International Journal of Hematology

, Volume 96, Issue 1, pp 65–73

Down-regulation of hematopoiesis master regulator PU.1 via aberrant methylation in chronic myeloid leukemia

Authors

  • Hui Yang
    • Department of Hematology, Xin Hua HospitalAffiliated to Shanghai Jiao Tong University (SJTU) School of Medicine
  • Hui Liang
    • Department of Hematology, Xin Hua HospitalAffiliated to Shanghai Jiao Tong University (SJTU) School of Medicine
  • Jing-song Yan
    • Department of HematologyThe Second Affiliated Hospital of Dalian Medical University
  • Rong Tao
    • Department of Hematology, Xin Hua HospitalAffiliated to Shanghai Jiao Tong University (SJTU) School of Medicine
  • Si-guo Hao
    • Department of Hematology, Xin Hua HospitalAffiliated to Shanghai Jiao Tong University (SJTU) School of Medicine
    • Department of Hematology, Xin Hua HospitalAffiliated to Shanghai Jiao Tong University (SJTU) School of Medicine
Original Article

DOI: 10.1007/s12185-012-1106-x

Cite this article as:
Yang, H., Liang, H., Yan, J. et al. Int J Hematol (2012) 96: 65. doi:10.1007/s12185-012-1106-x

Abstract

The PU.1 transcription factor is a crucial regulator of hematopoiesis, and its expression is altered in various leukemic processes. It has been shown that expression of PU.1 is severely impaired in patients with chronic myeloid leukemia (CML), but the mechanism underlying this effect remains unknown. Through bisulfite sequencing, semi-quantitative PCR, and indirect immunofluorescence and Western blot techniques, we found aberrant methylation in the promoter region of transcription factor PU.1 in CML patients both in the chronic and blast crisis phases, as well as in the CML blast K562 cell line. Of these, several CpG sites were more highly methylated in blast crisis than chronic phase, while no methylation of these sites was observed in healthy individuals. Interestingly, CML patients achieved complete cytogenetic remission under imatinib mesylate treatment, but the aberrant methylation status of PU.1 was not reversed. Down-regulation of PU.1 expression at the mRNA and protein levels was also observed in association with aberrant methylation. Thus, for the first time, we have revealed a potential epigenetic modification of PU.1 in CML, which may be responsible for the down-regulation of PU.1. These data suggest that aberrant methylation of PU.1 may play a role in CML pathogenesis, and may therefore serve as a useful biomarker and potential target for demethylating drugs.

Keywords

Chronic myeloid leukemiaTranscription factor PU.1DNA methylation

Introduction

Chronic myeloid leukemia (CML), a malignant clonal disorder of hematopoietic stem/progenitor cells, accounts for approximately 15 % of all adult leukemias. Typically the disease develops in 3 phases. An initial CP is characterized by massive expansion of the granulocytes lasting approximately 3–6 years. The disease then progresses, often through an accelerated phase (AP), to a terminal BC that is manifested by a block of cell differentiation resulting in accumulation of myeloid or lymphoid blasts in peripheral blood (PB) and/or bone marrow (BM) [1]. Treatments including allogeneic hematopoietic stem-cell transplantation and the administration of tyrosine kinase inhibitors such as imatinib mesylate can slow down disease progression and greatly improve clinical outcome, but the advent of blast transformation (AP/BC) remains a challenge for physicians.

PU.1 is a member of Ets family transcription factor which plays an important role in the development of lymphoid and myeloid cells and regulation of expression of lineage-specific genes. Prior studies had investigated that PU.1 was necessary for the development of B cell, T cell, neutrophils and mast cells [2]. In PU.1-deficient mice, B cell, T cell, monocyte and granulocyte show defects in differentiation [3]. Moreover, PU.1 is involved in various hematopoietic malignancies. The overexpression of PU.1 in erythroid precursors induces erythroleukemia in transgenic mice [4], whereas a reduction in the PU.1 expression level is associated to acute myeloid leukemias (AML) [5, 6], indicating that the PU.1 role in leukemic processes depends on its expression levels.

