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1 Department of Gene Regulation, and
2 Department of Biochemistry, Faculty of Pharmaceutical Sciences, Tokyo University of Science, Noda, Chiba 278-8510, Japan
3 Genome and Drug Research Center, Tokyo University of Science, Noda, Chiba 278-0022, Japan
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Abstract |
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 Top Abstract IntroductionResultsDiscussionExperimental proceduresReferences |
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Introduction |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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Metabolism of poly(ADP-ribose), which is a post-translational modification in eukaryotic cells catalyzed primarily by poly(ADP-ribose) polymerase (PARP) and poly(ADP-ribose) glycohydrolases (PARG), has been suggested to play important roles in regulating cell functions (Bonicalzi et al. 2005; Koh et al. 2005; Schreiber et al. 2006). Reversible poly(ADP-ribosyl)ation of chromosomal proteins has been suggested to be important in the regulation of nuclear functions, including DNA replication (Tanuma & Kanai 1982; Lonn & Lonn 1985), and repair (Durkacz et al. 1980; Kreimeyer et al. 1984; Tanuma et al. 1985), spindle assembly (Chang et al. 2004) and transcription (Kraus & Lis 2003). Moreover, it has been suggested that poly(ADP-ribose) metabolism plays important roles in carcinogenesis and in autoimmune diseases (Masutani et al. 2003, 2005; Oei et al. 2005). Recently, analysis of a loss-of-function mutant of the PARG gene in Drosophila revealed that PARG plays an important role in neurogeneration through degradation of poly(ADP-ribose) in the brain (Hanai et al. 2004). Studies of knockout mice showed that PARG is involved in DNA damage responses in pathological process (Cortes et al. 2004), and in embryonic development (Koh et al. 2004).
Previously, we observed that both expression and activity of PARG is transiently suppressed and gradually returns to initial levels during monocyte/macrophage-like differentiation of HL-60 cells treated with TPA (Uchiumi et al. 2004). Because poly(ADP-ribosyl)ation is known to modulate chromatin structure and transcription (Tulin & Spradling 2003; Kim et al. 2004), it is possible that poly(ADP-ribose) metabolism is involved in regulation of cell differentiation. At present, PARG is known as one of the primary enzyme that degrades poly(ADP-ribose) (Miwa et al. 1974; Tanuma & Endo 1990), therefore, transcription factors that affect PARG gene expression may be the key proteins that provoke HL-60 cells to undergo macrophage-like differentiation.
In the present study, we isolated a 2.1-kbp 5'-flanking region of the human PARG gene by polymerase chain reaction (PCR) with HL-60 genomic DNA as a template. Sequence analysis and primer extension experiments showed that the promoter has no TATA box but does possess several transcription start sites. Moreover, deletion analyses revealed that a 75-nt region in this 2.1-kbp fragment is required for basal promoter activity and positive response to TPA. TFSEARCH analysis indicated that multiple Ets family protein-binding motifs are located in the 75-nt region. The Ets family consists of 30 genes homologous to the proto-oncogene Ets-1 (Oikawa 2004). Electrophoretic mobility shift analysis (EMSA) and chromatin immunoprecipitation (ChIP) experiments indicated that PU.1 interacts with the 75-nt fragment. PU.1, which is also known as Spi-1 (Ray et al. 1990), regulates myeloid lineage commitment (Nerlov & Graf 1998), and it was reported that AML is induced by graded reduction of PU.1 (Rosenbauer et al. 2004). Moreover, transient transfection experiments indicated that the PU.1-binding motif is essential for the PARG promoter activity and responsiveness to TPA. Interestingly, co-transfection of a PU.1 expression vector with the PARG promoter–luciferase reporter constructs showed that the PARG promoter activity decreases by 24 h after TPA treatment. Western blot analysis indicated that expression of PU.1 transiently increases then gradually decreases after TPA treatment of HL-60 cells, suggesting that PU.1 acts as a negative regulator of PARG gene expression. Taken together, these findings suggest that PU.1-binding motifs are essential for PARG promoter activity and that PU.1 modulates PARG gene expression during differentiation of HL-60 cells.
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Results |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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A 2.1-kb fragment of the PARG gene from HL-60 DNA (Fig. 1A) was cloned into the pGL3-basic vector to generate pPARG-Luc#2. The nucleotide sequence of the 2.1-kb region is included in the sequences of NT 008583.16 (nts from 172656 to 174771), NT 035036.2 (nts from 185841 to 183729), and NW 001837980.1(nts from 1614 to 3728) and contains the 5' end of the cDNA sequence (NM 003631.1) of the PARG gene. The 2.1-kb fragment contains 0.43-kb of the bidirectional promoter region of TIM23 and PARG (XM 928114.3/XM 001715214.1/NM 006327.2) (Meyer et al. 2003).
