The Hinf-P/P220 Cell Cycle Signaling Pathway Controls Nonhistone

Research Article

The HiNF-P/p220NPAT Cell Cycle Signaling Pathway Controls Nonhistone Target Genes

Ricardo Medina, Margaretha van der Deen, Angela Miele-Chamberland, Rong-Lin Xie, Andre J. van Wijnen, Janet L. Stein, and Gary S. Stein

Department of Cell Biology and Cancer Center, University of Massachusetts Medical School, Worcester, Massachusetts

Abstract Whereas the HiNF-P/p220NPAT pathway plays a key role in HiNF-P and its cofactor p220NPAT are principal factors S-phase entry, the majority of data on cyclin E/CDK2 signaling is regulating histone gene expression at the G -S phase cell centered on the target genes of E2F factors. The E2F proteins had 1 been shown earlier to support S-phase entry and are activated cycle transition. Here,we have investigated whether HiNF-P controls other cell cycle– and cancer-related genes. We used following release from repressive RB-related pocket protein cDNA microarrays to monitor responsiveness of gene expres- complexes through CDK-dependent phosphorylation at the restric- sion to small interfering RNA–mediated depletion of HiNF-P. tion point (10, 11). E2F targets include a number of genes encoding Candidate HiNF-P target genes were examined for the enzymes that are required for nucleotide metabolism as well as presence of HiNF-P recognition motifs, in vitro HiNF-P binding proteins involved in DNA replication and cyclin subunits for CDKs. to DNA,and in vivo association by chromatin immunopreci- The recent realization that cyclin E/CDK2 signaling diverges into pitations and functional reporter gene assays. Of 177 the E2F and HiNF-P pathways (2) necessitates examination of proliferation-related genes we tested,20 are modulated in HiNF-P target genes to gain a more comprehensive understanding HiNF-P–depleted cells and contain putative HiNF-P binding of regulatory events that control the onset of S phase. motifs. We validated that at least three genes (i.e., ATM, To examine the role of HiNF-P in the transcription of nonhistone PRKDC,and CKS2) are HiNF-P dependent and provide data genes, we searched for other genes targeted by HiNF-P in indicating that the DNA damage response is altered in HiNF- asynchronous cells. Here, we have characterized HiNF-P target genes to identify nonhistone cell cycle–related pathways in which P–depleted cells. We conclude that,in addition to histone genes,HiNF-P also regulates expression of nonhistone targets HiNF-P may participate. We have characterized genes that are that influence competency for cell cycle progression. [Cancer modulated in response to HiNF-P small interfering RNA (siRNA) Res 2007;67(21):10334–42] treatment. Genes directly regulated by HiNF-P were defined by in vitro binding assays, reporter gene assays, and chromatin immunoprecipitations that establish binding of HiNF-P to Introduction endogenous chromatin-embedded promoters. One principal result HiNF-P is the gene regulatory end point for the cyclin E/cyclin- of this study is that HiNF-P interacts with divergently transcribed NPAT dependent kinase (CDK)-2/p220 pathway that activates genes encoding proteins that are involved in DNA damage multiple histone H4 genes and supports chromatin packaging of response pathways (e.g., ATM-p220NPAT and PRKDC-MCM4). We nascent DNA (1–7). This pathway may be a key component of a also show that HiNF-P deficiency impairs DNA repair. Based on broader cell cycle checkpoint (‘‘S point’’) defined, in part, by these findings, we suggest that HiNF-P may have important cell HiNF-P–responsive genes that are distinct from those encoding cycle regulatory functions that are complementary to its role in histones. The cell cycle regulatory role of HiNF-P in controlling controlling histone gene expression. histone gene expression is functionally coupled to formation of a HiNF-P/p220NPAT transcriptional coactivation complex (2). Whereas p220NPAT colocalizes with HiNF-P and histone genes at Materials and Methods the G1-S phase transition when transcriptional activation occurs, HiNF-P depletion by siRNA. For siRNA-mediated knockdown of HiNF-P this protein seems to be spatially restricted to two to four large mRNA, human T98G glioblastoma, human HeLa S3 cervical adenocarcino- subnuclear foci (related to Cajal bodies; refs. 2, 4–6). Our ma, or human U-2 OS osteosarcoma cells were transfected in six-well plates immunofluorescence data reveal that a significant fraction of with either siCONTROL Non-Targeting siRNA#1 (Dharmacon), Silencer HiNF-P is also located at a large number of other smaller Negative Control #1 siRNA (Ambion, Inc.), or HiNF-P–specific double- subnuclear foci (2). We have shown that expression of HiNF-P is stranded siRNA oligonucleotides (Ambion, Inc.) with Oligofectamine proliferation related (1, 8, 9). Depletion of HiNF-P decreases histone according to the manufacturer’s instructions (Invitrogen). H4 mRNA levels in asynchronous cells and delays progression into Cell cycle and cancer gene expression arrays. DNA-free total RNA S phase following serum stimulation of quiescent cells (1). These samples isolated from control and HiNF-P siRNA–treated T98G cells were used for probe synthesis in the presence of [a-32P]dCTP according to the data show that HiNF-P is a critical component of the cell cycle– manufacturer’s protocol (SuperArray Bioscience Corp.). This probe was dependent mechanisms that respond to cyclin E/CDK2 to achieve subsequently used to hybridize human cell cycle and cancer GEArray competency for S-phase entry. Q Series cDNA gene arrays following the instructions of the same manufacturer. Spot intensities were quantified with a Storm 840 PhosphoImager using ImageQuant 5.0 software (Molecular Dynamics) and normalized to Requests for reprints: Gary S. Stein, Department of Cell Biology and Cancer the controls provided in the array. Hierarchical clustering was used to Center, University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, analyze the data, and cDNAs with normalized signal intensity values that MA 01655. Phone: 508-856-5625; Fax: 508-856-6800; E-mail: [email protected]. I2007American Association for Cancer Research. differed by at least 1.4-fold between siCONTROL and HiNF-P siRNA doi:10.1158/0008-5472.CAN-07-1560 oligonucleotides were further investigated.