Considering the significance of PU.1 in the procedure of hematopoiesis and that CML blast crisis is the accumulation of immature blast cells, we investigated whether PU.1 is involved in the initiation and progression of CML. In our previous work, we have screened the promotor region and coding sequence of PU.1 in CML patients and found no existing mutations [7]. During the same period, another research group has demonstrated that expression of PU.1 is severely impaired in patients with chronic myeloid leukemia (CML), but its underlying mechanism is still unknown [8].

DNA methylation in promoter region can transform the expression of specific gene [9], and is the most common modification pattern in epigenetics which is the critical mechanism for the expression of genes. In hematologic malignancies, many kinds of tumor suppressor genes and candidate suppressor genes are epigenetically inactivated [10]. Previous investigation had claimed that many genes were inactivated by methylation in CML, such as hPER3, P15, C-ABL, SOCS-1 and JunB [1114]. Here, In our present work, for the first time we found aberrant methylation in the promoter region of transcription factor PU.1 in CML-CP at diagnosis, CML-CP with TKI, CML-BC, as well as CML blast K562 cell line, while totally no methylation was observed in normal individuals counterparts. Thus, we unveiled epigenetic modification of PU.1 in CML patients and K562 cell line, which may help us to further understand the pathogenesis of CML and develop new treatment.

Materials and methods

Patients

Peripheral blood and bone marrow samples were collected from CML patients at Hematology Department, Xin Hua Hospital, Affiliated to Shanghai Jiao Tong University (SJTU) School of Medicine. CML clinical diagnosis and stage system meet with WHO criteria. Peripheral blood and bone marrow samples from healthy volunteers were used as normal controls. Written informed consent was obtained according to the Declaration of Helsinki.

Cell culture

K562 cells were cultured in RPMI 1640 (Gibco) supplemented with 10 % fetal bovine serum (Invitrogen), 100 U/mL penicillin, and 100 μg/mL streptomycin, and grown at 37 °C with 5 % CO2.

DNA extraction

DNA was extracted from peripheral blood and bone marrow samples using Wizard Genomic DNA purification Kit (Promega), according to the manufacturer’s instruction.

Bisulfite treatment

Bisulfite treatment of informative DNA samples was performed according to the following protocol: 3–4 μg of genomic DNA was denatured in 30 μL of 3 M NaOH for 20 min at 55 °C. Then, 666 μL of freshly prepared 10 mM hydroquinone (Sigma) was added onto the sodium bisulfite (Sigma), 900 μL of the mixture at pH 5.0 were added onto the DNA mixture. The samples were overlaid with mineral oil and incubated at 55 °C for 6 h. The bisulfite-treated DNA was purified using Wizard DNA Clean-Up System (Promega). By adding 50 μL of warm water and 4.0 μL of 3 M NaOH for 15 min at 37 °C DNA was eluted. The DNA was then precipitated overnight with ammonium acetate and ethanol and glycogen, centrifuged, and washed with 70 % ethanol before being resuspended in 50 μL of ddH2O.

PCR and bisulfite sequencing

Bisulfite-modified DNA was amplified by PCR using the primer designed for detection of methylation status in promoter region of PU.1 gene as follows: 5′-GAG ATT TTT TGT ATG TAG TGT AAGA-3′ (forward), 5′-TAA CTT CCC ACT AAT AAC AAA CCA-3′ (reverse) [15], using the housekeeping gene GAPDH as internal control and the primer sequences 5′-TTT ATG ATA ATT TTG GTA TTG TG-3′ (forward) and 5′-TATCAC TAT TAA AAT CAA AAA-3′ (reverse). PCR were conducted under the following conditions: 10 μL bisulfite-treated DNA, 2.5 μL 10× PCR buffer, 2 μL dNTP mixture and 0.5 μL of each primer in a final volume of 25 μL. Cycling conditions were as follows: 5 min at 95 °C, followed by 40 cycles of 30 s at 94 °C, 30 s at 55 °C (PU.1)/49 °C (GAPDH), and 1 min at 72 °C, and a final 7-min extension step at 72 °C. The PCR products were purified using QIAquick Gel Extraction Kit (QIAGEN) according to the manufacturer’s instructions. The purified DNA was subcloned into the pGEM-T-EASY vector (Promega). Clones for each phase and cell line were sequenced for detection of methylation status in promoter region.

