PTIP epigenetically regulates DNA damage-induced cell cycle arrest by upregulating PRDM1

Nakata, Yuichiro; Nagasawa, Shion; Sera, Yasuyuki; Yamasaki, Norimasa; Kanai, Akinori; Kobatake, Kohei; Ueda, Takeshi; Koizumi, Miho; Manabe, Ichiro; Kaminuma, Osamu; Honda, Hiroaki

doi:10.1038/s41598-024-68295-w

PTIP epigenetically regulates DNA damage-induced cell cycle arrest by upregulating PRDM1

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Published: 03 August 2024

Volume 14, article number 17987, (2024)
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Scientific Reports

PTIP epigenetically regulates DNA damage-induced cell cycle arrest by upregulating PRDM1

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Yuichiro Nakata¹^na1,
Shion Nagasawa¹^na1,
Yasuyuki Sera²,
Norimasa Yamasaki³,
Akinori Kanai⁴,
Kohei Kobatake⁵,
Takeshi Ueda⁶,
Miho Koizumi²,
Ichiro Manabe¹,
Osamu Kaminuma³ &
…
Hiroaki Honda²

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Abstract

The genome is constantly exposed to DNA damage from endogenous and exogenous sources. Fine modulation of DNA repair, chromatin remodeling, and transcription factors is necessary for protecting genome integrity, but the precise mechanisms are still largely unclear. We found that after ionizing radiation (IR), global trimethylation of histone H3 at lysine 4 (H3K4me3) was decreased at an early (5 min) post-IR phase but increased at an intermediate (180 min) post-IR phase in both human and mouse hematopoietic cells. We demonstrated that PTIP, a component of the MLL histone methyltransferase complex, is required for H3K4me3 upregulation in the intermediate post-IR phase and promotes cell cycle arrest by epigenetically inducing a cell cycle inhibitor, PRDM1. In addition, we found that PTIP expression is specifically downregulated in acute myeloid leukemia patients. These findings collectively suggest that the PTIP-PRDM1 axis plays an essential role in proper DNA damage response and its deregulation contributes to leukemogenesis.

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Introduction

Cells have evolved highly orchestrated signaling pathways to repair DNA damage, since accumulation of DNA damage increases the risk of cancer¹. In the DNA damage response (DDR), cells recognize lesions and transiently arrest cell cycle progression to allow time for repair². DDR leads to cell cycle arrest, DNA repair, as well as apoptosis³ and prolonged cell cycle arrest following DNA damage is essential for effective DNA repair⁴. DNA double-strand breaks (DSBs) are the most cytotoxic type of DNA damage that occurs following DNA replication, exposure to ionizing radiation (IR), and treatment with chemotherapeutic drugs⁵. DSBs are repaired by either non-homologous end joining (NHEJ) or homologous recombination (HR)⁶. 53BP1 plays central roles in the NHEJ pathway—it is recruited to DSBs, protects DNA ends from excessive resection, and promotes end-joining of DNA ends⁷.

PTIP (PAX transcription activation domain interacting protein, also known as PAXIP1) contains multiple BRCA1 C-terminus (BRCT) domains that can interact with transcription factors and it functions as an adaptor protein downstream of p53-binding protein 1 (53BP1)^8,9. PTIP is recruited by phosphorylated 53BP1 to promote the NHEJ pathway and PTIP-deficient cells are sensitive to DNA damage, indicating that PTIP is directly involved in DNA repair via formation of the 53BP1 complex^8,10,11,12. On the other hand, PTIP functions as a component of the mixed-lineage leukemia (MLL) histone methyltransferase complexes that catalyze trimethylated histone H3 at lysine 4 (H3K4me3), a mark associated with gene activation^13,14. Indeed, PTIP is required for transcriptional activation via epigenetic regulation, such as peroxisome proliferator-activated receptor gamma (PPARγ) activation in preadipocytes and osteoclasts¹⁵ and sphingosine-1-phosphate receptor 1 (S1PR1) in T lymphocyte egress¹⁶. Moreover, PTIP is associated with Igh germline transcription in B cell class switch recombination (CSR) by regulating H3K4me3 and promotes CSR^17,18,19. Therefore, it is strongly suggested that PTIP contributes to cellular homeostasis through different functional properties.