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cDNA synthesis and real-time quantitative PCR. Total RNA from Chromatin immunoprecipitations. T98G cells were washed twice with siCONTROL Non-Targeting siRNA#1 (Dharmacon) and HiNF-P siRNA– ice-cold 1 PBS and subsequently cross-linked with 1% formaldehyde in treated HeLa cells was either subjected to DNase I digestion and purified by 1 PBS for 10 min at room temperature with gentle agitation. To quench column chromatography (DNA-free RNA Kit, Zymo Research) or purified the cross-linking reaction, 0.125 M glycine in 1 PBS was added for 5 min. using the RNeasy Plus Mini Kit (Qiagen) and used to prepare cDNA with the Cells were then washed twice with ice-cold 1 PBS, scraped in lysis buffer SuperScript First-Strand Synthesis System (Invitrogen). Quantitation was [50 mmol/L Tris-HCl (pH 8.0), 150 mmol/L NaCl, 1% NP40, 2 Complete determined using a 7000 sequence detection system (Applied Biosystems) protease inhibitor] and incubated on ice for 20 min. Lysates were sonicated and SYBR Green chemistry (SYBR Green PCR Master Mix, Applied to an average DNA size of 100 to 500 bp and then cleared by centrifugation Biosystems). The relative mRNA expression was determined by the DDCT at 14,000 rpm for 15 min at 4jC. Crude rabbit antiserum (3 AL) against method. The following primer pairs were used for human mRNAs HiNF-P, pre-bleed serum (3 AL), or 2 Ag of RNA polymerase II antibody were (in 5¶-3¶ direction): HiNF-P forward, GAGGAGGATGACCCACTTGA, and added and rotated at 4jC overnight. One-tenth volume of protein A/G reverse, TCAGCTTGGTGTGGTAGCAG; H4/n forward AGCTGTCTATCG- beads (Santa Cruz Biotechnology) was added for 1 h at 4jC. Beads were GGCTCCAG, and reverse, CCTTTGCCTAAGCCTTTTCC; ATM forward, then washed consecutively with the following buffers: low-salt [20 mmol/L CTGTGGTGGAGGGAAGATGT, and reverse, GTTGATGAGGGGATTGCTGT; Tris-HCl (pH 8.0), 150 mmol/L NaCl, 1% Triton X-100, 2 mmol/L EDTA, p220NPAT forward, TCCAGCCTGCTTACTGTCCT, and reverse, AGCAAA- 1 Complete protease inhibitor], high-salt [20 mmol/L Tris-HCl (pH 8.0), CCTTGGGGAACTTT; PRKDC forward, TGCAGCTGATTCACTGGTTC, and 500 mmol/L NaCl, 1% Triton X-100, 2 mmol/L EDTA], LiCl [10 mmol/L reverse, TCGAATACACCGACCACAAA; MCM4 forward, TGAAGCCATT- Tris-HCl (pH 8.0), 250 mmol/L LiCl, 1% deoxycholate, 1% NP40, 1 mmol/L GATGTGGAAG, and reverse, GGCACTCATCCCCGTAGTAA; CKS2 forward, EDTA], and three washes with Tris-EDTA [10 mmol/L Tris-HCl (pH 8.0), ACCGGCATGTTATGTTACCC, and reverse, TGTGGTTCTGGCTCATGAAT; 1 mmol/L EDTA]. Protein/DNA complexes were eluted twice with elution and RB1 forward, TTCACCCTTACGGATTCCTG, and reverse, AGTCCC- buffer (1% SDS, 100 mmol/L NaHCO3) at room temperature. One-tenth GAATGATTCACCAA. volume of 3 mol/L sodium acetate (pH 5.2) was added and samples were Differences in RNA levels between control and HiNF-P–specific siRNA incubated at 65jC overnight to reverse cross-links. Genomic DNA was oligonucleotides were elevated using general linear mixed models (12). purified by phenol-chloroform extraction followed by isopropanol precip- Models were fit by restricted maximum likelihood estimation (13) using the itation with 5 to 20 Ag of glycogen carrier and dissolved in resuspension SAS Proc Mixed procedure (14). In the presence of significant differences buffer (10 mmol/L Tris-HCl, pH 8.0). among means, pairwise comparisons were made using Fisher’s least Analysis of chromatin immunoprecipitations by real-time quanti- significant difference test using the estimated covariance matrix to account tative PCR. Quantitation of DNA from chromatin immunoprecipitation for correlated observations (15). A Bonferroni correction was applied to the samples was achieved by real-time quantitative PCR using a 7000 sequence pairwise P values to compensate for the additive error due to multiple detection system (Applied Biosystems) and SYBR Green chemistry (SYBR comparisons. The distributional characteristics of outcome measures were Green PCR Master Mix, Applied Biosystems). The amount of DNA was evaluated by applying the Kolmogorov-Smirnov goodness of fit test for determined using a standard curve and is expressed as percentage of input. normality (16) to residuals from fitted linear models and by inspection of PCR conditions were 95jC for 10 min and 40 cycles of 95jC for 15 s and frequency histograms of these residuals. In some cases, natural logarithms 60jC for 1 min. We also carried out a dissociation protocol to ensure that a of outcomes were applied to better approximate normally distributed single peak was obtained. The following primer pairs were used for residuals. All computations were done using the SAS version 9.1.3 (SAS chromatin immunoprecipitation analyses: H4/n forward, AGCTGTC- Institute, 2006) and SPSS version 14 (SPSS, Inc., 2005) statistical software TATCGGGCTCCAG, and reverse, CCTTTGCCTAAGCCTTTTCC; ATM- packages. Statistical significance is defined as P < 0.05. p220NPAT (+51) forward, TCCTTCTGTCCAGCATAGCC, and reverse, GT- Electrophoretic mobility shift assay. Binding of HiNF-P to the GGTTCCTGCTGTGGTTTT; p220NPAT-ATM (+241) forward, GGTTCAATT- specified oligonucleotides was assessed with in vitro transcribed/translated CAGGGCGTTTA, and reverse, TTGACTCCTCCCTCTCCTCA; PRKDC- HiNF-P protein produced by the TNT Coupled Reticulocyte Lysate System MCM4 (+129) forward, CGCGACAAGGACAAGCTC, and reverse, CCGCC- (Promega). In vitro DNA binding reactions were done by combining 1 ALof TTTCCACGGTAAC; and CKS2 (160) forward, CAGATCTCTGATTGGCT- in vitro transcribed/translated HiNF-P protein in 8 AL of protein buffer GACC, and reverse, CCAACGATCCGGATTTGA. [final concentrations of 8 mmol/L HEPES pH 7.5, 0.08 mmol/L EDTA pH 8.0, Reporter gene assays. Segments spanning the intergenic region of the 50 mmol/L KCl, 10% glycerol, 1 Complete protease inhibitor (Roche), ATM and p220NPAT loci were cloned into the pGL2-basic luciferase vector A NPAT 1 mmol/L NaF, and 1 mmol/L Na3VO4] with 10 L of a DNA mixture (Promega) using the following primers (in 5¶-3¶ direction): for p220 containing 20 fmol of labeled, double-stranded oligonucleotide in DNA promoter, GCTGCAACGCGTATCCCGACTCCTCTCGCCT (the MluIsiteis buffer (final concentrations of 0.1 Ag/AL of salmon sperm DNA, 1 mmol/L underlined) and CTTACCAGATCTCAATACAAGCCGGGCTACGTCCGAGGG- DTT, 0.5 mmol/L MgCl2, 0.1 mmol/L ZnCl2). Where indicated, unlabeled TAACAAcaaGATCAA (the BglII site is underlined and the mutated ATG is in oligonucleotide competitor (in 1 AL) was added at 25- to 100-fold molar lowercase); for ATM promoter: TTGGCCACGCGTGTCCTTCTGTCCAGCA- excess. Mixtures were incubated for 20 min at room temperature, and the TAGCCG (the MluI site is underlined) and GCGCAAGATCTAGGGTTCAATT- protein/DNA complexes were then separated on a 4% (40:1) native CAGGGCGTTTA (the BglII site is underlined). The splice acceptor site was polyacrylamide gel using 1 Tris-borate EDTA as running buffer at 4jC. mutated using the following primer (only top strand is shown) in 5¶ to 3¶ Gels were dried and exposed to BioMax XAR (Kodak) films at 80jC. In direction: TTCCGAGTGCAGaGcTAGGGGCGCGGAGGC (the mutated affinity competition experiments, signal intensities of bound DNA nucleotides are in lowercase). The ATG translational start codon and splice [electrophoretic mobility shift assay (EMSA) bands] were quantified with acceptor sites were mutated to prevent formation of unintended fusion an AlphaImager 2200 (Alpha Innotech Corp.) and expressed as percentage protein and chimeric mRNAs. The PRKDC promoter was cloned into the of wild-type binding in the absence of any specific competitor. Oligonu- pGL3-basic luciferase vector (Promega) using the following primers (in 5¶-3¶ cleotides used were (in 5¶-3¶ direction) an optimized HiNF-P binding site direction): CGACGCGTAGTGCTCGGAGTACCTG (the MluI site is underlined) based on its recognition sequence in site II of the H4/n gene, CTTCAGG- and GGTAGATCTCCCGGACCCGGAAATG (the BglII site is underlined). TTTTCAATCTGGTCCGATACT; HiNF-P mutant, CTTCAGGTTTT- Transient transfection of U-2 OS cells with FuGENE6 (Roche) was carried CAATCTTCTACGATACT (mutated nucleotides are underlined); ATM, out in six-well plates, with cells seeded at a density of 0.12 106 per CAATACAAGCCGGGCTACGTCCGAGGGT; p220NPAT, GAGGAGGT- well. The next day, cells were transfected with 200 ng of the luciferase TATTGGCCAAGTCCGCTAAG; PRKDC-MCM4, CGCGGAGCCGACGG- construct p220NPAT-luc, ATM-luc, or PRKDC-luc, or the negative control GAACGTCCGCGCTG; CKS2, TCGCCGCCGCCTCGCAAAGTCCGCTTCC; promoterless Luc vector (pGL-luc), and cotransfected with the expression RB1 (proximal), TTCCGCCCGCGGCGTCACGTCCGCGAGG; RB1 (distal), vector FLAG-HiNF-P (25 ng), p220NPAT (200 ng), or empty vector. The total GGTTCTGGGTAGAAGCACGTCCGGGCCG; and a nonspecific competitor, amount of DNA was kept the same in every transfection with empty vector ATTCGATCGGGGCGGGGCGAGC. plasmid DNA. Cells were harvested 24 to 28 h after transfection and cell www.aacrjournals.org 10335 Cancer Res 2007; 67: (21). November 1, 2007