RNA extraction

Total RNA was extracted from the BMNCs of CML patients by the guanidinium thiocyanate/acid phenol method using Trizol reagent (Invitrogen, USA) in accordance with the manufacturer’s standard method.

Semiquantitative reverse transcription-PCR

2 μg of total RNA was reverse transcribed into cDNA using random primers and Oligo dT(TaKaRa), 20 μL RT reaction was performed at 37 °C for 15 min, then at 85 °C for 5 s. cDNA was stored at −20 °C for use. Semiquantitative PCR analysis was done with the following primers: 5′-AAA GTC ATC CCT CTC AGT CCC AGC TTCC-3′ and 5′-TCT CAG GCC AGGAGT TCC AGG GAG GAAA -3′ (PU.1) and 5′-TCC ATG ACA ACT TTGGTA TCG TG-3′ and 5′-TGT CGC TGT TGA AGT CAG AGGA-3′ (GAPDH). PCR was conducted under following conditions: 1 μL cDNA, 2.5μL 10× PCR buffer, 2 μL dNTP mixture, 0.125 μL TaKaRa Taq and 0.5 μL of each primer in a final volume of 25 μL. Reaction conditions were as follows: 5 min at 95 °C, then 35 cycles of 30 s at 94 °C, 30 s at 66 °C, and 1 min at 72 °C, and a final 7-min extension step at 72 °C. GAPDH was undergoing 30 cycles at 58 °C. PCR products were subjected to electrophoresis in 1.5 % agarose gel stained with Gelred.

Immunofluorescence

For cytospin preparation, 2 × 105 cells were cytocentrifuged onto glass slides and fixed in 4 % paraformaldehyde solution in PBS for 30 min, permeabilized with 0.5 % Triton X-100/PBS for 20 min, and blocked for nonspecific binding with goat serum. Cells were then incubated with PU.1 antibody (1:1000 dilution; Cell signaling) at 4 °C overnight, then treated with FITC-conjugated goat-anti-rabbit fluorescent secondary antibody (1:100 dilution) at 37 °C for 1.5 h. The slides were washed in PBS for 5 min. Slides were embedded in antifade, cover slipped, and analyzed with a conventional fluorescence microscope (Olympus).

Western blot

Cell pellets were lysed in RIPA buffer containing 50 mM Tris pH 8.0, 150 mM NaCl, 0.1 % SDS, 0.5 % deoxycholate, 1 % NP-40, 1 mM DTT, 1 mM NaF, 1 mM sodium vanadate, and protease inhibitors cocktail (Sigma, St. Louis, MO, USA). Protein extracts were quantitated with the Bradford method, and loaded on an 8–12 % SDS-PAGE gel, electrophoresed, then transferred to a nitrocellulose membrane (Millipore, Bedford, MA, USA). The membrane was incubated with primary antibody, washed, and incubated with anti-rabbit or anti-mouse HRP-conjugated secondary antibody. The following antibodies were used: anti-PU.1 antibody (Cell signaling 1:500) and anti-Actin antibody (Beyotime 1:1000). Detection was performed using a chemiluminescent Western detection kit (Cell Signaling).

Statistical analysis

Statistical analysis was performed using SAS software. Wilcoxon rank sum test was carried out to compare the difference of frequencies between groups of patients. For all analyses, the P values were two-tailed, and a P < 0.05 was considered statistically significant.