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TFSEARCH was used to screen the 2.1-kb region for characteristic recognition sequences of known transcription factors, and several were found (Fig. 1A). Interestingly, GATA-1 (–279 to –270), AP-4 (–158 to –149), Elk-1 (–115 to –102), STAT-X (–98 to –90), c-Ets–Elk-1 (–83 to –70, –59 to –46, –51 to –38), CREB (–37 to –28) and Myo D (+25 to +34) sequences were located within an only 0.3-kb region (Fig. 1A,B). No sequence similar to a TATA or CCAAT box was found in the 2.1-kb region. Just upstream of the minor transcription start sites, three c-Ets–Elk-1 (Treisman et al. 1992; Woods et al. 1992) and CREB (Benbrook & Jones 1994) motifs are located in a 75-bp region (Fig. 1B). The region may form a cruciform structure (Fig. 1D).
Identification of promoter activity of the PARG gene in HL-60 and Jurkat cells
To examine promoter activity of the 2.1-kb 5'-flanking sequence of the PARG gene, pPARG-Luc#2 was transfected into HL-60 (Fig. 2A,B) and Jurkat (Fig. 2D,E) cells. The Luc activity of pPARG-Luc#2 transfected cells was equal to or half of that of pGL3-promoter vector-transfected cells (Fig. 2). We also transfected pPARG-Luc#2 into ML1 and HeLa cells and observed the Luc activity (data not shown). The results strongly suggested that the isolated 2.1-kb fragment contained a functional PARG promoter.
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-ray irradiated (Fig. 2A,D). Although, pPARG-Luc#2-transfected cells showed slight induction of Luc activity 2 h after irradiation, similar responses were observed in pGL3-promoter-vector-transfected cells. Therefore, induction of Luc activity after the irradiation may be because of specific sequences within the pGL3 vector.
Previously, we analyzed changes in PARG gene expression in HL-60 cells treated with TPA (Uchiumi et al. 2004) and found that the level of PARG transcript decreased to 20% that in untreated cells after 4 h of TPA treatment and then returned to the initial level. As shown in Fig. 2B, PARG promoter activity decreased to 80% of that in untreated cells then increased more than twofold after 24 h of TPA treatment. The pattern of induction of the PARG promoter by TPA treatment is absolutely different from that of the HIV promoter, in which NF-
B elements are located (Fig. 2C). Luc activity of the pGL3-promoter vector-transfected cells unchanged for 4 h and then by 24 h decreased to 50–70% of that in untreated cells. Luc activities of pGL3-promoter-, pPARG-Luc#2-, and pGL3-HIVLuc-transfected Jurkat cells increased in a similar manner after TPA treatment (Fig. 2E,F), suggesting that the response to TPA is not specific to the PARG promoter in T cells.
Deletion analysis of the promoter region of the human PARG gene
To identify DNA sequences that are essential for PARG promoter activity and the response to TPA, various deletion constructs were introduced into HL-60 and Jurkat cells, which were then treated with or without TPA (Fig. 3A). The promoter activity and TPA response were prominent in both cell lines with a deletion to position –152. Thus, the sequence between –152 and +170, which includes c-Ets–Elk-1 motifs and transcription start sites, is primarily responsible for PARG promoter activity and the TPA. Because the construct with the –565 to –153 deletion showed increased promoter activity, this 0.4-kb region may contain negative regulatory elements.
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1Luc) and disappeared completely when –102 to –58 (compare pKBST1Luc with pKBST2Luc and pKBST5Luc with pKBST7Luc), suggesting that positions –152 to –57 carries at least one positive regulatory element. In contrast, deletion of +78 to +170 of pKBST1Luc and deletion of –27 to +77 of pKBST5Luc increased basal promoter activity, indicating that the DNA sequence from –27 to +170 includes an inhibitory sequence(s). These results suggest that the 75-nt sequence from –102 to –28 (Fig. 1B,D) is the minimum promoter that affects TPA responsiveness as well. Binding of nuclear factors to the c-Ets–Elk-1 elements of the PARG promoter
We examined whether any nuclear factors bind to the sequence between –102 and –28, which contains c-Ets–Elk-1 binding motifs. Nuclear extracts prepared from TPA-treated HL-60 cells were incubated with a 75-bp double-stranded DNA (probe D, Fig. 5E) end labeled with 32P, and EMSA was carried out. As shown in Fig. 4A, mobility-shifted bands C1–C4 were observed. Quantification of each band signal revealed that formation of DNA–protein complexes with C1, C2 and C4 after TPA addition was essentially equal, peaked at 4–6 h after treatment, and then declined gradually over the next 24 h (Fig. 4B). The amount of complex C3 did not change after TPA treatment, suggesting that the protein(s) contained in the C3 complex differs from those contained in C1, C2 and C4.
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Sequence-specific binding of PU.1 to the 75-nt region
The above data indicated that multiple DNA–protein complexes were formed when the 32P-labeled 75-bp (probe D) was incubated with HL-60 nuclear extracts. To investigate if these DNA–protein complexes result from DNA sequence-specific binding of proteins, EMSA competition analysis was performed (Fig. 5A) with various double-stranded DNA probes as competitors (Fig. 5E). Formation of complexes C1, C2 and C3was reduced in a dose-dependent manner by the addition of DNA probes A, B, C and D (Fig. 5A,B). In contrast, DNA probes 1, 2 and 3 did not compete with the 32P-labeled 75-bp sequence for formation of C1, C2 and C3. Interestingly, formation of C4 was inhibited by competitor probes A, B, D, 2 and 3 (Fig. 5A). The binding affinities were estimated to be A = B = D = 2 > 3 > 1 > C from the competition profile (Fig. 5B, C4).
