Histone deacetylase inhibitors deplete enhancer of zeste 2 and associated polycomb repressive complex 2 proteins in human acute leukemia cells

During development, the polycomb group (PcG), together with trithorax group of proteins, play an important role in the regulation of expression of homeotic (HOX) genes (1, 2). Additionally, deregulated expression and function of PcG proteins has been linked to altered cellular proliferation and cancer (3, 4). PcG proteins function in multimeric protein complexes and repress HOX genes (2–4). Two of the best-characterized PcG complexes are the polycomb repressive complex (PRC) PRC1 and PRC2 (2, 3). These complexes localize at specific sites called polycomb repressive elements and organize the chromatin into a repressive structure that is unresponsive to chromatin remodeling factors and basal- and gene-specific transcription factors (2–4). Of the two complexes, PRC2 is smaller and contains the PcG proteins enhancer of zeste 2 (EZH2), embryonic ectoderm development (EED) that serves to recruit to the chromatin the PRC1 complex, suppressor of zeste 12 (SUZ12), Yin Yang 1, and the histone-binding protein RbAp46 (2–5). Recently, within the PRC2, EZH2 was shown to interact with and modulate the DNA methyltransferases DNMT1, DNMT3a, and DNMT3b, which affect their binding to the EZH2 target gene promoters (6, 7). Thus, EZH2 is required for DNA methylation of EZH2-targeted promoters. EZH2 may serve as a recruitment platform for DNA methyltransferases, thereby mechanistically linking the two epigenetic silencing systems (6, 7). EZH2 is crucial during embryonic development, as depletion of EZH2 from developing mouse embryos resulted in severe growth retardation and early embryonic lethality (8, 9). EZH2 contains an evolutionarily conserved, COOH-terminal Su(var)3-9, enhancer of zeste, and trithorax domain, which mediates the histone methyltransferase (HMTase) activity specific for lysine (K)-27 of histone H3 (H3K27) that is associated with gene repression (10–12). Besides its catalytic HMTase domain, EZH2 contains highly conserved NH₂-terminal protein interaction domains responsible for binding to EED and a nuclear localization domain (11). EED recruits HDAC activity to the PRC2 complex (12).

There is accumulating evidence that deregulated expression of PcG proteins is involved in cellular transformation and promotes aggressiveness and high proliferation rates in cancers (13, 14). EZH2 overexpression has been especially linked to aggressive tumor formation and poor prognosis in both prostate and breast cancers (15–18). In a subset of cancers, EZH2 locus is amplified resulting in EZH2 overexpression (16–18). This can promote cellular transformation and confer invasiveness in immortalized breast epithelial cells (16). With SUZ12 and EED, EZH2 has been established as the minimum functional PRC2 core complex exerting gene silencing through methylation of H3K27 and is transcriptionally regulated by the pRB/E2F pathway (3, 13, 19). Activated p53 has also been shown to suppress EZH2 expression (20). Recently, AKT was shown to phosphorylate EZH2 at Ser²¹ and suppresses its methyltransferase activity by impeding EZH2 binding to histone H3 (21). This results in decreased levels of trimethylated H3K27 and derepression of the silenced genes. EZH2 has also been reported to be up-regulated in mantle cell lymphoma as well as coexpressed with BMI-1 (a polycomb protein in the PRC1 complex) in Reed-Sternberg cells in Hodgkin's disease and non-Hodgkin's lymphoma (22–24). Whereas normal bone marrow plasma cells do not express EZH2, during disease progression in primary multiple myeloma, EZH2 expression is induced and correlates with the tumor burden (25). Additionally, a recent report showed that growth factor independence in multiple myeloma cells with N- or K-Ras mutation requires the activity of EZH2. It was also shown that the in vivo oncogenic activity of EZH2 depends on the function of its Su(var)3-9, enhancer of zeste, and trithorax domain (25).