Figure 1. Identification of HiNF-P target genes by siRNA depletion using cDNA arrays. A, DNA-free total RNA samples isolated from control and HiNF-P siRNA– treated T98G cells were used for synthesis of radioactive cDNA pools that were hybridized in duplicate to cell cycle and cancer gene arrays. A total of 10 and 18 HiNF-P–dependent genes were identified in cell cycle arrays and 6 and 32 genes in cancer arrays, which are consistently up-regulated (") or down-regulated (#). Subsets of these genes contain the HiNF-P core motif 5¶-GTCCG-3¶. HiNF-P protein depletion was confirmed by Western blotting. Bottom, protein samples of control (siC) and HiNF-P (siPB) siRNA oligonucleotide–treated T98G cells (CDK2 is shown as a loading control). B, cell cycle–related functions of six HiNF-P target genes that we selected for further analysis. lysates were measured for luciferase activity and normalized to Renilla Immunofluorescence microscopy and ;-irradiation of cells. U-2 OS (phRL-null) activity (dual-luciferase reporter assay system, Promega). cells were grown in six-well plates with or without coverslips (Fisher Significant differences were determined in three independent experiments Scientific), seeded at a density of 80,000 per well, and treated after by Student’s t test (**, P < 0.01). 24 h with siRNA oligonucleotides for 48 h as described above. DNA