Because anti-PU.1 antibody reduced C4 (Fig. 4C) formation, we speculated that PU.1 is involved in the C4 complex. Therefore, we used a GST–PU.1 fusion protein purified by glutathione-Sepharose column chromatography for EMSA. A DNA–protein complex was formed when GST–PU.1 was incubated with the 32P-labeled probe D (Fig. 5C, lanes 1, 14, 15 and 28). DNA–protein complex formation showed greater competition in the presence of unlabeled probes A, B and D than probes 1, 2, 3 and C. Although the order of binding affinity of GST–PU.1 was not completely identical to that with HL-60 nuclear extracts (Fig. 5B), these results suggest that the C4 complex contains PU.1.
PU.1 associates with the PARG promoter in HL-60 and ML1 cells
To examine whether PU.1 binds to 75-nt of the human PARG promoter, ChIP analysis was performed (Fig. 6). Anti-PU.1-immunoprecipitated chromatin fractions of HL-60 and ML1 cells contained the 75-nt region (lanes 2 and 7), whereas IgG-precipitated chromatin fractions did not (lanes 3 and 8). These results suggest that PU.1 associates with the 75-nt sequence of the PARG promoter in HL-60 and ML1 cells. In this experiment, chromosomal DNAs were treated by sonication. However, slight amount of unshared large DNAs might be remained in the input and unbound fractions. The large DNAs might hold PCR product to give rise to 2-kbp background bands (lanes 1, 4, 5, 6, 9 and 10).
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EMSA and ChIP analysis strongly suggested that PU.1 binds to the 75-nt region of the PARG promoter. As shown in Fig. 3, the activity of the human PARG promoter increased 24 h after TPA treatment. To examine whether the c-Ets–Elk-1 motifs in the 75-nt element respond to TPA or PU.1, various Luc-expression constructs and PU.1 expression vector were co-transfected into HL-60 cells, and the cells were treated with TPA (Fig. 7A). The Luc activities of pKBST-Luc- and pKBST6Luc-transfected cells were enhanced by TPA (Fig. 7A, columns 5–8 and 9–12). However, co-transfection with pcDNAPU.1 reduced the positive effect of TPA (compare columns 6 with 8 and 10 with 12), suggesting that PU.1 acts as a negative regulator of TPA-induced PARG promoter activity. In contrast, the PU.1 expression vector had no obvious effect on the SV40 promoter (Fig. 7A, columns 1–4). These results suggest that PU.1 suppresses TPA-induced PARG promoter activity.
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6Luc, pGL3-75WT and 75mt0 with pcDNA3.1B showed almost the same level of Luc activities. The promoter activity decreased with the increase of mutations in the c-Ets–Elk1 motifs (Fig. 7B and open columns in Fig. 7C). However, the Luc activities from these reporter constructs-transfected cells were not changed in the presence of PU.1 expression vector 24 h after transfection (Fig. 7C). Luc activities of pKBST6Luc-, pGL3–75WT-, 75mt0- and 75mt1-transfected cells were activated by the TPA treatment as that of pKBST- or pPARG-Luc#2-transfected cells (Figs 2 and 7A,D). The data also show that co-transfection of pcDNAPU.1 reduced Luc activities of TPA-treated cells (Fig. 7D, compare bars 2, 6 and 10 with 4, 8 and 12, respectively). Although the relative Luc activity of cells transfected with pGL3–75mt1 was lower than that of the pGL3–75WT- or pGL3–75mt0-transfected cells and did not respond to PU.1, the 75mt1 sequence still responded to TPA positively. Therefore, the down-stream c-Ets–Elk-1 motifs in the 75-nt are essential for TPA responsiveness. In contrast, the upstream c-Ets–Elk-1 motif responded negatively to PU.1 (Fig. 7D, bars 25–28) more apparent than downstream motifs in pGL3–75mt1 (Fig. 7D, bars 13–16). Moreover, these c-Ets–Elk-1 motifs together play a role in a basal promoter function. Because promoter activity was absent completely with the pGL3–75mt12 construct (Fig. 7D, bars 29–32). To test whether these mutations affect PU.1 binding, EMSA competition was carried out with GST–PU.1 and 32P-labeled 75WT probe (Fig. 8). As summarized in Fig. 8B, unlabeled 75mt0 and 75mt1 probes competed for the 32P-labeled 75WT/GST–PU.1 complex formation. In contrast, competition ability was reduced by 75mt2 and 75mt12 mutations. These results are consistent with the results of transient transfection assays, suggesting that PU.1 can bind to the downstream c-Ets–Elk-1 motifs which are responsible for TPA response, although it cannot enhance the TPA responsiveness (Fig. 7D). However, the 75mt2 probe provided greater competition than the 75mt12 probe, suggesting that the promoter suppressing effect of PU.1 results from its interaction with the upstream c-Ets–Elk-1 motifs (Fig. 7D, bars 25–28).