Although overexpression of EZH2 has been reported to increase aggressiveness in the various types of cancers, EZH2 expression has not been evaluated previously in acute leukemia. The PRC2 complex has been shown to bind to the promoter and regulate the expression of homeobox A9 (HOXA9), a gene linked to poor prognosis in acute myelogenous leukemia (AML; ref. 3). HOXA9 is the most frequent partner in a fusion oncoprotein with Nup98 (Nup98-HOXA9), resulting from the chromosomal translocation t(7;11)(p15;p15) observed in human leukemia (26–28). HOXA9 is also known to interact with myeloid ecotropic viral integration site 1 homologue (MEIS1), a member of the three–amino acid loop extension subclass of homeodomain-containing proteins (29). The MEIS1-HOXA9 interaction has been reported to rapidly accelerate leukemogenesis in mice (30–32), making these proteins and their interaction a potential therapeutic target for further evaluation. Additionally, in the PRC2 complex, EZH2 is known to interact with histone deacetylases (HDAC) 1 and 2 through the EED protein (13, 33, 34). This suggests that based on the cellular context, the transcriptional repression by the PRC2 complex may be mediated through the activity of the HDACs (18). Therefore, in the present studies, we determined the effect of the hydroxamate HDAC inhibitors (HA-HDI) LBH589 and LAQ824 on EZH2 and PRC2 complex proteins and their HMTase activity in cultured leukemia cell lines K562, U937, and HL-60 as well as in primary human chronic myelogenous leukemia (CML) and AML cells. We also determined the effects of small interfering RNA (siRNA)–mediated knockdown of EZH2 and/or HA-HDI on EZH2 mRNA and protein levels as well as on the clonogenic survival and differentiation of human AML cells.

LAQ824 and LBH589 were kindly provided by Novartis Pharmaceuticals, Inc. (East Hanover, NJ). Anti-EZH2 monoclonal antibody was purchased from BD Transduction Laboratories (San Jose, CA). Polyclonal anti-SUZ12, anti-EED, anti-HOXA9, Anti-MEIS1/MEIS2/MEIS3, monoclonal anti–trimethylated H3K27, polyclonal anti–dimethylated H3K27, polyclonal anti–acetylated H3K27, polyclonal anti–acetyl histone H3, and polyclonal anti–acetyl histone H4 antibodies were purchased from Upstate (Lake Placid, NY). Anti-p21 antibody was purchased from NeoMarkers (Fremont, CA). Anti–β-actin was purchased from Sigma-Aldrich (St. Louis, MO). Anti-pRb antibody was purchased from Cell Signaling (Beverly, MA).

K562, HL-60, and U937 cells were cultured in complete RPMI 1640 and incubated at 37°C with 5% CO₂ as described previously (35). Medium was changed every 2 to 3 days, and cells were passaged at a density of 0.25 × 10⁶/mL. Logarithmically growing cell cultures were used for all experiments described below.

Primary AML and CML cells were obtained with informed consent as part of a clinical protocol approved by the Institutional Review Board of the University of South Florida. As described previously (36, 37), peripheral blood samples were collected in heparinized tubes or obtained from leukophoresis units, and mononuclear cells were separated using Lymphoprep (Axis-Shield, Oslo, Norway), washed once with complete RPMI 1640, resuspended in complete RPMI 1640, and counted to determine the number of cells isolated before their use in the various experiments. The purity of the blast population was confirmed to be 80% or better by morphologic evaluation of cytospun cell preparations stained with Wright stain.