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Figure 2. Validation of HiNF-P target genes by real-time quantitative PCR. A, two distinct siRNA oligonucleotides (siRNA_HiNF-P A and siRNA_HiNF-P B) were used to deplete HiNF-P in HeLa cells. Total RNA was converted to cDNA and analyzed by real-time quantitative PCR to validate HiNF-P target genes. The mRNA levels of each gene are presented relative to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA levels. HiNF-P mRNA was effectively depleted by the siRNA treatment with a commensurate decrease in the expression of the prototypical HiNF-P target gene histone H4/n. HiNF-P protein depletion was confirmed by Western blotting. Bottom, protein samples of control (siC) and two HiNF-P siRNA–treated (siPA and siPB) HeLa cells (CDK2 is shown as a loading control). B, each of the genes selected from the array analysis was evaluated for mRNA levels in cells depleted for HiNF-P by two distinct siRNAs. Bottom, a drawing that shows HiNF-P core motifs (y) within the genomic organization of the selected genes. For statistical analysis, see Materials and Methods. Four independent experiments were carried out (**, P V 0.01; *, P V 0.05). damage was achieved with g-irradiation (2 Gy; 137Cs). Cells were allowed the cell cycle array and 15 from the cancer array contain the to recover for 6 h and prepared for either Western blotting (whole-cell HiNF-P core binding motif; of these, two genes are in common to extract) or in situ immunofluorescence microscopy. Antibody staining was both types of arrays (Fig. 1). Thus, 20 genes modulated by HiNF-P done by incubating whole-cell preparations with mouse monoclonal anti– siRNA depletion contain a putative HiNF-P binding site. phospho-histone H2A.X (Ser139), clone JBW301 (Upstate), at 1:2,000 dilution We selected six genes for follow-up studies based on their for 1 h at 37jC. The secondary antibody [Alexa 568 goat anti-mouse linkage to S-phase functions. Four of these genes represent two immunoglobulin G (IgG); Molecular Probes] was used at 1:800 dilution. NPAT Immunostaining of cell preparations was captured with an epifluorescence pairs of divergently transcribed genes (i.e., ATM/p220 and microscope Zeiss Axioplan II equipped with a charge-coupled device PRKDC/MCM4) that share short intergenic promoter regions of camera. Digital images were acquired and analyzed with MetaMorph 545 and 750 bp, respectively. In addition, we also analyzed two soli- software (Molecular Devices). tary genes (CKS2 and RB1); the RB1 gene was selected because Sekimata and Homma (19) have previously shown that MBD2- binding zinc finger (MIZF)/HiNF-P is able to bind to and repress Results the RB1 promoter. We validated HiNF-P responsiveness of these HiNF-P regulates expression of nonhistone genes. To assess genes by real-time quantitative PCR analysis of total cellular RNA whether the histone gene activator HiNF-P can participate in from HiNF-P–depleted cells. HeLa cells were treated with two regulation of nonhistone genes, we monitored a panel of cell growth distinct HiNF-P specific siRNA oligonucleotides at 25 nmol/L regulatory genes for changes in expression in HiNF-P–depleted each. Each siRNA oligonucleotide reduces HiNF-P mRNA levels NPAT cells. HiNF-P uses the G1-S phase–related cofactor p220 to by 68% to 76% and leads to a f44% decrease in mRNA levels for activate histone H4 genes, but other cofactors may synergize with the two identical histone genes H4/n and H4/o (Fig. 2A) that HiNF-P in other cell cycle stages. Because one cannot assume a represent prototypical HiNF-P–responsive genes (1–3, 20). We find priori that nonhistone targets are only regulated at the G1-S that HiNF-P siRNA treatment reduces the levels of the ATM and transition, these studies were done using asynchronous cells. PRKDC mRNAs, as well as CKS2 mRNA (Fig. 2B), in a statistically Total RNA from siRNA-treated T98G cells was used to carry out significant manner, although minor changes observed in MCM4 duplicate screens with two distinct cDNA micro-arrays, with each and p220NPAT levels are not significant. Taken together, these containing 96 cell cycle– or cancer-related genes; 15 genes are results indicate that HiNF-P depletion regulates the expression of common to both arrays (Fig. 1). Of 177 independent genes, several nonhistone genes. depletion of HiNF-P modulates expression of 64 genes. To identify Identification of HiNF-P binding sites in nonhistone genes. direct targets among these genes, we searched for similarities with To determine whether the six identified HiNF-P–responsive the HiNF-P consensus motif present in the cell cycle regulatory site genes directly bind HiNF-P, we carried out EMSAs. We used II of human histone H4 genes. Our search strategy focused on the putative HiNF-P recognition motifs located in the target gene core motif 5¶-GTCCG-3¶ (with redundant flanking sequences, see promoters as probes. As previously established (17), HiNF-P Fig. 3B), which encompasses key base pair contacts for HiNF-P binds with high affinity to its recognition motif in the H4/n gene (17, 18). We determined that seven HiNF-P–responsive genes from (Fig. 3A). This HiNF-P protein/DNA complex is competed www.aacrjournals.org 10337 Cancer Res 2007; 67: (21). November 1, 2007