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The above data suggest that PU.1 is a negative regulator of PARG expression by binding to the 75-nt region. As shown in Fig. 2B, PARG promoter activity in HL-60 cells was reduced 4 h after TPA treatment, and then increased further after incubation for 20 h. We therefore examined the expression of PU.1 protein during HL-60 differentiation induced by TPA (Fig. 9). The PU.1 signal detected by Western blotting peaked 2–6 h after TPA addition and then declined. This expression profile of PU.1 was the opposite of that of PARG promoter activity (Fig. 2B). Expression of PARP protein in response to TPA was observed within 1 h but decreased after 24 h significantly (Fig. 9A). The poly(ADP-ribosyl)ated proteins accumulated and peaked 6 to 8 h after addition of TPA (Fig. 9B, lower panel) and then decreased gradually. The profile is consistent with the changes in promoter activity, gene expression and enzyme activity of PARG (Uchiumi et al. 2004). Thus, changes in PARP and PARG activities may control expression of various genes during TPA-induced differentiation. The balance between PARP and PARG may influence poly (ADP-ribosyl)ation of various proteins during TPA-induced HL-60 differentiation.
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Discussion |
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PU.1 is a member of the Ets family of proteins that mediate binding to the purine-rich 5'-GGA(A/T)-3' element (Wasylyk et al. 1993) and is known as a hematopoietic transcription factor that controls the expression of B cell- and macrophage-specific genes (Moreau-Gachelin 1994; Oikawa 2004). PU.1-binding sequences are found in promoter regions of proteins, including macrophage colony-stimulating factor (M-CSF) receptor (Zhang et al. 1994), and granulocyte-macrophage colony-stimulating factor (GM-CSF) receptor 
(Hohaus et al. 1995). The proteins encoded by these genes, which are positively regulated by PU.1, play important roles in lymphoid/myeloid cells. It is noteworthy that most of these lymphoid/myeloid-specific promoter regions are TATA-box less similar to the PARG promoter. PU.1 binds specifically to TATA-binding protein in vitro (Hagemeier et al. 1993). Therefore, PU.1 binding to TATA-less promoters (including the PARG promoter) may tether general transcription machinery. Studies are needed to confirm this. In contrast, PU.1 could work as a negative transcription regulator (Lloberas et al. 1997; Harendza et al. 2000; Suzuki et al. 2003). Recent studies showed that PU.1 suppresses promoter activity of the µ opioid receptor gene (Hwang et al. 2004). Moreover, PU.1 modulates TAL-1 gene expression by binding to its silencer (Le Clech et al. 2006). Thus, PU.1 can induce or suppress expression of specific genes. In our experimental system, in which macrophage-like differentiation of HL-60 cells was induced by TPA treatment, PU.1 reduced the PARG promoter activity. This is suggested because PARG promoter activity decreased (Fig. 2B) as PU.1 protein expression (Fig. 9A,B) and C4 formation (Fig. 4A,B) increased and vice verse.
In the present study, TPA responsive positive element of the PARG promoter was not indicated directly. The positive response to TPA disappeared with a mutation in the two overlapping c-Ets–Elk1 motifs (pGL3-75mt2, Fig. 7D), indicating that a TPA responsive factor or factors recognize this sequence. It is possible that PARG promoter activity is regulated by PU.1 in a precise manner. Low PU.1 expression levels may stimulate promoter activity, whereas high levels suppress it. Another protein factor that is involved in C3 complex (Fig. 4) is also a candidate positive regulatory factor, because the ratio of the DNA–protein complexes C3/(C1 + C2 + C4) (Fig. 4A,B) corresponds to the PARG promoter activity, yet this must be verified by further experiments. Several transcription factors, including Jun D, Fos B, NKX2.1, TCF3, KLF4 and Rab2, have been identified as early response genes (Zheng et al. 2002). They may induce gene expression of an activator(s) of the PARG promoter, which is activated at 24 h after TPA treatment (Fig. 2B). Moreover, PU.1 is known to interact with other transcription factors such as c-Jun, GATA-1, C/EBPβ and Maf B (Friedman 2007). Among these, GATA-1 is unique in that the interaction with PU.1 inhibits both proteins’ functions (Rekhtman et al. 1999; Zhang et al. 2000). Because a putative GATA-1-binding site is located upstream of the PU.1-binding motifs in the PARG promoter (Fig. 1A), GATA-1 might regulate the PARG promoter through an interaction with PU.1. CREB motif is also located in the 75-nt region (Fig. 1A). Therefore, CBP binding to the motif and its interaction with PU.1 protein may also affect PARG promoter activity.
