SHPRH regulates rRNA transcription by recognizing the histone code in an mTOR-dependent manner

Deokjae Lee, Jungeun An, Young-Un Park, Hungjiun Liaw, Roger Woodgate, Jun Hong Park, and Kyungjae Myung
PNAS April 25, 2017 114 (17) E3424-E3433; first published April 11, 2017 https://doi.org/10.1073/pnas.1701978114

  1. Edited by Karlene Cimprich, Stanford University, Stanford, CA, and accepted by Editorial Board Member Philip C. Hanawalt March 13, 2017 (received for review February 8, 2017)

Significance

Transcription of ribosomal RNA (rRNA), which composes the ribosome with other proteins, is tightly regulated to maintain the right number of ribosomes. Many DNA repair proteins have functions in addition to their role in DNA repair. We provide evidence that SHPRH functioning in DNA repair at stalled DNA replication forks recognizes epigenetic histone codes of rDNA through its plant homeodomain (PHD) and modulates 47S rRNA transcription. SHPRH bound to rDNA promoters and promoted RNA polymerase I recruitment for rRNA transcription. SHPRH localization to the rDNA promoter was inhibited by trimethylation of histone H3 lysine 4, which is a mark of the rDNA promoter at poised status on starvation. Collectively, we suggest a mechanism controlling 47S rRNA transcription by SHPRH in a histone methylation-dependent manner.

Abstract

Many DNA repair proteins have additional functions other than their roles in DNA repair. In addition to catalyzing PCNA polyubiquitylation in response to the stalling of DNA replication, SHPRH has the additional function of facilitating rRNA transcription by localizing to the ribosomal DNA (rDNA) promoter in the nucleoli. SHPRH was recruited to the rDNA promoter using its plant homeodomain (PHD), which interacts with histone H3 when the fourth lysine of H3 is not trimethylated. SHPRH enrichment at the rDNA promoter was inhibited by cell starvation, by treatment with actinomycin D or rapamycin, or by depletion of CHD4. SHPRH also physically interacted with the RNA polymerase I complex. Taken together, we provide evidence that SHPRH functions in rRNA transcription through its interaction with histone H3 in a mammalian target of rapamycin (mTOR)-dependent manner.

Human ribosomal DNA (rDNA) is composed of hundreds of tandem repeats of 42.9-kb rDNA units that are organized into transcribed and intergenic regions (1). About one-half the 47S precursor ribosomal RNA (pre-rRNA) genes are actively transcribed, and the other half remain silent (2, 3). Transcription, processing of rRNA, and the assembly of ribosomes take place in the nucleoli (2, 4). Once transcribed in the nucleoli, pre-rRNA is immediately processed into small mature 28S, 18S, and 5.8S rRNAs that, together with ribosomal proteins, make a ribosome. Tight regulation of ribosome biogenesis, including rRNA transcription and synthesis of ribosomal proteins, is important in many biological processes such as cell proliferation, apoptosis, and autophagy (5⇓–7), and is closely associated with metabolic processes. Because of its importance in many metabolic pathways, dysregulation of ribosomal biogenesis is linked to aging and diverse diseases, including anemia and cancers (8⇓⇓⇓–12). 47S pre-rRNA is transcribed by the RNA polymerase I complex, whose activity is controlled by cellular responses to nutritional states, cellular stresses, growth, differentiation, and cell cycle (9). Posttranslational modifications of transcription factors, for example, phosphorylation of upstream binding factor (UBF), help regulate rRNA transcription (13). In addition to posttranslational modifications of transcription factors, nucleolar remodeling complex, NuRD (nucleosome remodeling and deacetylation) complex, and energy-dependent nucleolar silencing complex also affect rRNA transcription by modifying epigenetic signatures of rDNA, as well as histones in the rDNA promoter (14⇓–16). In addition to conventional active and silent histone signatures, the rDNA promoter has another histone signature called a poised state. CHD4 and CSB-containing NuRD complex establish a poised chromatin signature of rDNA that represses but primes rRNA transcription by marking histone H3 with both active (H3 K4me3) and inactive (H3 K27me3) modifications (16). However, it is unclear how these epigenetic changes control the transcription of rRNA.

The mammalian target of rapamycin (mTOR) pathway is a master pathway that controls overall cellular activities, including autophagy, macromolecule biosynthesis, and cell cycle in response to nutrients, stress, and growth factors (17, 18). Indeed, the mTOR pathway controls rRNA transcription and production of ribosomal proteins in response to nutrient, growth factors, and serum (19⇓–21).

Previously, we found SHPRH as a mammalian RAD5 homolog (22, 23). RAD5 functions to avoid the collapse of the DNA replication fork by promoting the bypass of DNA damage that would normally stall the DNA replication fork. RAD5 and its mammalian homolog SHPRH polyubiquitylate proliferating cell nuclear antigen (PCNA) to promote DNA damage bypass via an uncharacterized recombination-dependent pathway (24, 25). Furthermore, there are two RAD5 homologs in mammals, which suggests that functions other than DNA damage bypass might have been developed during evolution (26⇓–28).

