The Histone Demethylase JMJD2B Plays an Essential Role in Human Carcinogenesis through Positive Regulation of Cyclin-Dependent Kinase 6

Covalent histone modifications, including acetylation, methylation, phosphorylation, ubiquitination, glycosylation, and sumoylation can modulate chromatin dynamics and affect multiple cellular functions (1–3). Among these modifications, histone methylation is associated with activated or repressed transcription (3). Five lysine residues (H3K4, H3K9, H3K27, H3K36, and H4K20), located in the N-terminal tails of histones, are reported as representative lysines which can become mono-, di-, or trimethylated. According to recent findings, H3K9, H3K27, and H4K20 methylation mainly represses transcription, whereas methylation of H3K4 and H3K36 is associated with activated transcription (3). Although histone methylation had been thought to be irreversible, lysine specific demethylase 1, LSD1, was discovered to be the first example of a demethylase that can reverse histone H3 lysine 4 methylation status (4, 5). Later, the Jumonji C (JmjC) domain containing protein family, which catalyzes the hydroxylation of a lysine methyl group via a radical-based mechanism, has been identified as histone demethylases which differ from LSD1 (6, 7). Although our knowledge of the physiologic functions of histone demethylases is increasing, it still remains unclear how deregulation of the enzymes is involved in human diseases such as cancer.

We previously reported that the histone methyltransferase SMYD3 plays a critical role in human carcinogenesis (8–10). Besides our research, other groups also clarified that dysfunction of histone methylation status contributes to human carcinogenesis (11–13), but the detailed relationship between abnormal histone demethylation and human carcinogenesis is unclear. To find demethylases that contribute to human carcinogenesis, we examined the expression profiles of a number of proteins containing JmjC histone demethylase domains in clinical tissues and found that expression levels of JMJD2B were significantly upregulated in cancer tissues, compared with those in corresponding normal tissues. JMJD2B, also known as KDM4B, was identified in silico (14) and shown to be one of the demethylases capable of removing the trimethyl group from histone H3 lysine 9 on pericentric heterochromatin in mammalian cells (15). A line of recent reports indicated that hypoxic conditions can induce the expression of some JmjC family members, including JMJD2B (16, 17). In fact, JMJD2B has been shown to harbor HIF binding sites in their promoter sequences (17). However, the significance of JMJD2B in oncogenesis and cancer progression is not fully understood so far.

Here, we showed a critical role for JMJD2B in carcinogenesis, through the regulation of cancer-related downstream genes, and suggested the possibility that JMJD2B might be a novel therapeutic target for several types of cancer, especially bladder and lung cancer.

Bladder tissue sampling and RNA preparation were described previously (18). Briefly, 76 surgical specimens of primary urothelial carcinoma were collected, either at cystectomy or transurethral resection of bladder tumor, and snap frozen in liquid nitrogen. Twenty specimens of normal bladder urothelial tissue were collected from areas of macroscopically normal bladder urothelium in patients with no evidence of malignancy. Vimentin is primarily expressed in mesenchymally derived cells and was used as a stromal marker. Uroplakin is a marker of urothelial differentiation and is preserved in up to 90% of epithelially derived tumors (19). Use of tissues for this study was approved by Cambridgeshire Local Research Ethics Committee (Ref 03/018).

NIH3T3, CCD-18Co, SW780, SCaBER, A549, H2170, SW480, HCT116, LoVo, HepG2, HeLa, HFL1, and 293T cells were from American Type Culture Collection in 2001 and 2003 and tested and authenticated by DNA profiling for polymorphic short-tandem repeat (STR) markers. RERF-LC-AI, SBC5 and Huh-7 cells were from Japanese Collection of Research Bioresources (JCRB) in 2001 and tested and authenticated by DNA profiling for polymorphic STR markers. The 253J cells were from Korean Cell Line Bank in 2001 and tested and authenticated by DNA profiling for polymorphic STR markers. ACC-LC-319 cells were from Aichi Cancer Center in 2003 and tested and authenticated by DNA profiling for SNP, mutation, and deletion analysis. All cell lines were grown in monolayers in appropriate media: Dulbecco modified Eagle medium (DMEM) for RERF-LC-AI, HepG2, Huh-7, NIH3T3, and 293T cells; Eagleminimal essential medium (EMEM) for 253J, CCD-18, SCaBER, HeLa, SCaBER, and SBC5 cells; McCoy 5A medium for HCT116 cells; Leibovitz L-15 for SW480 and SW780 cells; RPMI-1640 medium for A549, H2170, and ACC-LC-319 cells, all supplemented with 10% FBS and 1% antibiotic/antimycotic solution (Sigma). LoVo cells were cultured in Ham F-12 medium supplemented with 20% FBS and 1% antibiotic/antimycotic solution. HFL1 cells were cultures in F-12K medium supplemented with 10% FBS, 1% antibiotic/antimycotic solution, 2 mmol/L l-glutamine, and 1,500 mg/L sodium bicarbonate. Cells were maintained at 37°C in humid air with 5% CO₂ condition (RERF-LC-AI, HepG2, Huh-7, NIH3T3, 293T, HeLa, SCaBER, SBC5, HCT116, A549, H2170, and ACC-LC-319) or without CO₂ (SW480 and SW780). Cells were transfected with FuGENE6 (Roche Applied Science) according to the manufacturer protocol.