Very recently, it was reported that poly(ADP-ribose) polymer induces cell death (Andrabi et al. 2006). Moreover, XRCC1 was suggested to regulate poly(ADP-ribose)-mediated apoptosis (Keil et al. 2006). In contrast, delayed poly(ADP-ribose) degradation in PARG-silenced cells protects against H2O2-induced cell death (Blenn et al. 2006). These observations suggest that poly(ADP-ribosyl)ation must be controlled precisely. Namely, cells require an appropriate level of poly(ADP-ribose) to survive. As shown in Fig. 9, the amount of poly(ADP-ribosyl)ated proteins was greatly increased by TPA within 2 h and decreased gradually over the next 48 h. Rapid induction of poly(ADP-ribosyl)ation after TPA treatment is explained by the activation of PARP-1 (Fig. 9A), as well as the decrease in PARG activity (Uchiumi et al. 2004). Given that accumulation of poly(ADP-ribosyl)ated proteins drive a death signal, induction of PARG gene expression and PARG activity may be required for cell survival. In the present study, PU.1 reduced PARG promoter activity. From the study of PU.1 knockdown, following reduction of c-Jun and Jun B was suggested to lead AML (Steidl et al. 2006). If PU.1 knockdown also induces PARG gene expression, induction of poly(ADP-ribose) degradation might protect cells from death. Furthermore, over-expression of PU.1 was shown to induce cell growth inhibition and apoptosis (Yamada et al. 1997). These observations suggest that PU.1 expression may activates poly(ADP-ribosyl)ation via suppression of PARG gene expression.
DNA microarray analysis showed that gene expression profiles are classified into four types, early, intermediate, late and biphasic, in TPA-induced HL-60 cell differentiation (Zheng et al. 2002). Our present data suggest that the proto-oncogene product PU.1 modulates PARG gene expression during HL-60 differentiation. Previous studies indicated that poly(ADP-ribosyl)ation influences chromatin structure and transcription (Kraus & Lis 2003; Schreiber et al. 2006). Nuclear events, including poly(ADP-ribosyl)ation of core histones and chromatin-associated proteins and PARP-1 binding to nucleosomes, have been suggested to alter chromatin structure to facilitate decondensation and transcription by pol II (Kraus & Lis 2003; Tulin & Spradling 2003; Kim et al. 2004). However, poly(ADP-ribosyl)ation of specific transcription factors has been shown to decrease DNA binding and transcriptional activation (Oei et al. 1998).
The molecular mechanism by which PU.1 influences differentiation of hematopoietic cells has been studied and explained through interactions with various transcription factors (Friedman 2007) and modulating transcription factor cascades (Laslo et al. 2006). Here, we propose that poly(ADP-ribose) metabolism is regulated at least in part by PU.1. Because the poly(ADP-ribose) levels in cells have been suggested to control cell death or survival, poly(ADP-ribose) metabolism may determine hematopoietic cell fate as well. Further studies, including analysis of the functions of PARP and PARG, may clarify physiologic roles of poly(ADP-ribosyl)ation during cell differentiation.
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Experimental procedures |
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TPA was purchased from Sigma Chemical Co. (St Louis, MO). [32P]ATP and poly(dI-dC) were from Amersham Biosciences (Piscataway, NJ). Restriction enzymes and other DNA modifying enzymes were from Takara (Kyoto, Japan) and Toyobo (Tokyo, Japan).
Cell cultures
HL-60 and ML1 cells are human promyelocytic and myeloblastic leukemia cell lines, respectively (Huberman & Callaham 1979; Rovera et al. 1979; Oka & Takeda 1997). Jurkat is a human acute lymphocytic T-cell line (Uchiumi et al. 1992). They all were cultured in RPMI 1640 medium supplemented with heat-inactivated 10% fetal calf serum (FCS), 2 mM L-glutamine, penicillin (100 IU/mL) and streptomycin (100 µg/mL). Differentiation was induced by TPA (5 ng/mL) in exponentially growing HL-60 cells at a density of 2 x 105 cells/mL.