Although the epigenetic signature and mTOR-dependent regulation for rRNA transcription are important, it is not clearly understood how RNA polymerase I can be directed to recognize such signatures to determine rRNA transcription. In this study, we report that SHPRH promotes rRNA transcription in a nutrient/mTOR-dependent manner by recruiting RNA polymerase I to the active rDNA promoter. SHPRH localized at the rDNA promoter in the nucleoli through the interaction between its plant homeodomain (PHD) and a specifically modified histone H3. Localization of SHPRH is redistributed into foci in the nucleoli under starvation or exposure to rapamycin that inhibits the mTOR pathway.

Results

SHPRH Forms Starvation-Induced Foci in the Nucleoli in an mTOR-Dependent Manner.

SHPRH and HLTF, homologs of yeast Rad5, were previously identified as E3 ubiquitin ligases that catalyze the polyubiquitylation of PCNA and participate in methyl methane sulfonate (MMS)-induced DNA damage repair (26⇓–28). In contrast to yeast in which Rad5-deficiency causes high sensitivity to DNA-damaging agents, silencing expression of SHPRH or HLTF in human cells or knockouts of SHPRH and HLTF in mice showed mild or no severe sensitivity to DNA damaging agents, respectively (29⇓–31) (Fig. S1 A and B). Therefore, we hypothesized that SHPRH would have a function other than DNA damage repair in mammals. To explore alternative functions, we first determined cellular localization of SHPRH. SHPRH was detected in the nucleus, as well as in the chromatin-bound fraction, when stained with an antibody (3F8) that specifically detects SHPRH (Fig. 1A and Fig. S2 A–C and S3). When cells were cultured for several days without changing media, SHPRH formed nucleolar foci, as judged by colocalization with a nucleolar protein, Fibrillarin, and these foci were not detected in fresh growth media (Fig. 1 A–C).

Fig. 1.
Fig. 1.

Starvation or rapamycin treatment induce the nucleolar SHPRH foci. (A) Nucleolar SHPRH foci were detected in HeLa cells. HeLa cells were immunostained with anti-SHPRH 3F8, anti-Fibrillarin antibody, and DAPI. In addition to broad staining of SHPRH detected in the nuclei, large and dense SHPRH foci were colocalized with the nucleolar marker Fibrillarin. (B and C) Nucleolar SHPRH foci were induced on treatment with 2.5 μg/mL rapamycin for 24 h (B) or starvation with HBSS for 2 h (C) in HeLa cells. Endogenous SHPRH (green) and Fibrillarin (red) were shown. (D) Nucleolar SHPRH foci were sustained during starvation. After starvation, the number of nucleolar SHPRH foci per nucleus was counted in HeLa cells after fixation at indicated times. The averages of foci numbers from two independent experiments were plotted with mean ± SEM. (E) SHPRH foci disappeared when cells were recovered from starvation. After starvation in HBSS for 2 h, cells were recovered by supplying normal growth media for indicated times. Foci were counted from more than 74 nuclei. Graphs are presented with mean ± SD.

Fig. S1.
Fig. S1.

Shprh−/−Hltf−/− MEFs did not exhibit significant sensitivity to MMS, cisplatin, and camptothecin. (A) Wild-type and Shprh−/−Hltf−/− MEFs were treated with MMS, cisplatin, and camptothecin as indicated concentration for 48 or 72 h. Cell survival was measured using CyQUANT Cell Proliferation Assay kit (Invitrogen). (B) Wild-type and Shprh−/− MEFs were treated with MMS, and surviving colonies were counted.

Fig. S2.
Fig. S2.

Antibody specifically recognizing SHPRH was generated. (A) HeLa cells were transfected with nontargeting siRNA or ON-TARGET plus siRNA of SHPRH (Thermo Scientific) and incubated for 2 d. Cells were stained with anti-SHPRH 3F8 antibody and DAPI. (B) Nontargeting (N.T.) or SHPRH siRNA-transfected 293T cell lysates were used for Western blotting. Endogenous SHPRH was detected with anti-SHPRH 3F8 antibody. (C) The levels of SHPRH in HeLa cells stably transduced with control and shSHPRH lentivirus (HeLa-pLL3.7/control and HeLa-pLL3.7/SHPRH).

Fig. S3.
Fig. S3.

Majority of SHPRH was located in the chromatin-bound fraction. Exogenously expressed SHPRH-myc-His was expressed in HeLa cells. Cytoplasmic, soluble nuclear, and chromatin-bound fractions were separated and analyzed by Western blotting with anti-myc 9E10 antibody.

If SHPRH foci were formed as a result of DNA damage in long culture condition, they should colocalize with other DNA damage-induced foci. However, we found no evidence that SHPRH colocalized with DNA damage response proteins, including phospho-RPA32, γH2AX, pol η, pol κ, PCNA, phospho-CHK1, 53BP1, XPA, and BRCA1 (Fig. S4 A and B). In addition, there were no SHPRH foci after treatment of cells with 60 J/m2 UV irradiation or 0.01% MMS treatment unless cells were cultured for several days before treatment (Fig. S4 C and D). HP1β is a major heterochromatin protein associated with nucleolar regions (32). The location of nucleolar SHPRH foci did not overlap with HP1β (Fig. S4E).