We established a genome-wide cDNA microarray with 36,864 cDNAs selected from the UniGene database of the National Center for Biotechnology Information (NCBI). This microarray system was constructed essentially as described previously (20). Briefly, the cDNAs were amplified by reverse transcriptase PCR (RT-PCR) using poly (A)⁺ RNAs isolated from various human organs as templates; the lengths of the amplicons ranged from 200 to 1,100 bp, without any repetitive or poly (A) sequences. Many types of tumor and corresponding nonneoplastic tissues were prepared in 8-μm sections, as described previously (20). A total of 30,000 to 40,000 cancer or noncancerous cells were collected selectively using the EZ cut system (SL Microtest GmbH) according to the manufacturer protocol. Extraction of total RNA, T7-based amplification, and labeling of probes were done as described previously (20). A measure of 2.5-μg aliquots of twice amplified RNA (aRNA) from each cancerous and noncancerous tissue was then labeled, respectively, with Cy3-dCTP or Cy5-dCTP.

As described previously, we prepared 76 bladder cancer and 20 normal bladder tissues in Addenbrooke's Hospital, Cambridge UK. For quantitative RT-PCR reactions, specific primers for all GAPDH (housekeeping gene), JMJD2B, HDAC1, AKT3, and MAP3K1 were designed (Primer sequences in Supplementary Table S1; ref. 21). PCR reactions were done using the LightCycler 480 System (Roche Applied Science) following the manufacturer protocol. Fifty percent SYBR GREEN universal PCR Master Mix without UNG (Applied Biosystems), 50 nmol/L each of the forward and reverse primers and 2 μL of reversely transcribed cDNA were applied. Amplification conditions were 5 minutes at 95ºC and then 45 cycles each consisting of 10 sec at 95ºC, 1 minute at 55ºC and 10 sec at 72ºC. Then, reactions were heated for 15 sec at 95ºC, 1 minute at 65ºC to draw the melting curve, and cooled to 50ºC for 10 seconds. Reaction conditions for target gene amplification were as described above and the equivalent of 5 ng of reverse transcribed RNA was used in each reaction. mRNA levels were normalized to GAPDH expression.

Immunohistochemical analysis was done using anti-JMJD2B antibody (A301-478A; Bethyl Laboratories) as described previously (22). For clinical bladder and lung cancer tissue microarray, EnVision kit/horseradish peroxidase (HRP; Dako) was applied. Briefly, slides of paraffin-embedded tumor specimens were processed under high pressure (125ºC, 30 seconds) in antigen-retrieval solution, high pH 9 (S2367; Dako), treated with peroxidase blocking regent, and then treated with protein blocking regent (X0909; Dako). Tissue sections were incubated with a rabbit anti-JMJD2B polyclonal antibody followed by secondary antibodies conjugated to peroxidase labeled dextran polymers (Dako). Antigen was visualized with substrate chromogen (Dako liquid DAB chromogen; Dako). Finally, tissue specimens were stained with Mayer's haematoxylin (Hematoxylin QS, Vector Laboratories) to discriminate the nucleus from the cytoplasm. Because the intensity of staining within each tumor tissue core was mostly homogeneous, the intensity of JMJD2B staining was semiquantitatively evaluated using the following criteria: negative (no appreciable staining in tumor cells) and positive (brown staining appreciable in more than 30% of the nucleus of tumor cells).