Downloaded from cancerres.aacrjournals.org on October 2, 2021. © 2007 American Association for Cancer Research. Cancer Research effectively by the unlabeled wild-type binding site but not by the cleotides (Fig. 3D). These competition experiments indicate that corresponding mutant site or a nonspecific oligonucleotide. We the relative affinity of HiNF-P for its targets is as follows: H4/n > assayed direct binding of HiNF-P to radiolabeled oligonucleotides RB1 > PRKDC-MCM4 > CKS2 > ATM > p220NPAT. With the containing each of the putative binding sites (Fig. 3A, right). We exception of RB1, the relative order of binding affinity correlates find that the sites derived from the PRKDC-MCM4 and RB1 with responsiveness to HiNF-P siRNA treatment. (proximal) genes exhibit robust formation of a HiNF-P protein/ In vivo interaction of HiNF-P with the shared regulatory DNA complex, comparable to that of the canonical HiNF-P regions of putative target genes. We carried out chromatin target gene (histone H4/n gene; Fig. 3A, left), whereas the sites immunoprecipitation analyses to investigate in vivo interaction of from the ATM, p220NPAT, and CKS2 genes form complexes that HiNF-P with the endogenous promoters of the paired ATM- are less intense. Hence, each of the HiNF-P–responsive genes we p220NPAT and PRKDC-MCM4 gene loci, as well as the CKS2 locus, examined contains a bona fide HiNF-P recognition motif, albeit within the context of nucleosomal organization. As we have the relative strength of binding differs among these genes. previously established (1–3), HiNF-P and RNA polymerase II both Alignment of these motifs reveals a nonhistone HiNF-P bind to the human histone H4/n promoter in actively proliferating consensus that is related to, but distinct from, the histone H4 cells (Fig. 4A). HiNF-P and RNA polymerase II each exhibit subtype–specific sequence (Fig. 3B). Competition assays with chromatin immunoprecipitation DNA enrichment for the H4/n HiNF-P binding elements in nonhistone genes reveal that sites in locus that is f30-fold above IgG background levels; because the the PRKDC-MCM4 and RB1 (proximal) genes compete with an primer set also amplifies a duplicate of the H4/n gene (i.e., H4/o; efficiency that is comparable to that of the H4/n-related ref. 20), the values we measured are doubled relative to single-copy element. Furthermore, the sites in the ATM, p220NPAT,CKS2, genes. We applied two probe sets that interrogate HiNF-P binding and RB1 (distal) genes exhibit moderate competition, suggesting to each of the two sites within the intergenic regulatory region of that HiNF-P interacts with reduced affinity with these latter the ATM-p220NPAT locus: one set that examines the single HiNF-P elements (Fig. 3C). Relative differences in strength of binding are element in the PRKDC-MCM4 locus and another set for the solitary also reflected by titration experiments with competitor oligonu- CKS2 gene. We observed strong binding to both ATM-p220NPAT