Fig. S4.

Nucleolar SHPRH foci were not associated with DNA damage repair proteins. (A) HeLa cells were fixed without exposure to exogenous DNA-damaging reagents and stained with anti-SHPRH 3F8 (Alexa488), anti-phospho-RPA32 S4/S8, anti-PCNA, anti-polymerase η, anti-53BP1, anti-γH2AX, anti-phospho-CHK1, anti-polymerase κ, or anti-BRCA1 antibodies (Alexa568). (B–D) HeLa cells either in starved (B and D) or in normal condition (C) were treated with 60 J/m2 UV or 0.01% MMS for 1 h and recovered for 4 h. Fixed cells were stained with anti-SHPRH 3F8, anti-phospho-RPA32 S33, anti-XPA, anti-γH2AX, anti-Fibrillarin antibodies and DAPI. (E) HeLa cells were stained with anti-SHPRH and HP1β. (F) HeLa cells were starved in HBSS for 4 h or treated with 2 mM hydroxyurea (HU) for 6 h. Proteins indicated were detected with antibodies specific for each protein. (G and H) HeLa cells were treated with 2.5 μg/mL rapamycin or starved for 0, 2, 4, 6, and 24 h, and the levels of SHPRH, phosphor-S6, S6, and α-Tubulin, were analyzed. (I) SHPRH foci disappeared when cells were recovered from starvation. After starvation in HBSS for 2 h, cells were recovered by supplying normal growth media for indicated times. Foci were counted from more than 14 nuclei. Graphs are represented as mean ± SEM.

SHPRH foci became more distinct in cells cultured for several days without changing growth media. We therefore hypothesized that cells experiencing nutrient restriction would generate SHPRH foci in the nucleolus. Consistent with our hypothesis, when cells were starved by incubating in HBSS, or treated with 2.5 μg/mL rapamycin that blocks the mTOR pathway and mimics starvation conditions (17, 18, 33), SHPRH foci were clearly detected in the nucleoli (Fig. 1 B and C). Starvation in HBSS did not increase DNA damage responses, including phosphorylation of RPA32 and CHK1; thus, the nucleolar SHPRH foci were not induced by DNA damage responses in starved condition (Fig. S4F). Starvation or rapamycin treatment did not change SHPRH level, suggesting SHPRH foci formation on starvation was not a result of the induction of SHPRH protein level (Fig. S4 G and H). We then investigated kinetics of nucleolar foci formation on starvation condition. SHPRH began forming nucleolar foci 2 h after cells were incubated in HBSS media (Fig. 1D). When HBSS media was replaced with normal growth media, the number of nucleolar SHPRH foci decreased in a time-dependent manner (Fig. 1E and Fig. S4I). Collectively, SHPRH forms nucleolar foci in response to inhibition of the mTOR pathway.

SHPRH Interacts with the Ribosomal DNA Promoter in an mTOR-Dependent Manner.

The mTOR-dependent pathway promotes ribosomal biogenesis and inhibits autophagy (17, 18, 20). Multiple repeated ribosomal DNA (rDNA) clusters are localized in the nucleoli, and some of them are actively transcribed (Fig. 2A) (2, 34, 35). Because SHPRH was found in the chromatin-bound fraction and localized in the nucleoli (Fig. 1A and Fig. S3), we investigated whether SHPRH would bind to specific regions of ribosomal DNA (Fig. 2A). When SHPRH was cross-linked with chromatin and immunoprecipitated, a ribosomal DNA promoter (H42.9) was enriched in precipitates (Fig. 2B). We did not detect the enrichment of the H42.9 region in precipitates of cells whose SHPRH expression was silenced by shRNA, suggesting the enrichment of SHPRH was specific (Fig. 2B and Fig. S2C). Chromatin immunoprecipitation (ChIP) experiments with two different human cell lines, K562 and HEK293, showed enrichment of SHPRH on the rDNA promoter (Fig. S5 A and B). Because starvation or rapamycin treatment induced the formation of nucleolar SHPRH foci (Fig. 1 B and C), we next investigated whether enrichment of SHPRH at the rDNA promoter is also affected by starvation. SHPRH enrichment at the promoter of rDNA was reduced on starvation (Fig. 2C). Instead, starvation as well as rapamycin treatment caused SHPRH enrichment at different locations in rDNA, including H8 (Fig. S5C). Simultaneously, a modified chromatin marker, H3 K4me2, was increased at rDNA H8 and decreased at rDNA H42.9 (Fig. S5 C–E), suggesting SHPRH is redistributed from the rDNA promoter to a different region in the rDNA cluster and forms foci structures when cells were starved. In addition, we compared the location of SHPRH foci with UBF, which binds to active rDNA (16). Although fewer nucleolar proteins and foci of SHPRH were found compared with those of UBF, which is an essential factor for rDNA transcription (36), SHPRH foci were partially colocalized with or outside the UBF positive regions (Fig. S5 F–H and Movie S1). Relative occupancies of UBF in the rRNA transcribed region (H1–H13) were decreased in starved cells where SHPRH was redistributed from the rDNA promoter (Fig. S5I). Thus, starvation induces relocalization of SHPRH to other regions from the active rDNA promoter. SHPRH also interacted with other promoters that are not related to rDNA transcription, although no significant change in their mRNA level by SHPRH depletion was detected (Fig. S5 J and K). Collectively, when the mTOR pathway is inhibited by starvation or rapamycin, SHPRH is released from the rDNA promoter and forms nucleolar foci at a different location in the nucleoli.