Construction of Luc expression plasmids
Extraction of DNA from HL-60 cells and PCR were performed as described previously (Uchiumi et al. 2004). Twenty nanograms of genomic DNA were used as template for PCR amplification. To amplify 2.1-kb fragment in the 5' promoter region of the PARG gene, the following primer sequences were determined from the GenBank database (Accession No. AL450342 [GenBank] ): PS-1939 (sense), 5'-GATATCGGTACCGGCTCTAAAATGGCATC-3' and PAR + 170 (antisense), 5'-GATATCTCGAGATGGACT GCGCGTCCTTC-3'. Amplification conditions were 30 cycles of 94 °C for 30 s, 55 °C for 30 s and 68 °C for 4 min. PCR products were digested with KpnI and XhoI and then separated on 0.9% agarose gels. After electrophoresis, DNAs of the correct length were excised from the gels and subcloned into the KpnI–XhoI site of the pGL3-basic vector (Promega, Madison, WI) to generate pPARG-Luc#2. The nucleotide sequences of the PCR products were determined directly with a Thermosequenase DNA sequencing kit (Amersham Bioscience) with FITC-labeled RV and GL primers and analyzed with a DSQ 2000 L DNA sequencing system (Shimadzu Corp, Tokyo, Japan). BLAST search program showed that the cloned 2.1-kb sequence is over 99% identical to sequences, NT 008583.16, NW 01837980.1 and NT 035036.2. However, the 2.1-kb sequence contains substitutions of nucleotides, –64G to A, –47C to G, +49G to A, +53G to A, which are reported in the sequence of the NW 001840231.1. The sequences identical to the NW 001840231.1 are contained in Luc reporter plasmids shown in Fig. 3, pGL3-75-mt0, and EMSA probe mt0. After sequencing, TFSEARCH software <http://www.cbrc.jp/research/db/TFSEARCH.html> was used to identify putative transcription factor-binding motifs in the 2.1-kb region. pPARG-Luc#2 was then treated with restriction enzymes and blunt ended with T4 DNA polymerase and re-ligated. The restriction enzymes used (and resultant plasmids) were as follows:
KpnI and AvrII (pKA-Luc), KpnI and SacI (pKS-Luc), KpnI and BstEII (pKBST-Luc), KpnI and NruI (pKN-Luc), and NruI and XhoI (pNX-Luc).
Other deletion derivatives were generated by PCR with pPARG-Luc#2 as a template and various primer sets. Sense primers were PA-102, 5'-GATATCGGTACCGGTTCCCGTTAA GCGCC-3'; PA-57, 5'-GATATCGGTACCCCGGAAGCT GGAAGCGC-3'; PA + 1, 5'-GATATCGGTACCCGAAT CAAAGCGGCGCA-3'; and PA + 36, 5'-GATATCGGTACC AGTGGAAGAGAGAAAGC-3'.
Antisense primers were PAR + 170, 5'-GATATCTCGAGAT GGACTGCGCGTCCTTC-3'; PAR + 77, 5'-GATATCTC GAGTCAGGCCGTAAACACTCG-3'; PAR-28, 5'-GATAT CTCGAGCTGCCGTCAGGCGCTTCC-3'; PCR products were amplified with the PA-102/PAR + 170, PA-57/PAR + 170, PA + 1/PAR + 170, PA + 36/PAR + 170, PA-102/PAR + 77, PA-102/PAR-28 and PA-57/PAR + 77 primer pairs. PCR products were then digested with KpnI–XhoI and subcloned into pGL3-basic to generate pKBST1 to 7Luc.
Mutations were introduced into the PU.1 elements in the PARG promoter by annealing two oligonucleotides as follows:
Sense oligos were 75WTS, 5'-CGGTACCGTTAAGCGCC ACACTTCCGCTTTGCGGCAATGTGCGCC-3'; PU.WTS, 5'-CGGTACCGTTAAGCGCCACACTTCCGCTTTGCG GCAGTGTGCGCC-3'; and PU.MS1, 5'-CGGTACCGTTAA GCGCCACACTTGGGGTTTGCGGCAGTGTGCGCC-3'.
Antisense oligos were 75WTA, 5'-TCTCGAGTCAG GCGCTTCCAGCTTCCGGGGCGCACATTGCCGCAA-3'; PU.WTA, 5'-TCTCGAGTCAGGCGCTTCCGGCTTCCG GGGCGCACACTGCCGCAA-3'; and PU.MA2, 5'-TCTCG AGTCAGGCGCTTGGGGCTTGGGGGGCGCACACT GCCGCAA-3'.
The following annealed oligonucleotide pairs were used 75WTS-75WTA, PU.WTS-PU.WTA, PU.MS1-PU.WTA, PU.WTS-PU.MA2 and PU.MS1-PU.MA2. The annealed oligos were filled in with Klenow enzyme, digested with KpnI–XhoI and then cloned into pGL3-basic vector to generate pGL3-75WT, pGL3-75mt0, pGL3-75mt1, pGL3-75mt2 and pGL3-75mt12, respectively. The nucleotide sequences of the inserted DNA fragments in these Luc reporter constructs were confirmed by sequencing as described above.
Primer extension analysis
Hybridization of a primer and extension reaction was carried out as described elsewhere (Triezenberg 1987). A 29-mer nucleotide (PAR + 77) carrying the antisense nucleotide sequence from +59 to +77 was 32P-5'-end labeled with T4 polynucleotide kinase and then hybridized to total RNAs (10 µg) from HL-60 and Jurkat cells in 1 x hybridization buffer (150 mM KCl, 10 mM Tris–HCl [pH 8.3], 1 mM EDTA) at 65 °C for 90 min. After the mixture was further incubated at 42 °C for 3 min, 37 °C for 3 min and 25 °C for 3 min, nucleotides were precipitated, air dried, and then dissolved in Reverse Transcriptase buffer (25 µL) containing 0.1% DPC, 1x RT buffer, 1 mM dNTP, 50 Units RNase Inhibitor (WAKO chemicals, Tokyo, Japan) and 125 Units ReverTra Ace (Toyobo), and incubated at 42 °C for 60 min. Next, 100 µL of RNase reaction mixture (100 mM NaCl, 10 mM Tris–HCl [pH 7.5], 1 mM EDTA, 100 µg/mL sermon sperm DNA, 20 µg/mL RNase A) was added, and the mixture was incubated at 37 °C for 15 min. The extended DNA strand from the 32P-labeled PAR + 77 was ethanol precipitated and analyzed with an 8 M urea-6% acrylamide sequencing gel.
