplemented with 1:100 protease inhibitor cocktail (Nacalai Tesque, Inc.). The protein concentration was quantified using the Pierce BCA Protein Assay kit (Thermo Fisher Scientific, Inc.) according to the manufacturer's instructions. Equal amounts of protein (50 µg per lane) were denatured in 1X Laemmli SDS sample buffer (GeneTex, Inc.) containing 8% β-mercaptoethanol and resolved via 8% SDS-PAGE. Proteins were transferred onto nitrocellulose membranes (GE Healthcare), which were blocked with 5% non-fat milk in 1× TBS containing 0.1% Tween-20 for 1 h at room temperature. The membrane was incubated with primary antibodies overnight at 4°C. Following incubation with rabbit anti-mouse IgG-HRP (1:1,000; cat. no. P0260; Dako; Agilent Technologies, Inc.) for 1 h at room temperature, signals were detected using Pierce™ ECL substrate (Thermo Fisher Scientific, Inc.) and imaged using the ChemiDoc Go Imaging System (Bio-Rad Laboratories, Inc.). Primary antibodies against Cas9 (1:500; cat. no. sc-517386) and β-actin (1:5,000; cat. no. sc-69879) (both Santa Cruz Biotechnology, Inc.) were used. The Cas9 antibody was used to detect the dCas9-HDAC3 fusion protein expressed in transduced cells. dCas9 is a catalytically inactive variant of Cas9 generated by introducing two point mutations in the nuclease domains (29). As dCas9 retains the Cas9 protein sequence, it is recognized by Cas9 antibodies and detected by western blotting.

Cells were cross-linked with 10 ml 1% formaldehyde for 10 min at room temperature with gentle rotation to preserve protein-DNA interactions. The cross-linking reaction was quenched by adding 1 ml 0.125 M glycine and incubating cells for 5 min at room temperature with rotation. Cells were pelleted by centrifugation at 1,200 × g for 5 min at room temperature and washed twice with 15 ml cold 1X PBS. After the final wash, cell pellets were either snap-frozen and stored at −80°C or used immediately for chromatin extraction. Frozen cells were thawed on ice and lysed in 2 ml lysis buffer composed of 20 mM Tris-HCl (pH 8.0), 150 mM NaCl, 2mM EDTA (pH 8.0), 0.1% SDS and 1% Triton X-100. The cell suspension was pipetted 10 times and incubated on ice for 5 min. Homogenization was performed on ice using a Dounce homogenizer. The homogenate was transferred to a 15-ml centrifuge tube for sonication. Chromatin was fragmented by sonication using a Branson Ultrasonics Sonifier at a frequency of 20 kHz (30 sec on/off for 15 cycles) while keeping samples on ice to minimize heat damage. The lysate was centrifuged at 18,000 × g for 20 min at 4°C and the supernatant containing sheared chromatin was collected. A 10 µl aliquot of lysate was set aside as input control for downstream qPCR.

For IP, 100 µl magnetic protein A/G beads (Thermo Fisher Scientific, Inc.) were equilibrated three times with 1 ml 0.5% BSA (Sigma-Aldrich; Merck KGaA) in 1X PBS. The beads were resuspended in 500 µl 0.5% BSA and conjugated with 5 µg anti-H3K27ac (cat. no. ab4729) or control IgG antibody (cat. no. ab37415) (both Abcam) for ≥4 h at 4°C on a rotating platform. The antibody-conjugated beads were washed three times with 0.5% BSA and incubated overnight at 4°C with 600 µl chromatin lysate on a rotating platform. Beads were washed three times with low-salt wash buffer (50 mM HEPES, pH 7.5, 140 mM NaCl, 1% Triton) and once with high-salt wash buffer (50 mM HEPES, pH 7.5, 500 mM NaCl, 1% Triton). All washing steps were performed on a magnetic rack (Thermo Fisher Scientific, Inc.). To reverse cross-links and elute bound DNA, 100 µl elution buffer was added and samples were incubated at 65°C with shaking at 1,000 rpm for 3 h. DNA was purified using the Monarch PCR & DNA Cleanup kit (New England BioLabs, Inc.) according to the manufacturer's protocol.