Untreated and siRNA or drug treated for 48 h, cells were harvested by centrifugation at 1,000 rpm in a swinging bucket rotor for 5 min. Cell pellets were washed with 1× PBS, gently resuspended in 200 μL lysis buffer [1% Triton X-100, 1 mmol/L phenylmethylsulfonyl fluoride, 10 μg/mL leupeptin, 1 μg/mL pepstatin-A, 2 μg/mL aprotinin, 20 mmol/L p-nitrophenyl phosphate, 0.5 mmol/L sodium orthovanadate, 1 mmol/L 4-(2-aminoethyl) benzenesulfonylfluoride hydrochloride], and incubated on ice for 30 min as described previously (38). Cell lysates were centrifuged at 12,000 rpm in a tabletop centrifuge for 15 min to remove the nuclear and cellular debris. An aliquot of each cell lysate was diluted 1:10 and quantitated using a bicinchoninic acid protein quantitation kit according to the manufacturer's protocol. Known concentrations of bovine serum albumin were used to establish a standard concentration curve. Proteins were loaded into a 96-well plate and the plate was incubated at 37°C for 20 to 30 min. Protein concentrations were read on a Bio-Rad (Hercules, CA) plate reader using microplate manager version 5.5.

One hundred micrograms of total cell lysate were used for SDS-PAGE. Western blot analyses of EZH2; SUZ12; EED; HOXA9; MEIS1; trimethylated, dimethylated, or acetylated H3K27; p21; α-tubulin; and β-actin were done on total cell lysates using specific antisera or monoclonal antibodies as described previously (35–38). The expression level of either β-actin or α-tubulin was used as the loading control for the Western blots. Blots were developed with a chemiluminescent substrate enhanced chemiluminescence (Amersham Biosciences, Piscataway, NJ).

For siRNA-mediated down-regulation of EZH2, an EZH2-specific SMARTPool was purchased from Dharmacon (Lafayette, CO). Nonspecific control siRNA was purchased from Ambion (Austin, TX). All siRNA experiments were done at a final concentration of 100 nmol/L duplex siRNA. For nucleofection into K562, U937, or HL-60 cells, Nucleofector Kit V was used (Amaxa, Gaithersburg, MD). For nucleofection, 5 million low-passage cells were mixed with duplex siRNA, and cell-specific nucleofection program (Amaxa) was used to deliver the siRNA duplex into the nucleus of the cells. Following nucleofection, the cells were incubated overnight for recovery. Following this, cells were treated with HDIs for Western blot analyses or cell cycle effects.

Histones were isolated by a modification of a previously described method (39). After the designated treatments, cells were harvested by centrifugation at 700 × g and incubated on ice-cold histone isolation buffer [10 mmol/L Tris-HCl, 50 mmol/L sodium bisulfite, 1% Triton X-100, 10 mmol/L MgCl₂, 8.6% sucrose (pH6.5)] on ice for 30 min. Cells were Dounce homogenized using a type B pestle. Released nuclei were pelleted, supernatant was removed, and the nuclei were washed briefly with the histone lysis buffer. The nuclei were resuspended in 100 μL ice-cold H₂O, 1.2 μL of concentrated H₂SO₄ were added, and the tubes were incubated on ice for 1 h. The acid-treated nuclei were centrifuged for 5 min at 15,000 rpm. The supernatants were removed to a clean microcentrifuge tube, and 1 mL acetone was added. Histones were precipitated from the acid extracts overnight at −20°C. The extracted histones were centrifuged briefly at 10,000 rpm, air dried, and resuspended in 50 μL water. Protein concentrations were quantitated as described above. For Western blot, 3 to 5 μg of purified histones were used per condition.

RNA was extracted from the cultured and primary cells using the Trizol method (Invitrogen, Carlsbad, CA). Purified RNA was quantitated and reverse transcribed using SuperScript II according to the manufacturer's protocol (Invitrogen). Resulting cDNAs were used in subsequent PCRs for EZH2, SUZ12, and EED (primer sequences available on request). PCRs for β-actin were used as an internal loading control for the PCRs. PCRs were carried out in a gradient Mastercycler (Eppendorf, Westbury, NY) and consisted of a 3-min denaturation at 95°C followed by 30 cycles of 95°C (30 sec), 52°C (30 sec), and 72°C (30 sec) with a final extension at 72°C (10 min). Amplified products were resolved on a 2% agarose gel and recorded with a UV/VIS gel box. Horizontal scanning densitometry was done with ImageQuant 5.2, and band intensity was compared with β-actin.