Studies reported that methylation of histones at certain sites is associated with accessibility and recruitment of repair factors. Indeed, H3K4me3 and methylated H3K36 were found to promote the DDR pathway by recruiting repair factors^20,21,22. In addition, H3K27me3, a gene repression mark, also promotes the NHEJ pathway through interaction with FANCD2, a key player in the DDR pathway²³. While methylated histone marks induce alteration of chromatin structure and recruit repair factors to the lesion, transcriptional regulation by histone modifiers is also essential for DNA repair. For instance, EZH2, the histone methyltransferase catalyzing H3K27me3, promotes DNA repair by DNA damage-induced transcriptional silencing during the early phase following DNA damage²⁴. In addition, H3K4me3 levels are upregulated to activate DDR genes at the intermediate phase of DNA damage²⁵.

Whether PTIP contributes to DDR not only as a component of the 53BP1 complex but also as an epigenetic factor regulating H3K4me3 levels is not known. Since a variety of mutations in DDR genes are associated with increased risk of developing leukemia²⁶, we explored whether PTIP-mediated H3K4me3 regulation has essential roles in DDR using human leukemia cell lines and mouse hematopoietic stem-progenitor cells (HSPCs). In this study, we provide evidence that at the intermediate post-IR phase, PTIP-mediated upregulation of H3K4me3 and subsequent induction of PRDM1 (also known as BLIMP1) are necessary for DDR mediated cell cycle arrest.

Results

DNA damage by IR induced alteration of global H3K4me3 levels

To examine whether H3K4me3 levels are altered following IR, we first performed fluorescent immunostaining and western blotting of phosphorylated H2AX (γH2AX), a marker of DNA damage, in human myeloid leukemia cell lines. In all three cell lines (HL-60, MOLM13, and K562), global H3K4me3 levels were significantly downregulated 5 min after IR but were significantly upregulated 180 min after IR in line with γH2AX accumulation, compared with 0 min (Fig. 1A–C and Supplementary Fig. 1A). Upregulated H3K4me3 levels gradually returned to baseline in a time-dependent manner (Supplementary Fig. 1B). To determine whether these observations were physiologically relevant, we performed the same experiment using primary mouse HSPCs (LSK cells; lineage [Mac-1, Gr-1, B220, Ter119, CD4 and CD8]⁻, c-kit⁺ and Sca-1⁺). Similar results were obtained at both time points (Fig. 1D), indicating that global H3K4me3 levels were downregulated in the early phase (5 min) following IR, likely due to DNA damage induced transcriptional silencing²⁷, but upregulated in the intermediate phase (180 min) following IR (hereafter, referred to as early post-IR and intermediate post-IR, respectively, two timepoints in DNA repair pathway activation) in normal and leukemic hematopoietic cells.

PTIP induces upregulation of global H3K4me3 levels at the intermediate phase after IR

Since PTIP mediates H3K4 methylation as part of a histone methyltransferase complex and the cell lines we tested do not possess mutations or deletions at the PTIP gene (COSMIC, https://cancer.sanger.ac.uk/cosmic), we examined whether PTIP contributes to the upregulation of H3K4me3 at intermediate post-IR. To this end, we produced PTIP-knockdown human leukemic cells and Ptip deficient mouse hematopoietic cells. For human cells, lentiviral-based control shRNA vector (shCtrl) and two independent shRNA vectors targeting PTIP (shPTIP #1 and #2) were transduced into HL-60 cells (Fig. 2A). In contrast to shCtrl-transduced HL-60 cells that exhibited significant upregulation of global H3K4me3 levels at 180 min after IR, no apparent changes were detected in shPTIP-transduced cells (Fig. 2B). The global H3K4me3 levels at 0 min after IR in shCtrl, PTIP (shPTIP #1 and #2)-transduced HL-60 cells were comparable, suggesting that PTIP deficiency may not affect H3K4me3 regulation at steady state (Fig. 2C). To obtain Ptip-deficient mouse hematopoietic cells, mice in which exon 7 was floxed (Ptip^flox/flox) were crossed with ERT2Cre⁺ mice in which Cre was activated by tamoxifen, and Ptip^flox/flox, ERT2Cre⁺ mice were treated with tamoxifen to generate Ptip-deficient (Ptip^Δ/Δ) mice (Fig. 2D). Tamoxifen-treated Ptip^flox/flox, ERT2Cre⁻ mice were used as a control (Ptip^+/+). As observed in human leukemic cells, Ptip^+/+ LSK cells exhibited marked upregulation of global H3K4me3 levels at 180 min after IR, which was observed to a lesser extent in Ptip^Δ/Δ cells (Fig. 2E). H3K4me3 upregulation in Ptip^Δ/Δ LSK cells was not completely diminished, potentially due to regulation of H3K4me3 by the COMPASS complex including SETD1A/B or by the COMPASS-like complex including MLL1/2. These results indicated that PTIP is required for upregulation of global H3K4me3 levels in both human and mouse hematopoietic cells at intermediate post-IR.