Figure 3. Interaction of HiNF-P protein and its binding core motif in selected target genes. A, oligonucleotides for each target gene were labeled and used independently as probes to detect direct HiNF-P binding. Left, HiNF-P binding to a probe spanning its classic site in the histone H4/n gene is shown as control. Right, binding of HiNF-P to radiolabeled probes of nonhistone target genes. Competition experiments with unlabeled oligonucleotides were done at 100-fold molar excess. The arrowheads (P) indicate the HiNF-P/DNA complexes (short exposure, 10 h; long exposure, 96 h). B, alignment of sequences was done using Web-based resources (Kalign and BoxShade). MIZF protein is identical to HiNF-P; the MIZF consensus binding sequence is taken from Sekimata and Homma (19). The nonhistone HiNF-P consensus sequence represents an all-inclusive redundant motif that is based on all the validated HiNF-P binding sequences presented here with the exception of the distal site of RB1, which is a weak binding site. Symbols used: V, Aor CorG;H, AorCorT;B, C or G or T; S, CorG;M, AorC;K, GorT.C, EMSAs were done with in vitro transcribed/ translated HiNF-P protein and radiolabeled probes spanning putative HiNF-P binding sites. The arrowhead (P) indicates the HiNF-P/DNA complex. Competition assays were done with 100-fold molar excess of unlabeled oligonucleotides spanning the HiNF-P binding core motif. The HiNF-P probe sequence, its binding core motif (bold), and the mutated nucleotides (below) are shown at the bottom. D, titration experiments with competitor oligonucleotides reveal relative differences in strength of binding.

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Figure 4. In vivo binding of HiNF-P to nonhistone genes. A, chromatin immunoprecipitation assays were done in T98G cells with a HiNF-P antiserum and its corresponding pre-bleed (control for nonspecific binding) as well as RNA polymerase II (RNA pol II) antibody to define direct HiNF-P target genes. HiNF-P binding to its target gene promoters in chromatin immunoprecipitation samples was detected by real-time quantitative PCR using specific primer sets and expressed as a percentage of input. The known HiNF-P target gene histone H4/n is shown as a reference. HiNF-P and RNA polymerase II show an f30-fold signal above background level (pre-bleed). Chromatin immunoprecipitation data are also presented for the ATM-p220NPAT (B), PRKDC-MCM4 (C), and CKS2 (D) loci. The promoter regions for each of these loci with the corresponding HiNF-P core motifs (y) are indicated on top. Arrowheads, primer sets used in quantitative PCR to amplify HiNF-P target gene promoters.