HeLa cells were transfected with pCAGGS-n3FC-JMJD2B. Forty-eight hours after transfection, cultured cells were fixed by 4% paraformaldehyde in 0.1 mol/L phosphate buffer (pH 7.4) at room temperature for 30 minutes and permeabilized with 0.5% Triton X-100 in PBS (Sigma). Fixed cells were blocked with 5% bovine serum albumin in PBS for 1 hour and incubated with primary antibodies overnight at 4ºC. Then they were incubated with Alexa Fluor–conjugated second antibodies (Molecular Probes; Invitrogen) and observed using a Leica confocal microscopy (23).

siRNA oligonucleotide duplexes targeting the human JMJD2B transcripts were purchased from Sigma Genosys. siEGFP and siNegative control (siNC), which is a mixture of 3 different oligonucleotide duplexes, were used as control siRNAs. The siRNA sequences are described in Supplementary Table S2. siRNA duplexes (100 nmol/L final concentration) were transfected into bladder and lung cancer cell lines with Lipofectamine 2000 (Invitrogen), and cell viability was examined at indicated time points using the Cell Counting Kit-8 (Dojindo) and by colony formation assay as described previously (24).

Whole-cell lysates were prepared from the cells with radioimmunoprecipitation assay–like buffer, and total protein was transferred to nitrocellulose membrane. The membrane was probed with anti-JMJD2B antibody (H-200; Santa Cruz Biotechnology), anti-JMJD2B antibody (A301-478A; Bethyl Laboratories), anti-H3K9me3 (ab8898; Abcam), and anti-FLAG antibody (F7425; Sigma). ACTB (I-19; Santa Cruz Biotechnology) or histone H3 (ab1791, Abcam) was used to ensure equal loading and transfer of proteins. Protein bands were detected by incubating with HRP-conjugated antibodies (GE Healthcare) and visualizing with Enhanced Chemiluminescence (GE Healthcare).

NIH3T3 cells, cultured in DMEM 10% FBS, were transfected with a p3xFLAG-JMJD2B wild-type vector or a p3xFLAG-JMJD2B ΔJmjC mutant vector. The transfected NIH3T3 cells were cultured for 2 days and seeded in 10-cm dish at the density of 10,000 cells per 10-cm dish in triplicate. Subsequently, the cells were cultured in DMEM 10% FBS containing 0.9 (mg/mL) geneticin/G-418 for 2 weeks until colonies were visible. Colonies were stained with Giemsa (Merck) and counted by Colony Counter software.

A 5′-bromo-2′-deoxyuridine (BrdU) flow kit (BD Pharmingen) was used to determine the cell-cycle kinetics and to measure the incorporation of BrdU into DNA of proliferating cells (25, 26). The assay was done according to the manufacturer protocol. Briefly, cells (2 × 10⁵ per well) were seeded overnight in 6-well tissue culture plates and treated with an optimized concentration of siRNAs in medium containing 10% FBS for 72 hours, followed by addition of 10 μmol/L BrdU and incubations continued for an additional 30 minutes. Both floating and adherent cells were pooled from triplicate wells per treatment point, fixed in a solution containing paraformaldehyde and the detergent saponin, and incubated for 1 hour with DNase at 37ºC (30 μg per sample). Fluorescein isothiocyanate (FITC)-conjugated anti-BrdU antibody (1:50 dilution in wash buffer; BD Pharmingen) was added and incubation continued for 20 minutes at room temperature. Cells were washed in wash buffer and total DNA was stained with 7-amino-actinomycin D (7-AAD; 20 μL per sample), followed by flow cytometric analysis using FACScan (Beckman Coulter) and total DNA content (7-AAD) was determined using CXP Analysis Software Ver. 2.2 (Beckman Coulter).

Microarray analysis to identify downstream genes were done as described previously (21). Purified total RNA was labeled and hybridized onto Affymetrix GeneChip U133 Plus 2.0 oligonucleotide arrays (Affymetrix) according to the manufacturer instructions. Probe signal intensities were normalized by RMA and Quantile (using R and Bioconductor). Because we could also confirm the microarray data of several randomly selected candidate genes using quantitative real-time PCR (Supplementary Fig. S4D), we assume our microarray data are reliable.

Chromatin immunoprecipitation (ChIP) assays were done using ChIP Assay kit (Millipore) according to the manufacturer protocol. Briefly, the fragment of JMJD2B and chromatin complexes was immunoprecipitated with anti-FLAG antibody 48 hours after transfection with pCAGGS-n3FC (mock) and pCAGGS-n3FC-JMJD2B vectors into 293T cells. After the bound DNA fragments to JMJD2B were eluted, a standardized amount was subjected to quantitative real-time PCR reactions. Primer sequences are shown in Supplementary Table S1.