Results

No methylation in promoter region of PU.1 in normal individuals

PU.1 was reported to be down-regulated through methylation of promoter and intron 1 in multiple myeloma patients and cell lines [15]. It has reported that PU.1 expression was down-regulated, and the activity of the BCR-ABL oncoprotein is not directly responsible for the reduced level of PU.1 mRNA seen in CML patients [8]. Hence, its underlying mechanism is still unknown. We investigated whether epigenetic inactivation of PU.1 gene could induce down-regulation of PU.1 gene expression. We designed methylation-specific PCR sequencing techniques to amplify and identify the methylation condition of 385-bp PU.1 promoter–intron 1 region containing 20 CpG islands. 385-bp PU.1 promoter–intron 1 region refers to −49 to +268 promotor nucleotides and 68 intron 1 nucleotides; this region is the hotspot of epigenetic modification (Fig. 1a). In 10 healthy donor samples, totally no methylation site was observed in these 20 CpG islands, as shown in the Fig. 1b. After bisulfite modification, all 20 C bases within and above 20 CpG islands had transited to 20 T bases.
https://static-content.springer.com/image/art%3A10.1007%2Fs12185-012-1106-x/MediaObjects/12185_2012_1106_Fig1_HTML.gif
Fig. 1

No methylation found in promoter region of PU.1 in normal individuals. a 385-bp PU.1 promoter–intron 1 region containing the 20 CpG islands. 385-bp PU.1 promoter–intron 1 region refer to −49 to +268 promotor nucleotides and 68 intron 1 nucleotides. b In 10 normal individuals, after methylation specific PCR, PCR product was cloned into pGEM-T-EASY vector, clones were sequenced to evaluate methylation status. Totally no methylation site was observed in these 20 CpG islands, sequencing result of a typical clone is shown. After bisulfite modification, all 20 C bases within 20 CpG islands have transited to 20 T bases

Aberrant methylation status in promoter region of PU.1 in chronic myeloid leukemia and K562 cell line

We further investigated promoter region of PU.1 in CML patient samples and K562 CML blast crisis cell line, using same methylation-specific PCR sequencing techniques, in order to evaluate the methylation status, respectively. To our great surprise, we have discovered different numbers of methylated CpG sites in CML-CP at diagnosis (14 samples), CML-CP with TKI (11 samples), CML-BC (10 samples) and in K562 cells. The number of methylated CpG sites range from 4/20 to 17/20. Here, we showed one typical clone of CML-CP at diagnosis, CML-CP with TKI, CML-BC and K562 cells, respectively. As shown in Fig. 2a, in CML-CP at diagnosis, 10 methylation sites were observed in these 20 CpG islands (site 6, 7, 8, 9, 14, 15, 16, 18, 19, 20). While in CML-BC clone, 17 methylation sites were observed in these 20 CpG islands (Fig. 2b, site 2, 3, 4, 5, 7, 8, 9, 10, 11, 12, 13, 14, 16, 17, 18, 19, 20). In K562 cell clone, 7 methylation sites were observed in these 20 CpG islands (Fig. 2c, site 5, 9, 13, 14, 18, 19, 20). Aberrant methylation status of PU.1 even existed in CML-CP with TKI (Fig. 2d). We further comprehensively showed methylation status of PU.1 in healthy donors, CML-CP at diagnosis, CML-CP with TKI, CML-BC samples (Fig. 3). Finally, we compared methylation status of PU.1 in CML-CP at diagnosis, CML-CP with TKI, CML-BC samples. Several CpG sites (site 3, 13) were highly methylated in CML-BC than CP at diagnosis; for CML-CP with TKI, there was statistical difference (see Table 1).
https://static-content.springer.com/image/art%3A10.1007%2Fs12185-012-1106-x/MediaObjects/12185_2012_1106_Fig2_HTML.gif
Fig. 2

Aberrant methylation status in promoter region of PU.1 in chronic myeloid leukemia and K562 cell line. Methylation-specific PCR sequencing techniques to amplify and identify the methylation condition of 385-bp PU.1 promoter–intron 1 region containing the 20 CpG islands in CML patient samples and K562 cell line. Sequencing results were analysed, and all the 20 CpG areas were truncated and combined in the figures. Methylated sites are marked with red circles, and unmethylated sites are marked with black circles. a In one of the typical CML-CP at diagnosis clones, 10 methylation sites were observed in these 20 CpG islands (site 6, 7, 8, 9, 14, 15, 16, 18, 19, 20). b In one of the typical CML-BC clones, 17 methylation sites were observed in these 20 CpG islands (site 2, 3, 4, 5, 7, 8, 9, 10, 11, 12, 13, 14, 16, 17, 18, 19, 20). c In one of the typical K562 cell clones, 7 methylation sites were observed in these 20 CpG islands (c, site 5, 9, 13, 14, 18, 19, 20). d In one of the typical CML-CP with TKI clones, Aberrant methylation status of PU.1 even detected

https://static-content.springer.com/image/art%3A10.1007%2Fs12185-012-1106-x/MediaObjects/12185_2012_1106_Fig3_HTML.gif
Fig. 3