and PRKDC-MCM4 loci with chromatin immunoprecipitation that the HiNF-P/p220NPAT complex regulates transcription of the signals that are 8- to 11-fold above background (Figs. 4B and C). ATM, p220NPAT, and PRKDC genes. Our findings establish these For comparison, in vivo binding of HiNF-P to the CKS2 locus is three genes as the first nonhistone targets of the HiNF-P/p220NPAT f6-fold above the nonspecific signal observed for IgG (Fig. 4D). signaling pathway. These chromatin immunoprecipitation data are in a similar range The DNA damage pathway is altered in HiNF-P–depleted (i.e., 0.23–0.70% of input) as histone H4 genes (0.94% of input for cells. To examine the physiologic consequences of HiNF-P H4/n). Therefore, our results validate the interaction of HiNF-P controlling the ATM and PRKDC genes, we tested whether with these nonhistone genes and establish that they are direct depletion of HiNF-P alters the DNA damage pathway. To assess targets of HiNF-P. Hence, HiNF-P controls the genes participating DNA damage, we monitored phosphorylation of H2A.X protein in cell cycle pathways that are distinct from genes required for (gH2A.X) in HiNF-P–depleted U-2 OS cells. Because gH2A.X histone gene expression. concentrates at sites of DNA damage, reduction of gH2A.X levels HiNF-P/p220NPAT complex bidirectionally activates the and disappearance of gH2A.X nuclear foci together permit NPAT divergently transcribed ATM-p220 and PRKDC loci. To test assessment of the recovery of cells from g-irradiation–induced whether HiNF-P, together with its coactivator p220NPAT,can DNA lesions. Cells were treated with control and HiNF-P–specific regulate the promoters of either the ATM or p220NPAT gene, we siRNA oligonucleotides for 48 h before g-irradiation (2 Gy) generated luciferase reporter vectors in which the intergenic region treatment. We monitored gH2A.X after 6 h of recovery and separating the genes was inserted in both orientations. We find observed that depletion of HiNF-P elevates gH2A.X protein levels that forced coexpression of HiNF-P and p220NPAT activates the by 1.9-fold (Fig. 5B). Furthermore, examination of HiNF-P siRNA– ATM promoter by 4.3-fold relative to basal luciferase activity treated cells by in situ immunofluorescence microscopy reveals (Fig. 5A). Similarly, the promoter for the p220NPAT gene in the that loss of HiNF-P increases the size and intensity of gH2A.X opposite orientation is activated 2.9-fold. In addition, the PRKDC nuclear foci (Fig. 5C). Thus, the DNA damage pathway is altered in promoter is activated 2.1-fold by HiNF-P/p220NPAT relative to HiNF-P–depleted cells. Taken together, these results show that control with empty expression vector, whereas the promoterless HiNF-P deficiency alters DNA damage pathways, perhaps in part by reporter plasmid is not appreciably enhanced. These data show controlling the ATM and PRKDC genes. www.aacrjournals.org 10339 Cancer Res 2007; 67: (21). November 1, 2007

Figure 5. Transcriptional activation of HiNF-P target gene promoter and alteration of the DNA damage pathway. A, transcriptional activation by the HiNF-P/p220NPAT complex of the promoters of the ATM, p220NPAT , and PRKDC genes. The empty pGL-luc vector was used as a negative control. Coexpression experiments were done with reciprocal amounts of expression vectors with or without cDNAs for HiNF-P and p220NPAT to maintain a constant level of DNA in each transfection. Firefly luciferase activity monitored as reporter readout was normalized to Renilla luciferase activity driven by a promoterless plasmid. Statistical significance was determined by the Student’s t test for three independent experiments (**, P < 0.01). Alterations in the DNA damage response were monitored by evaluating gH2A.X protein levels by Western blotting and in situ immunofluorescence microscopy. B, quantitation of band intensities (Western blots, left) for two independent experiments after g-irradiation and 6-h recovery. Cells were treated with control (NC) and HiNF-P–specific (siPB) siRNA oligonucleotides to deplete HiNF-P (Western blot, bottom) 48h before irradiation (2 Gy) treatment (*, P < 0.05). C, left, percentage of cells with low-intensity or high-intensity foci in U-2 OS cells treated with one of two HiNF-P (siPA and siPB) specific siRNA oligonucleotides or control siRNA (NC). Right, the three columns of images are representative microscopic fields of siRNA oligonucleotide–treated cells. The superimposed values represent percentages depicted on the graphs to the left. Images were captured as monochromatic light using MetaMorph software. The images on the top and bottom rows represent high-resolution images to show the appearance of gH2A.X in individuals cells. The images in the middle two rows were further processed using Adobe Photoshop. To improve the visualization of our data, the images were inverted (to yield black signals on a white background) and the intensity distribution (‘‘level’’) was linearly adjusted; all images were processed in the same manner. Arrowheads, high-intensity gH2A.X foci. Dashed boxes surround cells that were shown in the top and bottom rows as dark-field images. Cells were treated for siRNA/g-irradiation as described in (B). HiNF-P protein depletion was confirmed by Western blotting (top left); the blot shows protein samples of control (NC) and two HiNF-P siRNA–treated (siPA and siPB) U-2 OS cells (CDK2 is shown as a loading control).