qPCR was performed by adding the purified, de-crosslinked DNA to a reaction mix containing 2X PowerUp SYBR Green Master Mix (Thermo Fisher Scientific, Inc.) and 10 µM forward and reverse ChIP-qPCR iSE_1, ChIP qPCR iSE_2 and ChIP qPCR iSE_3 primers (Table I). The reaction was performed using the 7500 Fast Real-Time System (Applied Biosystems; Thermo Fisher Scientific, Inc.) with the following conditions: 50°C for 2 min, 95°C for 2 min for initial denaturation, followed by 40 cycles of 95°C for 3 sec and 60°C for 30 sec. Enrichment of ChIP DNA was quantified using the percent input (% input) method (30).

Statistical analysis was performed using GraphPad Prism software (version 10.6.0; Dotmatics). Data are presented as the mean ± SD from three independent experiments unless otherwise specified. Statistical significance was tested using one-way ANOVA followed by Tukey's or Dunnett's post hoc test or an unpaired t-test with Welch's correction. P<0.05 was considered to indicate a statistically significant difference.

As BRD4 serves pivotal roles in regulating the SE landscape and transcription of downstream genes (8), the present study evaluated whether ccRCC cells are sensitive to BRD4 inhibition by JQ1. 786-M1A, OS-LM1B and UOK101 human ccRCC cells were treated with increasing concentrations of JQ1 before cell viability assays at 72 h post-treatment. BRD4 inhibition significantly decreased ccRCC cell viability in a dose—dependent manner compared with that of the vehicle-treated cells (Figs. 1A and S1A). The present study also investigated the effect of BRD4 inhibition on the colony formation of ccRCC cells. JQ1 treatment, even at a concentration of 1 µM, significantly suppressed the clonogenic potential of ccRCC cells (Figs. 1B and S1B). Overall, ccRCC cells were sensitive to JQ1-mediated BRD4 inhibition, highlighting the key role of BRD4 in supporting ccRCC cell viability.

The p300 HAT is involved in regulating chromatin accessibility and transcription activation (12). Furthermore, p300 binding is also enriched at the SE regions, driving the expression of pro-oncogenes that promote cancer progression (12). The present study aimed to assess the effect of targeting p300 activity on ccRCC cell viability by treating 786-M1A and OS-LM1B cells with 1 and 10 µM A-485 for 72 h. In both ccRCC cell lines, inhibition of p300 with 10 µM A-485 significantly decreased the viability compared with that of vehicle-treated controls, whereas treatment with a lower concentration of A-485 did not result in a significant decrease in cell viability (Figs. 2A and S2A). By contrast, colony formation assays revealed that p300 inhibition with 1 or 10 µM A-485 significantly impaired the clonogenic capacity of ccRCC cells, which was consistent with the overall trend observed in the cell viability assays (Figs. 2B and S2B). Overall, ccRCC cells exhibited sensitivity to A-485-mediated p300 inhibition, which resulted in a marked reduction in cell viability and colony formation. This suggested that p300 serves a role in controlling the expression of genes that promote ccRCC cell viability.

KLF6 supports ccRCC cell proliferation by regulating cell networks through the transcriptional activation of platelet-derived growth factor β (PDGFβ), linking mTOR complex 1 (mTORC1) signaling pathway activity to lipid metabolism (25). While high expression of KLF6 is driven by one of the largest SEs in ccRCC (25), the roles of BRD4 and p300, key regulators of SEs and transcription activity, remain underexplored. Therefore, the present study investigated whether targeting of BRD4 and p300 with their respective inhibitors impairs KLF6 transcriptional activation in ccRCC cells. For BRD4 inhibition, 786-M1A and OS-LM1B cells were treated with 1 or 10 µM JQ1 for 6 h, which was a time point shown to not affect cell viability (data not shown), followed by assessment of KLF6 expression via qPCR analysis. Treatment with either 1 or 10 µM JQ1 significantly decreased KLF6 expression in both ccRCC cell lines compared with vehicle-treated controls. Although KLF6 downregulation was dose-dependent, the difference in the extent of reduction between 1 and 10 µM JQ1 was not significant (Fig. 3A).