PTIP suppresses cell cycle progression after IR via upregulation of PRDM1

A previous report identified IR-responsive genes that are associated with DDR and upregulated after IR²⁵. We thus hypothesized that PTIP contributes to DDR through a H3K4me3-mediated transcriptional program at intermediate post-IR. To investigate the expression changes of IR-responsive genes in leukemic cells, we first performed RT-qPCR using RNA extracted from HL-60 and MOLM13 cells before and after IR. We found that the expression of PRDM1, encoding a cell cycle inhibitor^28,29, was upregulated in both HL-60 and MOLM13 cells at 180 min after IR (Fig. 3A). Upregulation of PRDM1 at 180 min after IR was markedly attenuated in shPTIP-transduced HL-60 cells compared with shCtrl-transduced cells (Fig. 3B). These findings strongly suggest that PTIP contributes to upregulation of PRDM1 at intermediate post-IR. In addition, to explore the underlying transcriptional mechanisms, gene expression profiles of shCtrl- and shPTIP (#1 and #2)-transduced HL-60 cells at 0 min and 180 min after IR were examined by RNA-seq and the results were subjected to gene set enrichment analysis (GSEA) for pathway assessment (Fig. 3C and Supplementary Table. 1). In shCtrl-transduced HL-60 cells, gene sets associated with cell cycle proliferation (POSITIVE_REGULATION_OF_CELL_PROLIFERATION) were negatively enriched at 180 min after IR compared with 0 min, but no such effect was observed in shPTIP-transduced HL-60 cells (Fig. 3D, upper panels). Although no obvious changes were observed between shCtrl- and shPTIP-transduced HL-60 cells at 0 min after IR, these pathways were significantly positively enriched in shPTIP-transduced cells compared with shCtrl-transduced cells at 180 min after IR (Fig. 3D, lower panels). Following DNA damage, cell cycle checkpoints are activated and cell cycle arrest is transiently induced^2,7, suggesting that PTIP contributes to IR-induced cell cycle arrest.

To further investigate whether PTIP affects cell cycle activity via expression of PRDM1, we performed BrdU incorporation assays using IR-treated cells (Fig. 3E). We first examined the effect of IR on cell proliferation in HL-60 cells. As expected, IR-treated HL-60 cells exhibited markedly suppressed BrdU incorporation compared with untreated cells (Fig. 3F). We then performed the same assay using shCtrl- and shPTIP-transduced HL-60 cells. In contrast to shCtrl-transduced cells which exhibited significantly reduced BrdU incorporation, no such effects were detected in shPTIP-transduced cells (Fig. 3G). We further examined the effect of PRDM1 knockdown on cell proliferation following IR. We confirmed downregulation of PRDM1 with two different shRNAs (shPRDM1 #1 and #2) (Fig. 3H) and found that IR-induced cell cycle inhibition was not detected in shPRDM1-transduced cells (Fig. 3I). These findings collectively indicate that the PTIP-PRDM1 axis is essential for IR-induced cell cycle arrest.