Following the designated treatments, cells were harvested and washed twice with 1× PBS and fixed in ethanol overnight. Fixed cells were washed twice with 1× PBS and stained with propidium iodide for 15 min at 37°C. Cell cycle data were collected on a flow cytometer with a 488 nmol/L laser and analyzed with ModFit 3.0 as described previously (35–38).

Following the designated treatments, cells were harvested and washed twice with 1× PBS and ∼500 cells were plated in complete Methocult (Stemcell Technologies, Vancouver, British Columbia, Canada) and cultured for 7 to 10 days at 37°C in a 5% CO₂ environment. Colony growth was measured as a percentage of the control cell colony growth as described previously (38).

U937 cells were transfected with siRNA over 48 h as described above. Cells were washed with RPMI 1640 and treated with 5 nmol/L LBH589 for 5 days. Following this, cells were centrifuged at 1,000 rpm for 5 min and resuspended in 100 μL of 5% bovine serum albumin/PBS for 15 min. Following this, 5 μL Alexa 488–conjugated anti-CD11b antibody (BD PharMingen, San Diego, CA) and 10 μL propidium iodide (1:100 dilution) were added, and the cells were incubated on ice for 20 min. The percentage of CD11b-expressing cells was determined by flow cytometry. In addition, the percentage of morphologically differentiated cells was estimated using light microscopy of at least 200 Wright-stained cytospun cells. Morphologic criteria for differentiation included the presence of all of the following features: condensation of chromatin, indentation and lobation of the nucleus, as well as decreased basophilia and increased amount and eosinophilic granulation of the cytoplasm.

K562 cells were transfected with either control or EZH2 siRNA for 48 h. Following this, cells were first washed once with 1× PBS and subsequently washed in 1× PBS containing 10 μmol/L Na₃VO₄ and 50 μmol/L NaF. Cells were lysed in 1× hypotonic buffer, and nuclear extracts were prepared as described previously (40). To determine the HMTase activity, a previously described method was used (41). Five microgram of the nuclear extracts were combined with 2 μL of 5× HMTase buffer [250 mmol/L Tris (pH 9.0), 2.5 mmol/L DTT, 5 mmol/L phenyl-methylsulfonyl fluoride; Upstate], 0.55 μCi S-adenosyl-L-methionine, 1 μg recombinant histone H3, and diH₂O up to 10 μL. This mixture was gently mixed and incubated at 30°C for 30 min. Following this, 5 μL of each reaction were transferred to p81 phosphocellulose squares and air dried. The assay squares were washed thrice for 15 min with 10% trichloroacetic acid followed by one wash with 95% ethanol for 5 min. The assay squares were air dried and mixed with scintillation fluid, and the activity was measured with a scintillations counter. The HMTase activity of the samples was subtracted from the activity of a control sample without the nuclear extract.

We first determined the effects of the pan-HDIs LBH589 and LAQ824 on the levels of the core PRC2 complex components (i.e., EZH2, SUZ12, and EED) in the cultured CML K562 and LAMA-84 cells. Exposure to increasing concentrations of LBH589 or LAQ824 in a dose-dependent manner decreased the levels of EZH2, SUZ12, and EED in K562 cells (Fig. 1A). Whereas the depletion of SUZ12 was pronounced, EED was only modestly depleted (Fig. 1A). These effects were noted even after a 4-h exposure to 100 nmol/L LBH589 (Fig. 1B). Similar effects were observed in the cultured AML U937 cells (Fig. 1C). We next determined the effects of LBH589 and LAQ824 on the PRC2 complex components in primary CML and AML cells. Treatment with LBH589 again depleted EZH2 and SUZ12 more than EED in a sample each of primary CML-BC and AML cells (Fig. 1D and E). Similar effects were observed after exposure to LAQ824 (data not shown).