Fig. 2.
Fig. 2.

SHPRH is recruited to the 47S rRNA gene promoter. (A) Schematic representation of an individual rDNA unit. The black bar and number indicate the amplified regions for ChIP-qPCR (quantitative PCR) assay. (B and C) ChIP assay with anti-SHPRH 3F8 antibody showed that SHPRH binds to the rDNA promoter. (B) HeLa cells stably transduced with control or shSHPRH lentivirus (HeLa-pLL3.7 control and HeLa-pLL3.7/SHPRH, respectively) were used for ChIP assay with anti-SHPRH 3F8 antibody. Bars indicate the relative values of each rDNA region normalized to input DNA and SHPRH-interacting H1, and represented as mean ± SEM from two independent experiments. (C) Starvation reduced the enrichment of SHPRH at the rDNA promoter (H42.9). HeLa cells were starved with HBSS for 4 h. Precipitated DNA was amplified with H42.9 primers. Data are represented as mean ± SEM from five independent experiments.

Fig. S5.

SHPRH localized at the rDNA promoter in K562 and 293 cells. (A and B) K562 or 293 cells were used for ChIP assay to detect rDNA enrichment of SHPRH with anti-SHPRH 3F8 antibody. Bars indicate the relative values of each rDNA region normalized to input DNA (n = 3). (C and D) 293-TetON-SHPRH-myc-His cells were starved with HBSS for 4 h (C) or treated with 2.5 μg/mL rapamycin for 24 h (D). Expression of SHPRH-myc-His was induced by treatment of doxycycline for 2 d. SHPRH-myc-His was then precipitated using anti-6xHis antibody, and DNAs pulled down were analyzed in triplicate with the indicated primers. (E) 293 cells were grown in normal growth media (bright gray) or starved (dark gray), and histone H3 methylations at rDNA H8 or H42.9 were measured using ChIP assay (n = 3). Graphs are shown as mean ± SEM. (F–H) HeLa cells were grown in normal growth media or in starved condition (G and H) and fixed with 4% paraformaldehyde and stained with anti-SHPRH 3F8 (green), anti-UBF (red) antibodies and DAPI (blue). Images were merged to compare their localization. (G) Representative blow-up image of colocalized foci of UBF and shprh. (H) Merged image showing that there are some foci outside UBF positive signal. (I) The UBF occupancies were measured in normally growing or starved HeLa cells. The relative UBF occupancies were normalized with input and the relative occupancy in H1 with triplicates (mean ± SEM). (J) DNA fragments precipitated with anti-SHPRH in 293 cells were amplified with primer sets against promoters of randomly selected genes (locus A–C, G–K) or intergenic regions (locus D–F). (K) The mRNA levels of selected genes from J were measured using RT-qPCR (n = 4).

The PHD of SHPRH Interacts with Histone H3 When Lysine 4 of H3 Is Not Trimethylated.

In contrast to yeast Rad5 or human HLTF, SHPRH has a PHD (22, 28), which is generally known to interact with specifically modified histone H3 (Fig. 3A) (37⇓–39). To determine specificity of the interaction between SHPRH and histones, individually purified histone proteins were incubated with purified SHPRH (Fig. S6A). Similar to other proteins containing a PHD, only histone H3 was coprecipitated with a full-length SHPRH (Fig. 3B and Fig. S6B). To examine the interaction between SHPRH and histone H3 in more detail, interactions between GST-conjugated SHPRH PHD (GST-PHD) and differentially methylated histone H3 peptides were monitored. Unlike GST protein, which did not bind to any H3 peptide, GST-PHD interacted with various histone H3 peptides except the lysine 4 trimethylated H3 (H3 K4me3) peptide (Fig. 3C). Trimethylation of lysine enhances the hydrophobicity of lysine. We therefore hypothesized that the hydrophilic lysine 4 of H3 would be important for the interaction with SHPRH. To test this hypothesis, mutant H3 peptides (H3 K4A and H3 K9A) lacking hydrophilicity in their lysine residue were used to study in vitro interactions (Fig. 3D). In contrast to the interaction between H3 K9me3 and H3 K9A peptides with GST-PHD, both H3 K4me3 and H3 K4A did not interact with GST-PHD, suggesting that trimethylation at K4 with a high hydrophobicity inhibited the interaction between H3 and SHPRH.

Fig. 3.
Fig. 3.