Bisulfite sequencing of the PU.1 CpG island. The nucleotide range spans from −49 to +268 promotor nucleotides and 68 intron 1 nucleotides. Vertical lines are used to indicate the twenty individual CpG islands. Numbers at the bottom of the figure indicate the position of the CpG dinucleotides. Ten colonies are selected in normal individuals; and twenty colonies are selected in CML-CP, CML-BC and Imatinib-treated patients who acquired complete cytogenetics response, respectively. Each circle indicates a CpG site; unmethylated CpG site (open circles), methylated CpG site (closed circles)

Table 1

Statistical analysis of PU.1 methylation status in different stages

CpG site

1

2

3

4

5

6

7

8

9

10

Methylation percentage (CP)

0

3.70

3.70

96.30

96.30

11.10

11.10

11.10

100

96.30

Methylation percentage (BP)

0

17.80

28.60

64.30

57.10

25

21

28.50

78.60

60.70

Methylation percentage (TKI)

0

0

4.70

80.90

80.90

19

14.20

14.20

95

90.50

P value

0.9632

0.094

0.0124

0.0027

0.0005

0.1864

0.3079

0.1078

0.0114

0.0012

CpG site

11

12

13

14

15

16

17

18

19

20

Methylation percentage (CP)

96.30

7.40

7.40

11.10

11.10

11.10

7.40

11.10

7.40

11.10

Methylation percentage (BP)

60.70

25

39.30

32.10

10.70

21.40

21.40

14.30

17.80

21.40

Methylation percentage (TKI)

90.50

9.50

9.50

14.20

14.20

14.20

9.50

14.20

14.20

14.20

P value

0.0012

0.0791

0.0048

0.0596

0.9632

0.31

0.1434

0.7299

0.2531

0.31

Sequence clone number: CML-CP: 27; CML-BC: 30; CML-CP with TKI:21

PU.1 mRNA expression levels were lowered in CML patient samples and K562 cells

It has been demonstrated that expression of PU.1 was severely impaired in patients with CML at diagnosis [8]. We have discovered aberrant methylation status in promoter region of PU.1 in CML patient samples and K562 cells. To know whether this kind of aberrant epigenetic modification is associated with the impairment of PU.1 expression, we checked the mRNA expression levels in these patient samples and K562 cells which harbored aberrant methylation. Semiquantitative PCR was performed in order to assess the level of PU.1 mRNA expression (Fig. 4). We confirmed that the mRNA expression level of PU.1 in CML patient samples and K562 cells were really lower than their normal counterparts, but there was no obvious difference within CML-BC and CP.
https://static-content.springer.com/image/art%3A10.1007%2Fs12185-012-1106-x/MediaObjects/12185_2012_1106_Fig4_HTML.gif
Fig. 4

PU.1 mRNA expression levels were lowered in CML patient samples and K562 cells. Semiquantitative PCR was done to assess the level of PU.1 mRNA expression in CML patient samples and K562 cells which harbored aberrant methylation, GAPDH as internal control. From left to right: lane 1 DL2000marker, lanes2, 3, 4, 5 GAPDH for normal control individuals (CML-CP, CML-BC, and K562, respectively). lanes6, 7, 8, 9 PU.1 for normal control individuals (CML-CP, CML-BC, and K562, respectively

PU.1 protein expression levels were weakened in CML patient samples and K562 cells