A PTIP-containing MLL4 methyltransferase complex positively regulates H3K4me3 at the PRDM1 locus after IR

To explore the genome-wide distribution of H3K4me3, we performed CUT&RUN assays using HL-60 cells at 180 min after IR. In line with Fig. 1, genome-wide upregulation of H3K4me3 was observed in HL-60 cells after IR (Fig. 4A). We observed an increased number of H3K4me3 peaks (peak score > 3000, Supplementary Table. 2) at 180 min after IR compared with 0 min (Fig. 4B). In addition, the peaks were distributed genome-wide (Supplementary Fig. 2). The expression of H3K4me3 upregulated genes at 180 min after IR compared with 0 min, identified in Fig. 4B, was attenuated following PTIP knockdown (Fig. 4C). Motif analysis on these 1410 H3K4me3 peaks demonstrated greater enrichment for the E2F transcription factor, known to be associated with both cell cycle progression and cell cycle arrest³⁰. Indeed, the putative target genes (± 3 kb from the transcription start sites (TSSs)) of these peaks are involved in the cell cycle pathway, which are different properties compared with the H3K4me3 peaks at 0 min (Fig. 4D,E and Supplementary Table. 3).

To explore the direct relationship between PRDM1 expression and H3K4me3 upregulation, we analyzed whether H3K4me3 peaks accumulated at the PRDM1 locus. As expected, enrichment of H3K4me3 at PRDM1 was significantly increased downstream of the TSS in HL-60 cells at 180 min after IR compared with 0 min (Fig. 4F). In accordance with upregulation of H3K4me3, PTIP and MLL4 were also significantly enriched at this region at 180 min after IR compared with 0 min (Fig. 4G). In addition, H3K4me3 upregulation at 180 min after IR was attenuated by knockdown of PTIP (Fig. 4H). To investigate whether the regions downstream of the PRDM1 TSS are essential for transcription, dCas9-VP64 with 4 types of guide RNA (gRNA): #1 upstream of TSS, #2 and #3 putative PRDM1 regulatory regions, and #4 downstream of exon 2, were recruited with lentiviral vectors (Fig. 4I). We found that #2 and #3 regions positively regulated transcription in accordance with increased H3K4me3 (Fig. 4J). Together, these findings demonstrate that an MLL4 methyltransferase complex containing PTIP directly upregulates H3K4me3 levels downstream of the PRDM1 TSS, resulting in increased PRDM1 expression and inhibited cell cycle progression in HL-60 cells at intermediate post-IR.

PTIP is downregulated in human AML

Given that PTIP is associated with DDR directly via DNA repair and with cell cycle regulation via histone modification, PTIP is assumed to function as a tumor suppressor. To assess the role of PTIP in leukemogenesis, we compared the expression of PTIP in human leukemic samples and cell lines. We found that PTIP mRNA expression was significantly downregulated in acute myeloid leukemia (AML) samples with different karyotypes and cell lines compared with normal stem/progenitor cells (Fig. 5A–C), strongly suggesting that PTIP may play a tumor suppressive role, possibly through activation of both DDR pathways.

Discussion

It has been reported that trimethylation of H3K9 and H3K27, which changes the chromatin structure to a repressive state known as heterochromatin, plays essential roles for chromatin remodeling and transcriptional silencing in DDR^23,27,31. On the other hand, H3K4me3 mediated by the MLL4 methyltransferase complex containing PTIP, changes chromatin structure to an active state, called euchromatin, and is also thought to be one of the critical histone modifications for DDR. However, the mechanism underlying H3K4me3 regulation associated with PTIP in DDR has remained controversial^{20,21,25,32,33}.