We first examined expression levels of a number of jumonji histone demethylase genes in a small subset of clinical bladder cancer samples and found significant differences in expression levels between normal and cancer cells for the JMJD2B gene (data not shown). Then, we analyzed 76 bladder cancer samples and 20 normal control samples (British) and found significantly higher expression levels of JMJD2B in bladder cancer tissues compared with normal bladder (P < 0.0001, Mann–Whitney U test; Fig. 1A and B). Subclassification of tumors according to tumor grade, metastasis status, recurrence status, and smoking history identified no significant differences, whereas gender was significant (P < 0.0001, Mann–Whitney U test; Supplementary Table S3). Importantly, JMJD2B was notably overexpressed at an early stage of bladder cancer (Supplementary Fig. S1). Next, we evaluated JMJD2B protein expression levels in bladder tissues. After confirming the specificity of the antibody we used (Supplementary Fig. S2), we carried out immunohistochemistry using the specific JMJD2B antibody. This experiment showed strong nuclear staining of JMJD2B, specifically in bladder cancer tissues, whereas no significant staining was observed in normal bladder tissues (Fig. 1C). Among 29 bladder cancer tissue sections, 20 showed positive staining of JMJD2B (70.0%; Supplementary Table S4). In addition, our microarray expression analysis of a number of clinical samples indicated that JMJD2B was overexpressed in lung cancer (Fig. 2A and B). According to immunohistochemical analysis using clinical lung tissues, nuclear staining of JMJD2B was observed in cancer tissues, although no significant nuclear staining was evaluated in normal lung and placental tissues (Fig. 2B). Of 63 lung cancer cases, JMJD2B stained positively in 24 cases (38.1%; Supplementary Table S5). Together, expression levels of JMJD2B in bladder and lung cancer tissues are significantly higher than those in corresponding nonneoplastic tissues at the mRNA and protein levels.

To determine the significance of JMJD2B in human carcinogenesis, we examined whether JMJD2B is involved in the growth regulation of cancer cells. After confirming the elevated expression of JMJD2B in bladder and lung cancer cell lines, compared with the human airway epithelial cell line SAEC and the colonic fibroblast cell line CCD-18Co at the protein level (Fig. 3A), we inhibited JMJD2B expression in SW780 and A549 cells using 2 different siRNA oligonucleotide duplexes (Supplementary Table S2). As shown in Fig. 3B, the specific siRNAs clearly knocked down JMJD2B expression, and subsequently, we carried out a cell growth assay 72 hours after treatment with the same siRNAs. JMJD2B knockdown significantly suppressed growth of bladder and lung cancer cells (Fig. 3C), and this result was also confirmed by a colony formation assay (Fig. 3D). Similarly, growth suppression effect was confirmed in the other bladder cancer cell line SCaBER (Supplementary Fig. S3A and B). We then conducted BrdU and 7-AAD staining to analyze the detailed cell-cycle status of cancer cells after the knockdown of JMJD2B and confirmed that the proportion of cancer cells at the S phase was significantly decreased (Fig. 4A). Concomitantly, the percentage of cancer cells at G₁ phase significantly increased, indicating that JMJD2B could be a critical factor in the regulation of the G₁–S transition in cancer cells. These results suggested that JMJD2B plays an essential role in the growth regulation of cancer cells by modulating the G₁–S transition.

To examine the oncogenic activity of JMJD2B in more detail, we conducted a clonogenicity assay. A JMJD2B expression vector was constructed, capable of driving production of active enzyme within transfected cell lines (Fig. 4B). A wild-type JMJD2B (JMJD2B Wt) vector and an enzyme-dead JMJD2B (JMJD2B ΔJmjC) version of our vector were transfected into separate cultures of NIH3T3 cells. A clonogenicity assay was done on each culture (Fig. 4C; Materials and Methods). Wild-type JMJD2B protein showed stronger oncogenic activity than enzyme-dead JMJD2B protein; therefore, it is the demethylase activity of JMJD2B that promotes oncogenesis in cells. Because JMJD2B is overexpressed at an early stage in cancer progression, JMJD2B seems to play a crucial role in human carcinogenesis.