We next determined whether LBH589-mediated depletion of the components of the PRC2 complex is associated with the loss of H3K27 methyltransferase activity and decreased levels of trimethylated and dimethylated bulk H3K27. K562 cells were treated with LBH589 for 24 h and histones were extracted. As shown in Fig. 2A, exposure to LBH589 depleted the levels of trimethylated and dimethylated H3K27, with concomitant increase in the level of acetylated H3K27, suggesting a possible reciprocal relationship between acetylation and methylation status of H3K27; however, it is possible that these events may be unrelated. As was reported previously, LBH589 also induced the overall lysine acetylation of histone H3 and H4 (36). Similar effects on H3K27 methylation and acetylation were also observed in U937 cells (data not shown). PRC2 complex–mediated methylation of H3K27 is known to regulate the expression of HOXA9. Therefore, we determined the effects of LBH589 on HOXA9 as well as on MEIS1 because these two proteins collaborate in the pathogenesis of AML (30, 31). In K562 and U937 cells, exposure to LBH589 in a dose-dependent manner down-regulated HOXA9 and MEIS1 expressions (Figs. 2B and C). LAQ824 exerted a similar effect in U937 cells (data not shown). Both drugs also depleted HOXA9 and MEIS1 levels in two primary AML samples (Fig. 2D).

We next determined the effects of EZH2 siRNA on PRC2 complex and its activity in K562 cells. Transfection of EZH2 siRNA into the K562 cells resulted in down-regulation of the EZH2 mRNA levels in 24 h by ∼60%, without a similar effect on the mRNA levels of SUZ12 or EED (Fig. 3A). However, treatment with EZH2 siRNA not only decreased the protein levels of EZH2 but also resulted in modest depletion of the levels of SUZ12 and EED (Fig. 3A). To determine the combined effects of EZH2 siRNA and treatment with LBH589, we transfected K562 cells with the control or EZH2 siRNA over 48 h with or without the concurrent treatment with 100 nmol/L LBH589 during 24 to 48 h. Exposure to LBH589, along with the control siRNA, again depleted the levels of SUZ12, with less reduction in the levels of EED (Fig. 3B). However, under these conditions of culture, LBH589 was less effective in depleting the levels of SUZ12 and EED. In contrast, cotreatment with LBH589 enhanced EZH2 siRNA–mediated depletion of SUZ12 and EED levels. Concomitantly, siRNA to EZH2 also sensitized the cells to LBH589-mediated induction of p21. Treatment with LAQ824 (250 nmol/L) produced similar effects on the components of the PRC2 complex (data not shown). This suggests that EZH2 siRNA treatment renders K562 cells more sensitive to LBH589- or LAQ824-mediated decline in the levels of EED and SUZ12. We next determined the effects of EZH2 siRNA on HMTase activity in the nuclear extracts of K562 cells. Figure 3C shows that unlike the control siRNA, transfection of EZH2 siRNA produced marked reduction in the H3K27 methyltransferase activity in K562 cells. As was observed following exposure to LBH589, treatment of K562 cells with EZH2 siRNA was also associated with depletion of the levels of trimethylated and dimethylated H3K27, with concomitant increase in the level of acetylated H3K27 (Fig. 3D). We also determined the apoptotic effects of combined treatment of control siRNA or EZH2 siRNA and LBH589. The combination of EZH2 siRNA and LBH589 did not result in greater apoptotic effects than control siRNA-transfected cells treated with LBH589 (data not shown).