While several groups suggested the importance of H3K4me3 in DDR^25,33, others argued that the demethylation of H3K4me3 positively regulates DDR^20,21,32. These contradictory results raise the question as to which mark (methylated or demethylated H3K4) is beneficial for DDR. In this study, we demonstrated that global H3K4me3 levels were downregulated at early (5 min) post-IR but upregulated at intermediate (180 min) post-IR in hematopoietic cells, which is in line with the results by Rashi-Elkeles, S. et al.²⁵ (Fig. 1). Our data support the idea that the contributions of H3K4me3 to DDR are time dependent. Indeed, treatment with JIB-04, an inhibitor of KDM5B, a demethylase of H3K4, in early post-IR induced an accumulation of H3K4me3 and impaired DNA repair pathways in non-small cell lung cancer (NSCLC) cells²¹. On the other hand, when NSCLC cells were treated with JIB-04 at intermediate post-IR, DNA repair pathway impairment was attenuated²³. Additionally, highly compacted heterochromatin protects the genome from DNA damage but also suppresses repair efficiency and increases the mutation rate by creating a barrier to repair factor recruitment³³, indicating that chromatin structure should be altered at the proper phase in DDR. These results imply that accumulation of H3K4me3 is dispensable or toxic for DDR at early post-IR but crucial for DDR at intermediate post-IR.

PTIP deficiency suppressed accumulation of H3K4me3 and promoted cell proliferation pathways at intermediate post-IR (Figs. 2, 3A), suggesting that PTIP possibly fine-tunes cell cycle progression at intermediate post-IR via H3K4me3 regulation. Our findings also highlighted that PTIP mediates expression of PRDM1, which functions as both a cell cycle inhibitor^8,29 and an IRinduced protein²⁵ by upregulating H3K4me3 in cooperation with MLL4 (Figs. 3, 4). Thus, PTIP is crucial for temporal cell cycle arrest after IR and for the DNA repair process.

In this study, we demonstrated that following DNA damage, downregulation of H3K4me3 was observed as a primary event and upregulation of H3K4me3 by an MLL4 methyltransferase complex containing PTIP occurred as a secondary event to induce cell cycle arrest via PRDM1 activation (Fig. 5D). Understanding this mechanism may also shed light on the multifaceted functions of PTIP. In addition, to address the issue that PTIP functions as a tumor suppressive factor through activation of both DDR pathways in greater detail, future studies can aim to investigate whether Ptip^Δ/Δ mice develop leukemia spontaneously via in vivo mutagenesis using retroviral transduction assays and exposure to a chemical carcinogen.

Materials and methods

Cells and mice

Human leukemia cell lines (HL-60, MOLM13, and K562), provided by Dr. Toshiya Inaba (Hiroshima University, Hiroshima, Japan), were grown in RPMI-1640 medium containing 10% fetal bovine serum supplemented with penicillin and streptomycin. The detailed procedures for construction of the targeting vector and generation of the cKI mice are described in Fig. 2C. CreERT2 + mice (C57BL/6-Gt(ROSA)26Sortm1(cre/Est1)Arte) were purchased from Taconic Biosciences (Hudson, NY, USA). Cre activation was achieved by intraperitoneal administration tamoxifen (Sigma, T5648). The all animal experiments complied with the ARRIVE guidelines and were conducted in accordance with the Guide for the Care and Use of Laboratory Animals of Hiroshima University Animal Research Committee (permission no. A17-84). Cervical dislocation under light anesthesia was performed as a euthanasia method for mice according to the American Veterinary Medical Association (AVMA) Guidelines. Mouse HSPCs (LSK cells) were stained with antibodies listed in Supplementary Table. 4 and sorted with a FACSAria II (BD Biosciences). LSK cells were grown in S-clone SF-O3 serum-free medium (Iwai North America Inc.) containing 1% BSA supplemented with cytokines (100 ng/mL mouse SCF, 100 ng/mL mouse TPO, 100 ng/mL mouse FLT3L and 50 ng/mL mouse IL-6) (Pepro Tec). CB derived human CD34⁺ stem/progenitor cells (Lonza) were cultured in StemSpan (Stem Cell Technologies) with 50 ng/mL human SCF (PeproTech), 50 ng/mL human TPO (PeproTech), and 50 ng/mL human Flt3-ligand (PeproTech). The cells were maintained in a humidified, 5% CO₂ atmosphere at 37 ℃. Cultured CD34⁺ cells were stained with antibodies listed in Supplementary Table. 4 and sorted as described above.