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Discussion in vitro. Our functional siRNA results show that HiNF-P is rate- In this study, we have shown that the histone gene regulatory limiting for at least three of these genes (i.e., ATM, PRKDC, and complex HiNF-P/p220NPAT can control cell cycle– and cancer- CKS2). Furthermore, HiNF-P (1, 17, 33) is identical to the MIZF related gene targets beyond histones. By combined application of protein that was independently isolated in yeast two-hybrid screens cDNA gene arrays, siRNA depletion, EMSAs, chromatin immuno- (34). Sekimata and Homma (19) subsequently recognized that MIZF precipitations, and reporter gene assays, we have defined several has sequence-specific DNA binding activity and defined the binding genes that are directly regulated by HiNF-P/p220NPAT, including motif by CASTing analysis with bacterially produced recombinant ATM (21–23), PRKDC (24, 25), and CKS2 (26). Each of these targets glutathione S-transferase (GST)-MIZF/HiNF-P fusion protein. is either directly or indirectly related to competency for cell cycle These studies showed that MIZF is able to bind and repress the progression, indicating novel cell growth regulatory roles for the RB promoter in HEK293 cells. In our study, we have confirmed HiNF-P/p220NPAT complex. that HiNF-P/MIZF interacts with two recognition motifs (proximal HiNF-P regulation of the genes for ATM and PRKDC suggests and distal) in the RB promoter. However, siRNA depletion reveals functional linkage to pathways involved in repair DNA synthesis. that HiNF-P is not rate-limiting for endogenous RB1 expression in The ATM gene encodes an important kinase that is stimulated T98G and HeLa cells. In addition, the H4 subtype consensus that we after genomic insult and induces the p53-p21 DNA damage had earlier identified as the recognition element for HiNF-P (1, 3, 17, response pathway (27). Similarly, the PRKDC kinase participates 33) is significantly longer than the MIZF consensus site defined in the nonhomologous end joining pathway that facilitates repair by Sekimata and Homma (19). Here, using in vitro EMSAs and of double-strand DNA breaks to maintain genome integrity (28, 29). in vivo chromatin immunoprecipitation analysis with HiNF-P– Our data show that HiNF-P deficiency alters the DNA damage specific antibodies, we have experimentally delineated a distinct response, which is reflected by increased levels of gH2A.X protein nonhistone consensus element for HiNF-P. This motif is also larger as well as the intensity and size of gH2A.X nuclear foci. Combined than the MIZF consensus sequence, suggesting that endogenous with the direct regulation of the ATM and PRKDC genes by HiNF-P, HiNF-P/MIZF recognizes more extended sequence motifs than our findings suggest a new function for this transcription factor in the recombinant GST-MIZF fusion protein. In conclusion, our findings indicate that the cyclin E/CDK2/ DNA damage response pathways that may affect competency for NPAT cell cycle progression. p220 /HiNF-P pathway participates in the control of several We have also provided evidence indicating that the CKS2 gene is distinct cell cycle– and cancer-related genes. The identification of a nonhistone target of HiNF-P. CKS2 belongs to a family of small novel nonhistone target genes indicates that HiNF-P has regulatory proteins (9–18 kDa) with two human homologues (CKS1 and CKS2; roles beyond cell cycle control of histone genes at the G1-S phase SUC1 transition. Our data support the concept that the response of both related to p13 ) that modulate the levels and/or activities of NPAT the cyclin/cyclin-dependent kinase complexes. CKS1 serves as an the E2F/RB and p220 /HiNF-P pathways to cyclin E/CDK2 essential cofactor for the ubiquitin-mediated proteolysis of the signaling activates two distinct regulatory programs that together CDK inhibitor p27Kip1 and p130, which are critical regulators of support cell cycle progression. the G1-S phase transition (30, 31). CKS2 has been shown to be important in controlling the first metaphase-anaphase transition of Acknowledgments mammalian meiosis (32). Oocytes must generate storage pools of Received 4/27/2007; revised 7/27/2007; accepted 8/16/2007. maternal histone mRNAs in anticipation of the rapid cleavage Grant support: NIH grant GM032010. The costs of publication of this article were defrayed in part by the payment of page stages that follow fertilization. HiNF-P is known to be expressed in charges. This article must therefore be hereby marked advertisement in accordance oocytes and may thus be linked to CKS2-dependent regulatory with 18 U.S.C. Section 1734 solely to indicate this fact. events associated with mammalian meiosis. We thank Judy Rask for expert assistance in the preparation of the manuscript and the members of our research group, especially Jitesh Pratap and Kaleem Zaidi, for This study has defined multiple recognition motifs in at least six stimulating discussions. We also thank Kaleem Zaidi for assistance in acquiring nonhistone genes that interact with native HiNF-P in vivo and/or microscopy images.

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