We next determined the effects of treatment with EZH2 siRNA and/or the pan-HDI LBH589 on the levels of the components of the PRC2 complex and clonogenic survival of AML cells. HL-60 and U937 cells were nucleofected with 100 nmol/L control or EZH2 siRNA for 48 h with or without the concurrent treatment with LBH589 during 24 to 48 h. EZH2 siRNA treatment alone depleted the protein levels of EZH2, SUZ12, and EED in AML cells (Fig. 4A and B). As was seen for the CML K562 cells (Fig. 3B), cotreatment with LBH589 enhanced EZH2 siRNA–mediated decline in the levels of EZH2, SUZ12, and EED in U937 (Fig. 4A) and HL-60 cells (Fig. 4B). As compared with treatment with either agent alone, cotreatment with LBH589 and EZH2 siRNA also induced more p21 (Fig. 4A). Whereas treatment with EZH2 siRNA alone induced the expression of HOXA9, combined treatment with LBH589 (25 nmol/L) and EZH2 siRNA depleted HOXA9 and MEIS1 levels in U937 cells (data not shown). Notably, compared with treatment with either agent alone, cotreatment with LBH589 and EZH2 siRNA increased the loss of clonogenic survival of HL-60 and U937 cells (Fig. 4C). We next determined the effect of treatment with EZH2 siRNA and/or LBH589 on the differentiation of U937 cells. Differentiation was assessed by morphologic evaluation of Wright-stained cells and induction of CD11b expression by flow cytometry. Figure 5A and B shows that, compared with treatment of U937 cells with EZH2 siRNA alone, cotreatment with LBH589 and EZH2 siRNA resulted in significantly more CD11b expression and more morphologically differentiated cells (P < 0.05). Notably, in the clonogenic survival and differentiation studies, lower concentrations of LBH589 (2.5–5 nmol/L) were used (Figs. 4C and 5).

Findings presented here show that treatment with the HA-HDIs LBH589 and LAQ824 or EZH2 siRNA down-regulates EZH2 and the other core components SUZ12 and EED of the PRC2 complex in human acute leukemia cells. This correlated with decreased HMTase activity of EZH2 and reduced trimethylation and dimethylation of H3K27. The protein levels of PRC2 members are in part dependent on the presence of the other partners in the complex and are unstable outside of a functional PRC2 complex (12, 41). Previous studies have also shown that, although it contains the Su(var)3-9, enhancer of zeste, and trithorax domain for HMTase activity for H3K27, to gain catalytic activity EZH2 requires association with SUZ12 and EED (41). Therefore, the depletion of the PRC2 complex components due to treatment with HA-HDI or EZH2 siRNA results in the attenuation of HMTase activity of EZH2 and decreased trimethylation and dimethylation of H3K27 in the leukemia cells. Because EZH2 recruits class I HDACs through their interaction with EED to PRC2 and H3K27 (33), depletion of the PRC2 complex components by EZH2 siRNA and/or HA-HDI also concomitantly promotes the acetylation of H3K27. Whether the mechanism by which the HA-HDIs, such as LBH589, deplete EZH2 levels is transcriptional, post-transcriptional, or increased protein degradation is not described here but is currently under investigation in our laboratory.

EZH2 is often deregulated in many cancer cell types, including prostate, breast, bladder, cutaneous melanoma, and non-Hodgkin's lymphoma (3, 4, 13–18). Although EZH2 locus is amplified in some cancers (19), EZH2 is also induced by E2F transcription factors (19). Both pRB and p16 repress EZH2 (14, 19). Transcriptionally active p53 has also been shown to suppress EZH2 expression (20). Taken together, these observations explain why EZH2 expression is deregulated in a variety of cancers (14–18). Our preliminary findings show that both cultured and primary AML cells express significant levels of EZH2 protein. This has to be confirmed in a larger sample size of acute leukemia cells. In previous reports, EZH2 overexpression has been shown to increase proliferation, promote invasive potential, and inhibit differentiation (13–17, 34). Notably, EZH2 has oncogenic activity in breast cancer and multiple myeloma cells, where cell transformation and tumor formation requires HMTase activity of EZH2 (16, 25). Conversely, EZH2 down-regulation results in growth arrest and differentiation but not apoptosis (18, 33). In the present studies, we also observed an inhibition of colony growth but not apoptosis of the cultured AML cells following depletion of EZH2 as well as of SUZ12 and EED, by EZH2 siRNA. Treatment with levels of LBH589 and LAQ824 that results in growth arrest, differentiation, and apoptosis was also noted to deplete EZH2, SUZ12, and EED in the CML and AML cells (36, 38). However, the precise mechanistic link between the depletion of the PRC2 complex components and LBH589- or LAQ824-induced growth arrest, differentiation, and loss of clonogenic survival was not established. As noted above, previous reports have shown that EZH2 interacts with and recruits class I HDACs to PRC2 through EED (33). Importantly, EZH2-mediated gene silencing and invasive potential of breast cancers was also noted to be dependent on HDAC activity (16, 18). Our findings are consistent with this in showing that cotreatment with the HA-HDI LBH589 augments EZH2 siRNA–mediated depletion of the PRC2 complex components as well as induction of differentiation. Furthermore, cotreatment LBH589 also enhanced EZH2 siRNA–mediated loss of clonogenic survival of the AML cells.