To examine the role of p300, 786-M1A and OS-LM1B cells were treated with 1 or 10 µM A-485 for 16 h. qPCR analysis revealed that treatment with 1 µM A-485 did not significantly downregulate KLF6 expression in either ccRCC cell line. However, treatment with the higher concentration of A-485 decreased KLF6 expression by 50–60% in both 786-M1A and OS-LM1B cells (Fig. 3B). These findings underline the involvement of BRD4 and p300 in the regulation of KLF6 expression in ccRCC.

The present study investigated whether the downregulation of KLF6 following BRD4 and p300 inhibition was associated with changes in the chromatin landscape at the KLF6 SE region. H3K27ac ChIP-qPCR was performed to assess H3K27ac signal levels in JQ1- and A-485-treated 786-M1A cells. Constituent enhancers near the KLF6 locus (SE_1, SE_2 and SE_3) were selected from our previous H3K27ac ChIP-Seq dataset in ccRCC (25) based on their enrichment signal and proximity to the KLF6 promoter (Fig. 4A). Inhibition of BRD4 with 10 µM JQ1 decreased H3K27ac signals at all three constituent enhancers compared with those of vehicle-treated cells (Fig. 4B).

Similarly, treatment with the p300 inhibitor A-485 also decreased H3K27ac signals at these enhancer regions (Fig. 4C). As a negative control, IgG ChIP-qPCR was performed in parallel under the same conditions, which showed minimal background enrichment across all loci, confirming antibody specificity and negligible non-specific binding (Fig. 4B and C). Collectively, inhibition of BRD4 and p300 reduced H3K27ac signals, a marker of open chromatin and active transcription (7,10), at SE_1, SE_2 and SE_3 constituent enhancers, which potentially contributed to KLF6 downregulation.

A CRISPR-based approach was used to deacetylate the SE_1, SE_2 and SE_3 regions and assess the effects on chromatin accessibility and KLF6 expression. Prior to generating the 786-M1A cells that stably express dCas9-HDAC3, the plasmid was modified to remove the stop codon between the dCas9-HDAC3 and blasticidin-resistant gene reading frames (Fig. 5A). Site-directed mutagenesis was used to replace the stop codon (TAA) with alanine (AAA), allowing for blasticidin selection of cells that were successfully transduced with the dCas9-HDAC3-expressing plasmid (Fig. 5B and C). dCas9-HDAC3 protein expression in the transduced 786-M1A cells was confirmed by western blotting (Fig. 5D). These stable dCas9-HDAC3-expressing 786-M1A cells were transduced with either a non-targeting sgRNA control or iSE_1, iSE_2 and iSE_3 sgRNAs, which specifically target the SE_1, SE_2 or SE_3 enhancer regions, respectively. All sgRNAs were cloned into an sgRNA expression vector as described in our previous study (25). Western blot analysis confirmed stable expression of dCas9-HDAC3 in 786-M1A cells transduced with each sgRNA (Fig. 6A).

qPCR demonstrated ~25% KLF6 expression downregulation in dCas9-HDAC3-expressing 786-M1A cells transduced with iSE_1 sgRNA. However, there was no significant decrease in KLF6 expression in the iSE_2- and iSE_3-transduced cells (Fig. 6B). The downregulation of KLF6 in the iSE_1-transduced cells suggested that the decreased KLF6 expression may result from dCas9-HDAC3-mediated deacetylation of the SE_1 region. To assess the deacetylation efficiency, H3K27ac ChIP-qPCR was performed on the iSE_1 sgRNA-transduced cells. Decreased H3K27ac signals at the iSE_1-targeted enhancer indicated deacetylation of this region, which consequently altered chromatin accessibility and impaired KLF6 transcription (Fig. 6C). As a negative control, parallel IgG ChIP-qPCR demonstrated low background signal at this locus, confirming the specificity of the antibody and minimal non-specific binding.