Radiation treatment

Cells were irradiated with X-rays using a CP-160 (Acrobio). Total radiation dose was as follows: 5 Gy in human leukemia cell lines and 2 Gy in LSK cells from Ptip^+/+ and Ptip^Δ/Δ mice. After IR, cells were incubated for the indicated time periods at 37 ℃.

Immunofluorescent staining

Cells were plated on poly-L-lysine-coated glass slides (Matsunami Glass), fixed with 4% paraformaldehyde, permeabilized with 0.5% Triton X-100 and treated with Protein block serum-free (Dako) to block non-specific binding. Subsequently, the cells were stained with anti-H3K4me3 and anti-γH2AX antibodies (Supplementary Table. 4). After washing with PBS, the cells were stained with Alexa Fluor 488- or 555-conjugated anti-rabbit and mouse IgG antibodies (Invitrogen). The nuclei were counterstained with Hoechst 33342 (Dojindo Laboratories). Images were taken with an FV-1000 confocal microscope (Olympus) and the fluorescence intensity of individual cells was computationally quantified using ImageJ64 software and were normalized against the mean fluorescence intensity of each control cell type (0 min after IR), as the relative fluorescence intensity³⁴.

Lentiviral vector transduction

To produce lentivirus containing supernatant, pLV-H1-shRNA-EF1α-GFP-Bsd (Biosettia), EF1α-dCas9-VP64-EGFP, Lenti-gRNA-EF1α-BleoR (kindly provided from Dr. Hoshii, Chiba university), Gag-Pol, Rev, and VSV-G expression vectors (Addgene) were transduced into Lenti-X 293 T cells (Takara Bio Inc.) with Lipofectamine 2000 (Invitrogen) and concentrated with Lenti-X concentrator (Takara Bio Inc.). In knockdown experiments, HL-60 cells were infected with lentivirus containing supernatant carrying shCtrl, shPTIP (#1 and #2) and shPRDM1 (#1 and #2). The cell lines were established and expanded before radiation treatment. In dCas9 experiments, HL-60 cells were infected with lentiviral supernatant containing dCas9-VP64 and gRNA (#1 to #4). The shRNA and gRNA sequences are listed in Supplementary Table. 5.

Quantitative real-time PCR

Total cellular RNA was isolated with TRIZOL reagent (Invitrogen) and reverse-transcribed with a SuperScript VILO master mix kit (Invitrogen) according to the manufacturer’s protocol. Quantitative real-time PCR was performed with power SYBR Green PCR Master Mix (Thermo Fisher Scientific) on a StepOnePlus Real-Time PCR system (Applied Biosystems). The primer sequences are listed in Supplementary Table. 5. All data are presented relative to GAPDH.

Histone extraction and western blotting

Cells were resuspended in pH 6.5 lysis buffer (10 mM Tris- HCl at pH 8, 50 mM sodium bisulfite, 1% Triton X-100, 10 mM MgCl2, 8.6% sucrose, 10 mM sodium butyrate). After the supernatant was discarded, the pellet was washed with pH 7.4 wash buffer (10 mM Tris, 13 mM EDTA). After the supernatant was discarded, the pellet was dissolved in 0.4 M H₂SO₄ for 1 h on ice. The supernatant was collected into a new tube including 80% of acetone and incubated overnight at − 20 ℃. After the supernatant was discarded, the pellets were air-dried and resuspended in water. Histone concentration was assayed using Bradford assay. Three micrograms of total histone extract were loaded into a 15% SDS-PAGE gel. Western blotting was performed as previously described³⁵.

RNA-seq and analysis

Total RNA from duplicate samples (shPTIP #1 and #2) was extracted with RNeasy mini kit (QIAGEN) and RNA-Seq libraries were prepared with the TruSeq Stranded mRNA Library Prep Kit (Illumina). Transcriptome analysis was performed using a next-generation sequencer (HiSeq 2500; Illumina) according to the manufacturer’s instructions. The generated sequence tags (more than 2.0 × 10⁷ reads for each sample) were mapped onto the human genomic sequence (UCSC Genome Browser, version hg19) with the sequence alignment program Tophat. Transcriptomes were compared with FPKM values and analyzed with Gene Set Enrichment Analysis (GSEA_4.3.3) software (Broad Institute, https://www.gsea-msigdb.org/gsea/index.jsp). Gene sets with a false discovery rate (FDR) q-value of < 0.25 were considered statistically significant.