Although not studied here, the binding to chromatin and gene silencing by PRC1 complex, which consists of >10 subunits, including BMI-1 and the HPC proteins, requires PRC2 complex–mediated trimethylation of H3K27 (6, 42, 43). Notably, PRC1, PRC2, and trimethylated H3K27 cooccupy >1,000 silenced genes, but 40 genes are derepressed in human embryonic fibroblasts depleted of PRC2 components and BMI-1 (44). Of these, HOXA9 is repressed by PRC2 complex–mediated trimethylation of H3K27 and induced by the knockdown of SUZ12 (5). Recently, PRC1 complex has been shown to induce H2AK119 ubiquitin E3 ligase activity, which is associated with the repression of HOX genes (45). Therefore, our findings that treatment with EZH2 siRNA, which depletes the levels of PRC2 components and trimethylation of H3K27, increases the levels of HOXA9 is consistent with these reports. In addition, our additional findings that treatment with HA-HDIs depletes HOXA9 levels are also consistent with the previous report that HDAC activity is essential for the expression of HOXA9 and for the endothelial commitment of progenitor cells (46). We further show that treatment with HA-HDIs alone or cotreatment with EZH2 siRNA depleted HOXA9 and MEIS1 levels but induced p21 in the AML cells. This was associated with significantly diminished clonogenic survival but increased differentiation of the AML cells. Transplantation of murine bone marrow cells retrovirally transduced with HOXA9 or the nucleoporin (NUP98)-HOXA9 fusion leads to AML, which is greatly accelerated by the coexpression of MEIS1 gene (30–32). Therefore, by depleting HOXA9 and MEIS1 levels, cotreatment with EZH2 siRNA and LBH589 may disrupt the biological synergy between the two genes in inducing myeloid leukemia. This suggests that combined treatment with antagonists of PRC2 complex and HA-HDIs may be an attractive targeted therapy against AML with coexpression of HOXA9 and MEIS1.

Whereas BMI-1 is necessary for the self-renewal of hematopoietic stem cells, EZH2 has also been shown to prevent stem cell exhaustion in hematopoietic tissue (47, 48). By down-regulating EZH2 and PRC2, and thereby impairing BMI activity, cotreatment with LBH589 and EZH2 siRNA may also target and deplete leukemia stem cell. However, this has to be directly probed and established. In addition, as noted above, EZH2 can interact with and recruit DNA methyltransferases and thereby promote DNA methylation of EZH2-targeted promoters and their repression (6, 7). Taken together with the findings presented here, these observations create a strong rationale to develop and test the in vivo efficacy of a combination of anti-EZH2 and a HA-HDI against AML because this would abrogate three separate mechanisms of epigenetic silencing that promote growth and suppress differentiation. Our preclinical findings support the development of a targeted therapy against EZH2 for AML. This could potentially be of two kinds. One that inhibits the binding of EZH2 with EED, a critical interaction required for EZH2 HMTase activity. The second could be an inhibitor of the COOH-terminal Su(var)3-9, enhancer of zeste, and trithorax domain of EZH2, which mediates the histone lysine methyltransferase activity of the PRC2 complex. This therapeutic strategy has become more attractive, given the recent approval of the DNMT1 inhibitors decitabine and azacytidine for the treatment of advanced stages of myelodysplastic syndromes that often culminate in AML (49).