We next analyzed downstream genes and pathways associated with JMJD2B to clarify the mechanism by which JMJD2B regulates cell-cycle progression. After treatment with JMJD2B siRNA in SW780 and A549, we carried out Affymetrix GeneChip analysis. To exclude the secondary effects of growth suppression, we picked statistically significant signals which were commonly decreased at 12 and 24 hours after siRNA treatment. The knockdown of JMJD2B was clearly confirmed at the protein level (Fig. 5A, right top). Among candidate genes, we observed a significant downregulation of cyclin-dependent kinase 6 (CDK6), one of the key regulators for the G₁–S transition (Fig. 5A, left and right bottom and Supplementary Fig. S4A; ref. 27). Quantitative real-time PCR also confirmed a significant downregulation of CDK6 at both 12 and 48 hours following JMJD2B siRNA treatment (Fig. 5B and Supplementary Fig. S4B). Likewise, downregulation of CDK6 after JMJD2B knockdown was confirmed at the protein level (Supplementary Fig. S4C). In addition, we confirmed a significant elevation of CDK6 in HeLa cells transfected with pCAGGS-n3FC-JMJD2B compared with mock-treated cells (Fig. 5C). To evaluate the possibility that JMJD2B directly regulates CDK6 expression at the transcriptional level, we conducted a ChIP assay. JMJD2B protein was highly enriched at the promoter region of CDK6 after transfection with a pCAGGS-n3FC-JMJD2B vector, accompanied by decreased levels of H3-K9 trimethylation in the region (Fig. 5D). Consistently, signal pathway analysis using the Gene Ontology database indicated that JMJD2B can regulate cell-cycle checkpoint pathways, as well as other important pathways involved in carcinogenesis (Supplementary Table S6). The data revealed that JMJD2B directly activates the expression of CDK6 through demethylation of histone H3 at lysine 9.

Because it is well-known that p16INK4a is an important regulator of CDK6 (28–32), we examined the relationship between p16INK4A and JMJD2B in regulating CDK6. To compare expression levels of p16INK4a, JMJD2B, and CDK6, we conducted quantitative real-time PCR analysis on 14 different cell lines (Fig. 6A). Although p16INK4A is considered an important inactivator of CDK6, we did not find a significant inverse correlation between p16INK4a and CDK6 expressions; instead, a slight positive correlation was observed (Fig. 6B, left). This implies that p16INK4a is unlikely to regulate transcription levels of CDK6 to inactivate the cell-cycle dependent-kinase. On the contrary, expression levels of JMJD2B and CDK6 were significantly positively correlated in 14 cell lines (Fig. 6B, right), which is consistent with our results (Fig. 5). Intriguingly, expression levels of JMJD2B and CDK6 were strongly correlated in cell lines expressing low levels of p16INK4a (Fig. 6C; r = 0.976, P < 0.0001). According to our data, JMJD2B-dependent transcriptional regulation of CDK6 seems to be affected by expression levels of p16INK4a.

Among posttranslational modifications on histones, methylation has been shown to be involved in transcriptional regulation, such as X-inactivation or genomic imprinting (1, 2). Although it had been unclear whether histone methylation marks could be reversed, LSD1, a flavin-dependent amine oxidase histone demethylase, was identified as the first histone demethylase (4). Unlike LSD1, the JmjC family, harboring the conserved JmjC domain, require Fe (II) and α-ketoglutarate to exert demethylase activity, and this demethylation process results in the generation of formaldehyde and succinate (7). Among these enzymes, the JMJD2 family demethylate H3K9 and H3K36 (6), and JMJD2B selectively removes H3K9me2 and H3K9me3 (15, 17). Functional studies showed that JMJD2B contains hypoxia response elements in its promoter region and, therefore, JMJD2B is induced by Hypoxia-inducible factor (HIF; refs. 16, 17), but its significance in human diseases such as cancer remains to be elucidated. In this study, we showed significant upregulation of JMJD2B in various types of cancers, including bladder and lung, by quantitative real-time PCR, cDNA microarray, or immunohistochemistry. In addition, a series of our experiments clarified that JMJD2B serves a critical role in the growth regulation of cancer cells, especially at the G₁–S transition (Fig. 3 and Supplementary Fig. S3). This JMJD2B-dependent cell-cycle regulation could be mediated by the downstream gene CDK6 through the demethylation of histone H3-K9 at the promoter region (Fig. 5), indicating that the enzyme activity of JMJD2B may be an important regulator for the G₁–S transition in cancer cells. Expression analysis examining the correlation between JMJD2B and the histone methyltransferase G9a, catalyzing histone H3-K9 methylation, showed that G9a expression was not changed after treatment with JMJD2B siRNA (Supplementary Fig. S5). We interpret this to mean that although both proteins regulate the methylation status of histone H3K9, the target genes may be different. In fact, we have recently published a paper analyzing the significance of G9a deregulation in human carcinogenesis (33). We reported that G9a is overexpressed in various types of cancer, similarly to JMJD2B. Our functional analyses showed that G9a and JMJD2B may regulate different genes (our previous data in ref. 33 and Supplementary Table S6). Therefore, the histone demethylase JMJD2B and the histone methyltransferase G9a both regulate the status of histone H3K9 methylation and play important roles in human carcinogenesis but may regulate different pathways.