BrdU incorporation analysis

BrdU incorporation was analyzed with a BrdU Flow Kit (BD Biosciences). Pre-cultured HL-60 cells 60 min after IR were treated with BrdU (10 μM) into cultured medium 240 min before the analysis. Cells were fixed, permeabilized and stained with an anti-BrdU antibody conjugated APC (Supplementary Table. 4). BrdU positive cells were measured with a FACSCanto II (BD Biosciences). Data were analyzed with FlowJo v10.10 software (BD Biosciences).

ChIP-qPCR analysis

Chromatin was enzymatically shred and immunoprecipitated with the ChIP-IT Express Enzymatic Kit (Active Motif) according to the manufacturer’s instructions. Antibodies are listed in Supplementary Table. 4. Subsequently, the chromatin was cleaned up with a QIAquick PCR purification kit (QIAGEN). Primer sequences are listed in Supplementary Table. 5.

CUT&RUN assay and analysis

Anti-H3K4me3 and anti-IgG antibodies, listed in Supplementary Table. 4, were used for CUT&RUN performed with the CUT&RUN assay kit (Cell Signaling), according to the manufacturer’s protocol. Libraries were generated using the NEBNext Ultra II DNA Library Prep Kit for Illumina (NEB, E7370L) and sequenced on an Illumina NovaSeq 6000. Paired-end fastq files were processed using the ENCODE Transcription Factor and Histone ChIP-Seq pipeline (https://github.com/ENCODE-DCC/chip-seq-pipeline2). Reads were trimmed using cutadapt v2.5. Reads were aligned to the hg19 genome using Bowtie2 v2.3.4.3, and SAMtools v1.9 was used to convert the output file to BAM format. Duplicates were removed using Picard Tools v2.20.7. Peak calling was performed with MACS2 v2.2.4, and the peaks compared with each IgG sample for subsequent analysis. Narrow peaks (1142–10293 bp, peak score: > 3000) were detected as H3K4me3 enriched peaks, listed in Supplementary Table. 1. Bigwig files were normalized by a scaling factor using Deeptools v3.3.1and bamCoverage tools and visualized in the UCSC genome browser (https://genome.ucsc.edu). The scaling factors were calculated by determining the number of spike-in DNA reads and normalized to total sequencing depth. Homer v4.11 was used for peak annotation and motif analysis. Bedtools v2.29.0 intersect was used to determine overlapping peaks (with at least 1 bp overlap) and assign target genes.

Statistical analysis

The statistical significance of differences between two samples was assessed by two-tailed Student’s t test. When multiple samples were compared, the statistical significance of differences was assessed using one-way ANOVA followed by Dunnett’s test. All statistical tests were performed with Prism software version 6 (GraphPad). All data were obtained from at least two independent experimental replicates. In quantitative real-time PCR, SDs were calculated with technical triplicates, thus statistical significance was not shown.

Data availability

The datasets generated and analyzed during the current study are available in the DNA Data Bank of Japan (DDBJ) repository (https://www.ddbj.nig.ac.jp/) (RNA-seq; DRA010427 and CUT&RUN; DRA016981 (rep1) and DRA018596 (rep2)).

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Acknowledgements

We thank Yuki Sakai, Sawako Ogata, and Megumi Nakamura for animal care, genotyping, and molecular experiments as well as Dr. Yoshinori Ohno for technical assistance. We also thank Dr. Junji Takeda (Osaka University, Japan) and RIKEN BRC for providing us with KY1.1 ES cells and B6.Cg-Tg(CAG-Cre)CZ-MO2Osb mice (RBRC01828), respectively. This work was, in part, supported by a JSPS KAKENHI (18K16087), the Kato Memorial Bioscience Foundation and AMED-CREST (JP24gm1310006).