We also examined the expression level of PU.1 protein approximately via indirect immunofluorescence method as shown in Fig. 5. We use the rabbit anti-PU.1 monoclonal antibody as the first antibody (1:1000 dilution), FITC-conjugated goat-anti-rabbit secondary antibody (1:100 dilution) to detect the distribution and intensity of the fluorescence. Cells from BMNC of a healthy donor, a CML-CP patient and K562 cells were cytospinned onto slides and indirect immunofluorescence experiment was performed. As shown in Fig. 5, fluorescence immensely distributed in the cytoplasmic and perinuclear area, with high intensity in normal individuals (Fig. 5a 200×, d 1000×). While, fluorescence sparsely distributed in the cytoplasmic and perinuclear area, with lower intensity both in CML-CP cells and K562 cells at different degrees of magnification (Fig. 5b 200×, e 1000× for CML-CP, c 200×, f 1000× for K562). Thus, when compared with normal individuals, the fluorescence distribution scope was much less, and fluorescence intensity was obviously weaker in both CML-CP (Fig. 5b, e) cells and K562 cells (Fig. 5c, f). Further, we detected PU.1 protein expression level in CML-CP patients and K562 cells via Western blot technique, again, we found that the PU.1 protein expression level severely decreased both in CML-CP patient cells and K562 cells (Fig. 6).
https://static-content.springer.com/image/art%3A10.1007%2Fs12185-012-1106-x/MediaObjects/12185_2012_1106_Fig5_HTML.jpg
Fig. 5

PU.1 protein expression levels were weakened in CML patient samples and K562 cells. Indirect immunofluorescence method was used to examine the expression level of PU.1 protein in normal control individuals, CML-CP, K562 cells. Rabbit anti-PU.1 primary antibody (1:1000 dilution); FITC-conjugated goat-anti-rabbit secondary antibody (1:100 dilution). Cells from BMNC of a heath donor, a CML-CP patient and K562 cells were cytospinned onto a slide and indirect immunofluorescence experiment was performed and observed under the fluorescence microscope at different magnify degrees (ac ×200; df ×1000). Compared with normal individuals (a, d), the fluorescence distribution scope was much less, and fluorescence intensity was obviously weaker in both CML-CP (b, e) sample and K562 cells (c, f)

https://static-content.springer.com/image/art%3A10.1007%2Fs12185-012-1106-x/MediaObjects/12185_2012_1106_Fig6_HTML.gif
Fig. 6

PU.1 protein expression levels were weakened in CML patient samples and K562 cells. PU.1 protein expression levels were detected in CML-CP patient and K562 cells via Western blot technique, PU.1 protein expression level severely decreased both in CML-CP patient cells and K562 cells, actin as internal control. From left to rightlane 1 normal individuals, lane 2 CML-CP, lane 3 K562 cells

Discussion

Our present results displayed the following: (a) we confirmed the reduced expression of PU.1 both in CML patients and CML blast cell line K562 at molecular and protein levels; (b) we specifically unveiled its underlying mechanism: we found aberrant methylation in the promoter region of transcription factor PU.1 in CML patients both in chronic phase and blast crisis phase, as well as CML blast K562 cell line, while totally no methylation was observed in normal individual counterparts. This may be responsible for the down-expression of PU.1. Thus, aberrant methylation of PU.1 may play a role in CML pathogenesis; (c) CML patient acquired complete cytogenetic remission under imatinib mesylate treatment; aberrant methylation status of PU.1 was not reversed, demethylating drugs might be considered in the future.

The PU.1 transcription factor is a master regulator of hematopoiesis. PU.1 regulates the expression of G-CSF, M-CSF, and GM-CSF receptors [16] and interacts with other factors such as GATA-1 [17] and IL-7Rα [18] to manipulate the expression of genes. Loss of PU.1 function has been found to be related to human and experimental AML [19]; including loss-of-function gene mutation or reduced expression. Inactivating mutations in the PU.1 gene have been found in radiation-induced murine myeloid leukemias and human AML samples [20, 21] and PU.1-deficient adult mice developed myeloid leukemia [22]. Several leukemogenic proteins such as AML1-ETO and PML-RARa have been described to interfere with the PU.1 function or expression [23, 24]. Taken together these data indicates that PU.1 has tumor suppressor activity in myeloid cells.