CDK6 is a member of the cyclin-dependent protein kinase family and a catalytic subunit of the protein kinase complex that is important for cell-cycle regulation. This kinase promotes cell-cycle progression from G₁ phase to S phase through promoting RB phosphorylation and subsequent detachment of E2F1 from RB (27). CDK6 has been reported to possess various physiologic functions like T-cell development (34, 35). Importantly, our microarray data showed the aberrant expression of CDK6 in several types of human tumors, including bladder and lung cancers (Supplementary Table S7), and previous reports also described the involvement of this gene in human carcinogenesis (27, 35, 36). This study shows that CDK6 is transcriptionally activated by JMJD2B through demethylation of H3K9 and that elevated CDK6 is likely to promote cell malignancy. This is a novel example of how deregulated histone demethylation contributes to human carcinogenesis. To date, the tumor suppressor gene p16INK4a has been considered as an important negative regulator of CDK6 through direct interaction (30, 37). We found that an inverse correlation was not observed between p16INK4a and CDK6 expressions, and that JMJD2B expression was strongly correlated to CDK6 expression, especially in cell lines expressing low levels of p16INK4a. These results indicate that expression levels of p16INK4a are likely to be a regulator of JMJD2B-dependent CDK6 transcriptional activation. Further functional analysis is required to clarify the crosstalk between the JMJD2B-CDK6 and the p16INK4a-RB1 pathways.

Detailed expression analysis showed that expression levels of JMJD2B in bladder and lung cancers are significantly higher than those in corresponding normal tissues, and knockdown of JMJD2B resulted in the significant suppression of cancer cell proliferation. To explain the mechanism for increased expression of JMJD2B in human cancers, we looked at the genome annotation across the JMJD2B region, spanning Ch19 4,806,367 to 5,154,791 using the 1 Mb CGH array data of bladder cancer tissues. There are no single gains or amplifications; therefore, we cannot confirm there is a gain relating to gene dosage in bladder cancer. Beyer and colleagues previously reported that the HIF-1α binds to specific recognition sites in the gene encoding JMJD2B and induces its expression (17). All human tumors display genomic instability, aberrant transcriptional programs, and, very often, contain areas that are insufficiently perfused, resulting in a local shortage of nutrients and oxygen (hypoxia). This leads to an activation of the transcription factor HIF, the master regulator of oxygen homeostasis (38). These results imply that low-oxidation concentration in cells seems to be an important mediator in activating JMJD2B expression in human carcinogenesis. In addition, the Oncomine database (39) implies that JMJD2B is overexpressed in various types of cancer (Supplementary Fig. S6), so deregulation of JMJD2B plainly contributes to carcinogenesis in a variety of human tumor types. Importantly, expression levels of JMJD2B in various types of normal tissues are significantly low (Supplementary Fig. S7). These results indicate JMJD2B as an ideal target for cancer therapy. Our functional analyses were mainly based on in vitro models, so additional data from animal experimental models may reinforce the importance of JMJD2B as a therapeutic target in human cancer. Recently, inhibitors targeting DNA methyltransferases and histone deacetylases were approved by the FDA to use for patients with hematologic malignancies (40, 41). Furthermore, a novel inhibitor targeting the histone demethylase LSD1 has been developed and a suppressive effect on tumor growth, via reexpression of epigenetically silenced genes in colon cancer cells, was shown (42). On the basis of these results, it seems feasible to develop specific inhibitors targeting JMJD2B as anticancer agents. Because the development of inhibitors targeting histone methyltransferases and demethylases has begun, these reagents should be further validated against the functions of this enzyme, to assure the usefulness of this approach in the near future.

No potential conflicts of interest were disclosed.

The authors thank Professor Gillian Murphy and the members of her laboratory for substantial technical support and also thank Ms. Yuka Yamane and Ms. Haruka Sawada for technical assistance.

This work was supported by a grant-in aid for young scientists (A; 22681030) from the Japan Society for the Promotion of Science. Our biorepository is supported by funding NIHR and the Cambridge Biomedical Research Centre.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.