The PHD of SHPRH is important for the interaction with histone H3. (A) Schematic representation of human SHPRH, HLTF, and yeast Rad5 protein structures (27). Nuclear localization signal sequence (NLS), linker histone H1/H5 domain (H15), PHD, HIRAN, SWI2/SNF2 helicase domains and RING domain are indicated. (B) SHRPH interacted with histone H3. Purified SHPRH-myc-His was incubated with calf histones. Protein complexes were precipitated with Ni-NTA bead and analyzed by Western blotting. (C and D) Mixture of GST and GST-PHD was incubated with indicated synthetic histone H3 peptides. Indicated lysine was mono-, di-, or trimethylated or substituted with alanine. Biotinylated peptides were pulled down with Streptavidin Sepharose. GST and GST-PHD were detected with anti-GST antibody. (E) Alignment of PHDs. Sequences of SHPRH PHD are aligned with PHDs of K4 trimethylated H3-binding BPTF and ING2 and PHDs of unmethylated H3-binding CHD4-PHD2 and PHF21A. Highly conserved sequences are indicated with red or green. Critical H3-binding residues are indicated with dotted box. (F) Changes of the conserved glutamate to alanine (E660A; A in F) or tyrosine (E660Y; Y in F) of SHPRH PHD abolished the interaction of the PHD of SHPRH with H3 peptides. (G) Endogenous SHPRH interacted with histone H3 K4me2, but not with H3 K4me3. 293T lysates were immunoprecipitated with indicated antihistone H3 antibodies or control rabbit antibody. Western blotting was performed with anti-SHPRH 3F8 and antihistone H3 antibodies.

Fig. S6.
Fig. S6.

Purification and expression of SHPRH-myc-His proteins. (A) SHPRH-myc-His was expressed in Sf21 cells and purified by immobilized metal ion chromatography. Purified SHPRH-myc-His protein was analyzed by Coomassie blue staining (lane 2). (B) Purified SHPRH-myc-His and calf histones were mixed and SHPRH complexes were precipitated with anti-myc 9E10 antibody. (C) SHPRH WT or E660A was expressed in HEK293T cells. Indicated histone antibodies were used for immunoprecipitation, and the interaction of SHPRH was detected using anti-myc 9E10 antibody. (D) The levels of ectopically expressed wild-type and E660A mutant SHPRH-myc-His and precipitated wild-type and E660A mutant SHPRH-myc-His in ChIP assay were comparable.

There are two major classes of PHDs (38). The first class of PHD found in BPTF and ING2 interacts with histone H3 carrying K4me3. The second class of PHD found in CHD4 and PHF21A interacts with unmodified, K4me, or K4me2 H3. Negatively charged amino acids such as glutamate or aspartate in the PHD are critical for its interaction with histones (40, 41). To examine whether this requirement for a negatively charged amino acid in the PHD is conserved in SHPRH, we aligned the SHPRH PHD with other PHDs. We found a critical amino acid, a glutamate at amino acid residue 660 (E660) in SHPRH, aligned with negatively charged amino acids of PHDs that interact with unmodified, K4me, or K4me2 H3 (Fig. 3E). When E660 of SHPRH was mutated to a neutral amino acid, alanine (A) or tyrosine (Y), the mutant SHPRH PHDs no longer interacted with histone H3 in vitro (Fig. 3F). Similarly, the exogenously expressed SHPRH PHD mutant protein (E660A) showed less interaction with H3 K4me2 (Fig. S6C). Consistent with the in vitro interactions between H3 and SHPRH, in vivo immunoprecipitation with crosslinking by a histone H3 K4me2 antibody coprecipitated SHPRH, but not with a H3 K4me3 antibody (Fig. 3G).

We next investigated whether the SHPRH PHD is important for the enrichment of SHPRH at the promoter of rDNA. The localization of ectopically expressed wild-type and mutant SHPRH (E660A) proteins at the rDNA promoter was monitored by ChIP. As expected, although the ectopically expressed wild-type SHPRH protein was enriched at the rDNA promoter, the E660A SHPRH mutant was not (Fig. 4A and Fig. S6D).

Fig. 4.
Fig. 4.

rDNA promoter marked by H3 K4me3 was void of SHPRH. (A) SHPRH was targeted to rDNA through its PHD. HeLa cells were transfected with empty vector, SHPRH-myc-His wild-type, and SHPRH-myc-His PHD, which has an E660A mutation. Binding of SHPRH to rDNA promoter (H42.9) was analyzed with ChIP assay. SHPRH was precipitated with anti-6xHis antibody. Data are represented with mean ± SEM from four independent experiments. (B) HeLa cells were transfected with siRNA of nontargeting or SHPRH 3′UTR or (C) starved with HBSS for 6 h. ChIP assay was performed with anti-H3 K4me3 (Left) or anti-H3 K4me2 (Right) antibodies. Bars indicate the relative values amplified with H42.9 primers. Each targeting was normalized to input DNA. Graphs show mean ± SEM from five (B, Left) or four independent experiments.