Considering the significance of PU.1 in the procedure of hematopoiesis, we studied whether PU.1 is involved in the initiation and progression of CML or not. In our previous work, we have screened the promotor region and coding sequence of PU.1 in CML patients and found no existing mutations [7]. During the same period, another research group has claimed that expression of PU.1 is severely impaired in patients with chronic myeloid leukemia (CML), they also transfected BCR-ABL in murine myeloid 32D cells (which do not express endogenous BCR-ABL), no changes in PU.1 expression were observed in two 32D sublines with enforced expression of BCR-ABL, furthermore, siRNAs directed against BCR-ABL present in K562 cells resulted in the down-regulation of BCR-ABL, but increase in PU.1 expression was not observed, suggesting that the activity of the BCR-ABL oncoprotein is not directly responsible for the reduced level of PU.1 mRNA seen in CML patients [8]. So, up to now, its underlying mechanism is unknown. To shed light on the mechanism, we hypothesized whether epigenetic modification is responsible for it.

Hypermethylation of promoter region of gene leading to abnormal silencing of transcription is common in cancers [9]. It has been proved that 385-bp PU.1 promoter-intron 1 region containing 20 CpG islands is a methylation hotspot, and in multiple myeloma patients aberrant methylation existed in this area, which caused down-expression of PU.1 [15]. So, in our present study, we analyzed the methylation status of the PU.1 promoter–intron 1 region containing 20 CpG islands using bisulfite sequencing to calculate the number of methylated CpG sites. Different numbers of methylated CpG sites were discovered in CML-CP at diagnosis, CML-CP with TKI, CML-BC and CML leukemic cell line K562. The number of methylated CpG sites range from 4/20 to 17/20. Surprisingly, no methylation site was observed in normal individuals. Interestingly, we also found aberrant methylation status of PU.1 even in those CML patients who acquired complete cytogenetic remission via TKI treatment, suggesting that TKI agent can target BCR/ABL oncoprotein and inhibit cell proliferation, but cannot abrogate methylation modification. This phenomenon might be associated with a small portion of CML disease progression or relapse under TKI treatment. In addition, we also found no methylation in few K562 cell clones, but it was quite rare. This might belong to cell variation due to continuous cell culture and passage. At last, we compared methylation status of PU.1 in CML-CP at diagnosis, CML-CP with TKI, CML-BC samples. We found that several CpG sites were highly methylated in CML-BC than CP at diagnosis and CML-CP with TKI, which has obvious statistical difference (see Table 1). We then investigated the expression level of PU.1 mRNA and protein with the purpose to testify whether methylation aberration in promoter region of PU.1 could lead to down-regulation of PU.1 expression. From the results, we again confirmed that PU.1 was really down-regulated at CP and BC of CML, as well as in K562 cell line which harbored aberrant methylation. We found that two CpG islands were highly methylated in CML-BC than CP. Nevertheless, corresponding PU.1 mRNA or protein expression levels have no obvious difference. Taken together, we conclude from these data that aberrant methylation in promoter region of PU.1 may play a role in CML leukemogenesis, but not disease progression from CML-CP to BC.

In conclusion, we demonstrated that aberrant methylation in the CpG island of the promoter region of PU.1 gene is a common event at different stages of CML, even in those patients acquired complete cytogenetic remission via TKI treatment. It seemed that this kind of abnormal methylation status might be associated with CML pathogenesis. Further study will be needed to determine the definite role of PU.1 methylation in the development, and prognosis of CML. Epigenetic modification of the target agent might be needed in combination treatment of CML. Hopefully, it could be served as a useful biomarker and potential target for demethylating drugs. Demethylating agents in combination with TKI drugs might become a reasonable therapeutic regimen for patients with CML in future.

Acknowledgments

We thank all members of the Department of Hematology, Xin Hua Hospital, Affiliated to Shanghai Jiao Tong University (SJTU) School of Medicine, for their support. This work was supported in part by the Shanghai Jiao Tong University (SJTU), School of Medicine natural science Grant (09XJ077).

Conflict of interest

The authors have declared that no conflict of interest exists.

Copyright information

© The Japanese Society of Hematology 2012