Construction of PU.1 (Spi-1) expression plasmids
Molecular cloning of the cDNA from RT-PCR was carried out as described previously (Uchiumi et al. 2004). Total RNAs were extracted from HL-60 cells with Trizol reagent (Invitrogen, Carlsbad, CA). cDNAs were synthesized from 5 µg of total RNA with random primers and 200 U of MuLV reverse transcriptase (Toyobo). Then 1/10 volume of each cDNA product was amplified by PCR. Sequences for the human PU.1 (Spi-1) primers were determined from the GenBank database (Accession No. X52056 [GenBank] ) and were Spi-1S, 5'-GCGAATTCAATGGAAGGGTTTC CCCTCG-3' and Spi-1A, 5'-GCGAATTCGCGTGGGGCG GGTGGCGCCGC-3'.
Amplification conditions were 30 cycles of 94 °C for 30 s, 55 °C for 30 s and 68 °C for 4 min. PCR products (0.8-kbp) were digested with EcoRI and subcloned into the EcoRI site of the pBluescript II KS+ (Stratagene, La Jolla, CA). The resultant plasmid was designated pBS-PU.1#48. After determination of the nucleotide sequences of the PCR products, the EcoRI fragment was ligated into the EcoRI sites of the pcDNA3.1B (Invitrogen) and pGEX-3X (Amersham Biosciences) to generate pcDNAPU.1 and pGEX-PU.1, which were used for transient transfection assays and transformation of Escherichia coli. BL-21, respectively.
Expression and purification of a GST–PU.1 fusion protein
The GST–PU.1 fusion protein was extracted from transformed E. coli BL-21 cells and further purified by Glutathione Sepharose 4B (Amersham Biosciences) column chromatography as described (Uchiumi et al. 1996).
Electrophoretic mobility shift analysis (EMSA)
The binding reaction and electrophoresis were carried out as described previously (Uchiumi et al. 1996). In brief, 32P -labeled double-stranded oligonucleotides (1 x 104 cpm approximately 0.1 ng) were incubated with 50–400 ng of GST–PU.1 fusion protein (or 2 µg of nuclear extract from HL-60 cells) and 1 µg of poly (dI-dC) (Amersham Biosciences) in a solution (20 µL) containing 50 mM KCl, 25 mM Hepes–KOH (pH 7.9), 1 mM EDTA, 1 mM DTT and 10% (v/v) glycerol at 25 °C for 30 min. The antibodies used for EMSA were anti-Elk-1 (Sigma), anti-phosphorylated Elk-1 (Sigma) and anti-PU.1 (Santa Cruz Biotechmology, Santa Cruz, CA). The reaction mixture was subjected to electrophoresis on a 4% polyacrylamide gel containing TAE buffer (6.7 mM Tris–HCl [pH 7.5], 3.3 mM sodium acetate, 1 mM EDTA) at 4 °C 150 V constant voltage for 2 h. The gel was then dried and exposed to X-ray film with an intensifying screen at –80 °C for 24 h. DNA–protein complexes were quantified with a BAS2000 Image Analyzing System (Fuji Film Co., Tokyo, Japan).
Transient transfection assays
Plasmid DNAs were transfected into HL-60 or Jurkat cells with the DEAE-dextran method (Uchiumi et al. 1998). Cells (1 x 106) were treated with 0.2 mL TBS (25 mM Tris [pH 7.4], 137 mM NaCl, 5 mM KCl, 0.6 mM Na2HPO4, 0.7 mM CaCl2, 0.5 mM MgCl2) containing 1 µg of the Luc reporter plasmid and 500 µg per mL of DEAE-dextran for 30 min at room temperature. The cells were then washed with TBS to remove the unabsorbed DNA and cultured for another 24 h in DMEM/10% FCS. Next, TPA (5 ng/mL final concentration) was added to the culture and incubated for 24 h. Cells were then collected, and samples for analysis of Luc activity were prepared. Luc assays were performed with the Luciferase Assay System/Dual-Luciferase Reporter Assay System (Promega) as described in the manufacture's technical manual. In brief, the collected cells were lysed with 100 µL of 1 x Cell Culture Lysis Reagent (25 mM Tris-phosphate [pH 7.8], 2 mM DTT, 2 mM 1,2-diaminocyclohexane-N, N, N', N'-tetraacetic acid, 10% glycerol, 1% Triton X-100) or 1 x Passive Lysis Buffer (1 x PLB), mixed, and centrifuged at 12 000 g for 5 s. The supernatant was transferred to a new tube and stored at –80 °C until used. Luc assay reagent (45 µL) was added to 10 µL of protein sample and mixed briefly. For the dual-Luc assay, Luc assay reagent II (45 µL) was added to 10 µL of 1 x PLB extract and then added Stop & Glo reagent (45 µL). Chemiluminescence was immediately measured for 7.5 s with a luminometer Minilumat LB9506 (Berthold, Bad Wildbad, Germany). The light intensity (RLU, Relative Light Units) was referred to directly as Luc activity.