Unlike most promoters where H3 K4me3 represents an active state, H3 K4me3 at the rDNA promoter denotes a poised state, whereas H3 K4me2 with acetylated H4 represents the active state (16, 42). SHPRH is enriched at the rDNA promoter and released from it on starvation. Thus, we hypothesized that SHPRH depletion or starvation would change the status of methylation in H3 from K4me2 to K4me3. H3 K4me3 at the rDNA promoter was increased and H3 K4me2 decreased after depletion of SHPRH, or on starvation (Fig. 4 B and C). Consistent with our data, it has been reported that the occupancy of H3 K4me3 at the rDNA promoter is increased by serum deprivation (16, 43). Thus, we conclude that the PHD of SHPRH is important for the enrichment of SHPRH at the rDNA promoter by recognizing histone codes.

SHPRH Up-Regulates 47S rRNA Transcription.

Because SHPRH is recruited to the rDNA promoter, we hypothesized that SHPRH would affect the level of pre-rRNA transcription. Indeed, depletion of SHPRH with two individual siRNAs that target the 3′ UTR of SHPRH reduced 47S pre-rRNA levels (Fig. 5A and Fig. S7A). Conversely, SHPRH overexpression increased 47S pre-rRNA levels (Fig. 5B). The inhibition of the mTOR pathway by starvation or by the treatment with rapamycin similarly reduced 47S pre-rRNA levels, and there was no additive or synergistic reduction of pre-rRNA levels by SHPRH depletion (Fig. 5C and Fig. S7B). Thus, SHPRH enhances the level of 47S pre-rRNA in an mTOR-dependent manner. There are two possible ways that SHPRH could increase 47S pre-rRNA levels. One is to enhance pre-rRNA transcription, so as to produce newly synthesized rRNA, and the other is to increase the stability of existing pre-rRNA. To determine which process is promoted by SHPRH, we measured newly transcribed nascent rRNA by pulse-labeling cells with BrUTP that were detected in situ (44) after SHPRH knockdown. We found that depletion of SHPRH reduced the level of newly transcribed rRNA in situ (Fig. 5D and Fig. S7C). Similarly, starved cells showed low levels of newly transcribed nascent rRNA. There was no obvious synergistic or additive effect on the level of pre-rRNA or newly transcribed nascent rRNA on starvation and depletion of SHPRH (Fig. 5 C and D and Fig. S7 B and C). Thus, our observations indicate that SHPRH promotes pre-rRNA transcription and is directly inhibited by starvation.

Fig. 5.
Fig. 5.

SHPRH regulates 47S ribosomal RNA transcription. (A) SHPRH was depleted in HeLa cells by transfecting individual siRNA targeting the 3′UTR of SHPRH for 48 h. Nontargeting (N.T.) siRNA was used as a control. (B) Empty vector or SHPRH-myc-His-expressing vector ectopically expressed in HeLa cells for 24 h. (C) HeLa cells were transfected with nontargeting siRNA (N.T.) or siRNA targeting the SHPRH 3′UTR. Forty-eight hours after transfection, cells were starved with HBSS for 2 h and harvested. (A–C) The level of pre-rRNA was analyzed by reverse transcription and qPCR. RNA levels of pre-rRNA were normalized to β-actin. Data show mean ± SEM from independent two independent experiments for A, five experiments for B, and three experiments for C. (D) HeLa cells were transfected with nontargeting siRNA or siRNA targeting the SHPRH 3′UTR for 48 h. For staining of nascent rRNA, cells were permeabilized and rRNA was synthesized in vitro with Br-UTP. Amount of nascent rRNA was stained with anti-BrdU (BU1/75) antibody, and images were taken with a fixed exposure time. Intensities of nascent rRNA were analyzed with ImageJ (NIH).

Fig. S7.
Fig. S7.

Depletion of SHPRH negatively regulates 47S rRNA transcription. (A) HeLa cells were transfected with nontargeting siRNA, siRNA of SHPRH 3′UTR#1, or siRNA of SHPRH 3′UTR#3 for 2 d. The level of SHPRH mRNA was normalized to β-actin. Graphs show the mean ± SEM from two independent experiments. (B) HeLa cells in which SHPRH was depleted by transfecting siRNA of SHPRH were treated with 2.5 μg/mL rapamycin for 4 h. The level of pre-rRNA and SHPRH mRNA was normalized to β-actin. Graphs show the mean ± SEM from two independent experiments. (C) Representative images of Fig. 5D are shown.

SHPRH Interacts with and Regulates RNA Polymerase I Complex.