Funding

JSPS KAKENHI,18K16087,Kato Memorial Bioscience Foundation,AMED-CREST,JP24gm1310006.

Author information

These authors contributed equally: Yuichiro Nakata and Shion Nagasawa.

Authors and Affiliations

Department of Systems Medicine, Graduate School of Medicine, Chiba University, 1-8-1 Inohana, Chuo-Ku, Chiba-Shi, Chiba, 260-8670, Japan
Yuichiro Nakata, Shion Nagasawa & Ichiro Manabe
Field of Human Disease Models, Major in Advanced Life Sciences and Medicine,, Institute of Laboratory Animals, Tokyo Women’s Medical University, 8-1 Kawada-Cho, Shinjuku-Ku, Tokyo, 162-8666, Japan
Yasuyuki Sera, Miho Koizumi & Hiroaki Honda
Department of Disease Models, Research Institute for Radiation Biology and Medicine, Hiroshima University, 1-2-3 Kasumi, Minami-Ku, Hiroshima, 734-8553, Japan
Norimasa Yamasaki & Osamu Kaminuma
Laboratory of Systems Genomics, Department of Computational Biology and Medical Sciences, Graduate School of Frontier Sciences, The University of Tokyo, 5-1-5 Kashiwanoha, Kashiwa, Chiba, 277-8561, Japan
Akinori Kanai
Department of Urology, Institute of Biomedical and Health Sciences, Hiroshima University, 1-2-3 Kasumi, Minami-Ku, Hiroshima, 734-8553, Japan
Kohei Kobatake
Department of Biochemistry, Faculty of Medicine, Kindai University, 377-2 Ohnohigashi, Osakasayama-Shi, Osaka, 589-8511, Japan
Takeshi Ueda

Authors

Yuichiro Nakata
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Shion Nagasawa
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Yasuyuki Sera
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Norimasa Yamasaki
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Akinori Kanai
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Kohei Kobatake
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Takeshi Ueda
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Miho Koizumi
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Ichiro Manabe
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Osamu Kaminuma
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Hiroaki Honda
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Contributions

Y.N. and H.H. designed the study. H.H., N.Y., and M. K. constructed the targeting vectors and generated genetically engineered mice. Y.N. and A.K. performed RNA-seq analysis. Y.N. and K.K. performed immunofluorescent staining and flow cytometric analysis. Y.N., S.N., and Y.S. performed Cut &Run, ChIP-qPCR and dCas9 assays. Y.N., T.U., I.M., O.K., and H.H. wrote and revised the manuscript. All authors reviewed and agreed on the final version of the manuscript.

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Correspondence to Yuichiro Nakata or Hiroaki Honda.

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Nakata, Y., Nagasawa, S., Sera, Y. et al. PTIP epigenetically regulates DNA damage-induced cell cycle arrest by upregulating PRDM1. Sci Rep 14, 17987 (2024). https://doi.org/10.1038/s41598-024-68295-w

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Received: 31 August 2023
Accepted: 22 July 2024
Published: 03 August 2024
DOI: https://doi.org/10.1038/s41598-024-68295-w
Springer Nature Limited

PTIP epigenetically regulates DNA damage-induced cell cycle arrest by upregulating PRDM1

Abstract

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Introduction

Results

DNA damage by IR induced alteration of global H3K4me3 levels

PTIP induces upregulation of global H3K4me3 levels at the intermediate phase after IR

PTIP suppresses cell cycle progression after IR via upregulation of PRDM1

A PTIP-containing MLL4 methyltransferase complex positively regulates H3K4me3 at the PRDM1 locus after IR

PTIP is downregulated in human AML

Discussion

Materials and methods

Cells and mice

Radiation treatment

Immunofluorescent staining

Lentiviral vector transduction

Quantitative real-time PCR

Histone extraction and western blotting

RNA-seq and analysis

BrdU incorporation analysis

ChIP-qPCR analysis

CUT&RUN assay and analysis

Statistical analysis

Data availability

References

Acknowledgements

Funding

Author information

Authors and Affiliations

Contributions

Corresponding authors

Ethics declarations

Competing interests

Additional information

Publisher's note

Supplementary Information

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