ChIP and PCR analyses
ChIP assays were performed as described previously (Weinmann et al. 2001) with slight modifications. In brief, for each immunoprecipitation, 5 x 106 HL-60 and ML1 cells were cross-linked with a final concentration of 1% formaldehyde (WAKO chemicals) added directly to the media for 10 min at 25 °C. Cross-linking was stopped by the addition of 0.125 M glycine for 5 min at 25 °C. Cells were washed with 1 mL of ice-cold Phosphate Buffered Saline containing 2% FCS and 0.05% NaN3, suspended in 200 µL of SDS-lysis buffer (50 mM Tris–HCl [pH 8.0], 10 mM EDTA–NaOH [pH 8.0], 1% SDS), and kept on ice for 10 min. Chromosomal DNA was sheared to approximately 500-bp by sonication six times for 30 s (with 1-min intervals) with a Branson Sonifier. The lysates were centrifuged at 18 000 g for 10 min at 4 °C, and the supernatant (200 µL) was diluted with 1.35 mL of ChIP buffer (50 mM Tris–HCl [pH 8.0], 167 mM NaCl, 1.1% Triton X-100, 0.11% sodium deoxycholate) and stored at 4 °C (Input fraction). The chromatin lysate (approximately 1.8 mL) was pre-cleared by addition of 60 µL of 50% Protein A agarose/salmon sperm DNA slurry and rotated at 4 °C for 2 h. After centrifugation (8500 g for 10 min), 1 µg of anti-PU.1 or normal rabbit IgG (Santa Cruz) was added to the chromatin lysate (580 µL) and left overnight with rotation at 4 °C. Twenty microliters of 50% Protein A agarose/salmon sperm DNA slurry was then added and incubated 2 h with rotation at 4 °C. The lysate (Unbound fraction) was stored at 4 °C until use. Immunoprecipitates were washed in RIPA buffer (50 mM Tris–HCl [pH 8.0], 1 mM EDTA–NaOH [pH 8.0], 1% Triton X-100, 0.1% SDS, 0.1% sodium deoxycholate) containing 150 mM NaCl, then in RIPA buffer containing 500 mM NaCl, then in a LiCl wash solution (50 mM Tris–HCl [pH 8.0], 167 mM LiCl, 0.25 mM EDTA–NaOH [pH 8.0], 0.5% Nonidet P40, 0.5% sodium deoxycholate), and finally twice in TE. Immunocomplexes were treated with 200 µL of ChIP Direct Elution Buffer (10 mM Tris–HCl [pH 8.0], 300 mM NaCl, 5 mM EDTA–NaOH [pH 8.0], and 0.5% SDS) and incubated at 65 °C overnight. RNA was removed by the addition of 4 µg of RNase A (Sigma) and incubation at 37 °C for 30 min. Then Proteinase K (10 µg) (WAKO chemicals) was added and incubated at 55 °C for 1 h. Quick PrecipTM Plus Solution (Edge BioSystems, Gaithersburg, MD) was added, and DNAs were extracted with phenol–chloroform and ethanol precipitated. The pellets were then resuspended in 50 µL of TE, and a 5 µL aliquot was analyzed by PCR. Total input chromatin from 5 x 106 cells was resuspended in 150 µL of TE and used at a 1 : 10 dilution. PCR was carried out with the PA-102 and PAR-28 primers by 30 cycles of 94 °C for 30 s, 55 °C for 30 s and 68 °C for 2 min.
Western blot analysis
Western blot analysis was performed as described previously (Uchiumi et al. 1999). In brief, cells (1 x 106) were lysed with 100 µL of 1 x RIPA buffer (10 mM Tris–HCl [pH 7.5], 150 mM NaCl, 1 mM PMSF, 1% NP40, 0.1% sodium deoxycholate, 0.1% SDS) and 5 µg of protein was separated by 7.5% SDS-PAGE. Proteins were then transblotted onto nitrocellulose filters, and antibody reactions were performed at 25 °C for 1 h with anti-PU.1, anti-PARP1 (Santa Cruz), anti-β actin, and anti-poly(ADP-ribose) (Calbiochem, San Diego, CA) as primary antibodies. Blots were then incubated with Peroxidase-conjugated anti-rabbit IgG (Calbiochem). The filters were washed with TBST buffer, and chemiluminescence from Super Signal West-Pico (Pierce Biotech, Rockford, IL) was detected by exposing the filter to X-ray film.
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Acknowledgements |
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Footnotes |
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* Correspondence: uchiumi{at}rs.noda.tus.ac.jp
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Received: 24 May 2008
Accepted: 2 September 2008
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