RNA polymerase I transcribes 47S pre-rRNA (13). rRNA transcription is directly controlled by transcription factors, as well as indirectly controlled by epigenetic factors (3, 13). Because SHPRH interacted with the active rDNA promoter and increased rRNA transcription, it is possible that SHPRH could directly control rRNA transcription through an interaction with the RNA polymerase I complex. We therefore examined an interaction between SHPRH and the RNA polymerase I complex by coimmunoprecipitation. SHPRH-myc-His was induced in a cell line containing a stable doxycycline-inducible SHPRH construct and was immunoprecipitated with anti-6xHis antibody. The RNA polymerase I subunit, RPA194, UBF that functions to activate RNA polymerase I (13, 45) and histone H3, but not Fibrillarin, was specifically coprecipitated with SHPRH (Fig. 6A). Serial deletion mutant analysis of SHPRH revealed that amino acids between 784 and 1,288 amino acids were responsible for the interaction with RNA polymerase I (Fig. S8 A and B). Consistently, SHPRH without this region could not induce pre-rRNA transcription (Fig. S8C). SHPRH harboring a point mutation in the PHD or really interesting new gene (RING) domain could not induce pre-rRNA transcription, although they still interacted with RNA polymerase I (Fig. S8 B, D, and E). In addition, ChIP experiment showed that RNA polymerase I recruitment to rDNA promoter was down-regulated by SHPRH depletion or starvation (Fig. 6 B and C). Conversely, SHPRH overexpression increased the recruitment of RNA polymerase I to the rDNA promoter (H42.9) and the level of pre-rRNA, which was not observed with the SHPRH PHD mutant (E660A) or RING mutant (C1432A; Fig. 6D and Fig. S8 D and E). Changes of SHPRH PHD mutant or RING mutant's localization would be a result of defects of these mutant proteins localizing to the rDNA promoter (Fig. 4). Under starved condition, the interaction between SHPRH and RNA polymerase I was decreased (Fig. 6E and Fig. S8 F and G). Thus, the PHD and RING domains of SHPRH and its potential cooperation with RNA polymerase I are important for rRNA transcription by recognizing the status of rDNA promoters.

Fig. 6.
Fig. 6.

SHPRH is associated with RNA polymerase I complex in a starvation-dependent manner. (A) SHPRH interacted with RNA polymerase I complex. The expression of SHPRH-myc-His was induced by treatment of doxycycline to 293-TetOn-SHPRH-myc-His cells. Precleared lysates were immunoprecipitated with anti-6xHis antibody. Precipitates were analyzed by Western blotting with anti-6xHis conjugated with HRP, anti-RPA194, anti-UBF, anti-Fibrillarin, and antihistone H3 antibodies. (B–D) RNA polymerase I binding on rDNA was regulated by SHPRH or starvation. (B) SHPRH was stably depleted by infecting HeLa cells with lentivirus expressing shRNA of SHPRH. (C) HeLa cells were starved in HBSS for 4 h. (D) HeLa cells were transfected with empty vector, SHPRH-myc-His wild-type, or SHPRH mutant having an E660A mutation in PHD or a C1432A mutation in RING domain. ChIP assay was performed with anti-RPA194 antibody. Data are represented with mean ± SEM: B, n = 2; C, n = 6; and D, n = 3. A relative amount of binding was normalized to input DNA. (E) The interaction between SHPRH and RNA polymerase I was down-regulated in starved cells. 293-TetOn-SHPRH-myc-His cells were treated with 1 μg/mL doxycycline for 2 d to induce SHPRH-myc-His expression. Cells were starved for 4 h in HBSS and harvested for immunoprecipitation with anti-6xHis antibody.

Fig. S8.
Fig. S8.

Starvation blocked the interaction between SHPRH and RNA polymerase I. (A) Schematic representations of full-length SHPRH and truncated SHPRH. (B) Full-length, mutated or truncated SHPRH were transiently expressed in HEK293T cells. RNA polymerase I was immunoprecipitated, and the bindings of SHPRH and UBF were analyzed. (C) After SHPRH wild-type and serial deletion mutants (SHPRH-SmaI, SHPRH-EcoRV) were expressed, levels of pre-rRNA were measured. (D) The levels of ectopically expressed SHPRH-myc-His in ChIP assay were shown. (E) Empty vector or SHPRH wild-type or mutants (RINGm and PHDm) were transfected in HEK293T cells. The levels of pre-rRNA were measured using RT-qPCR. (F) HEK293T cells were transfected with nontargeting (N.T.) siRNA or siRNA targeting the 3′UTR of SHPRH (SH). Two days after transfection, cells were starved for 8 h and harvested for immunoprecipitation. SHPRH was precipitated with anti-SHPRH 3F8 antibody. (G) HeLa cells were starved for 5 h in HBSS. During immunoprecipitation, 100 μg/mL of ethidium bromide was added to the IP buffer to dissociate any nonspecific proteins binding through DNA. *P < 0.5; **P < 0.01; ***P < 0.001, respectively.

CHD4 Mediates the Relocalization of SHPRH at the rDNA Promoter.

CHD4/NuRD establishes a poised status of rDNA that pauses rDNA transcription by specific histone modifications. In starved cells, the amount of CHD4-occupied and poised rDNA marked by H3 K4me3 is increased and RNA polymerase I binding on the rDNA is reduced (16). Silencing of CHD4 expression decreases the level of H3 K4me3 and increases the CpG-methylation in the rDNA promoter, which results in the inactivation of rDNA (16). When CHD4 expression was silenced, targeting of SHPRH to the rDNA promoter was also diminished, which is consistent with the involvement of SHPRH at the active rDNA promoter (Fig. 7A). There was an increase of H3 K4me2 at H8 rDNA and decrease of H3 K4me2 at H42.9 rDNA when CHD4 was depleted, which would facilitate SHPRH relocalization (Fig. S9 A and B). The simultaneous knockdown of SHPRH and CHD4 did not further decrease pre-rRNA transcription (Fig. 7B), supporting the cooperation of pre-rRNA transcription by SHPRH and CHD4. To investigate whether blocking rRNA transcription alone would be enough to dissociate SHPRH from the rDNA promoter, SHPRH was monitored after cells were treated with the rRNA transcription inhibitor, actinomycin D. Although SHPRH did not significantly accumulate at the H8 region, SHPRH dissociated from rDNA promoter H42.9 by inhibition of rRNA transcription (Fig. S9C). Thus, epigenetic changes in rDNA, as well as inhibition of rRNA transcription, appear to affect the relocalization of SHPRH. Collectively, we concluded that CHD4 changes the status of the rDNA promoter to control rDNA transcription and the influences on SHPRH for rRNA transcription.

Fig. 7.
Fig. 7.

SHPRH is associated with CHD4 in rRNA transcription. (A) CHD4 controlled the targeting of SHPRH on rDNA. To deplete CHD4, HeLa cells were transfected with siRNA of CHD4 or nontargeting siRNA as control. ChIP assay was completed with anti-SHPRH 3F8 antibody, and precipitates were analyzed by qPCR in triplicate. (B) CHD4 and SHPRH cooperated in pre-rRNA transcription. CHD4 or SHPRH was depleted with siRNA in HeLa cells. The amount of pre-rRNA normalized to β-actin was measured by RT-qPCR. Data show mean ± SEM from four independent experiments.

Fig. S9.
Fig. S9.

The treatment of actinomycin D dissociated SHPRH from the rDNA promoter. (A and B) In control or CHD4-silenced HeLa cells, histone H3 methylations on rDNA H8 or H42.9 were measured by ChIP assay. (C) Doxycycline was treated to HEK293-TetOn-SHPRH-myc-His cells for 2 d, followed by 4 h treatment of 1 μM actinomycin (D) Enrichment of SHPRH-myc-His was measured by ChIP assay, using anti-6xHis antibody. Graphs show the mean ± SEM from four independent experiments.

Discussion

In this study, we present an unexpected role for SHPRH in rRNA transcription. SHPRH was enriched at the rDNA promoter in the nucleoli, where SHPRH promoted pre-rRNA transcription. Enhanced pre-rRNA transcription by SHPRH depends on the nutrient state of the cell and CHD4. SHPRH accumulates at the active rDNA promoters to assist the recruitment of RNA polymerase I for pre-rRNA transcription, whereas SHPRH is displaced from the rDNA promoter when histone H3 is trimethylated at K4 and relocalized to other rDNA sites including the H8 region of rDNA (Fig. S10).

Fig. S10.
Fig. S10.

Model for how SHPRH localizes at different locations of rDNA and functions in different media conditions. SHPRH exists in nucleoli as well as other locations of the nucleus in normal growth condition. SHPRH recruits RNA polymerase I through protein–protein interaction, an activity of RING domain, and recognition of histone codes of rDNA promoter. Changes of histone codes by starvation result in displacement of SHPRH from the rDNA promoter to other rDNA sites, including rDNA H8 and formation of SHPRH foci.

The majority of DNA damage repair proteins were identified and studied in depth after cells were challenged with DNA-damaging agents (46, 47). After DNA damage, many DNA repair proteins are redistributed in cells in response to posttranslational modifications, including phosphorylation, ubiquitylation, and acetylation, or their level is increased (48, 49). However, some repair proteins are nevertheless expressed at high levels in the absence of DNA damage, suggesting they may be required for other cellular functions in the absence of DNA damage. Indeed, the nonhomologous end joining DNA repair pathway functions during the development of lymphocytes (50) and variable, diversity and joining gene segments recombination that is required to produce antibody and T-cell receptor diversity (51). Likewise, nucleotide excision repair proteins participate in transcriptional activation in the absence of DNA damage (52). Embryonic lethality through the inactivation of many DNA repair proteins suggests those proteins are important for rapid proliferation. For example, knockout of the Nijmegen breakage syndrome gene leads to early embryonic lethality in mice (53). Although human ATAD5 regulates the level of ubiquitylated PCNA in response to DNA damage (54), ATAD5 also regulates the lifespan of DNA replication factories by modulating the level of PCNA on chromatin (55, 56). SHPRH is also expressed in most tissues regardless of exogenous DNA damage (57). Furthermore, although SHPRH and HLTF catalyze the ubiquitylation of PCNA, which is important for DNA damage bypass, disruption of SHPRH and HLTF genes in mice did not cause a significant cellular sensitivity to DNA damaging agents (29, 30). Thus, our current study suggests that the regulation of pre-rRNA transcription by SHPRH would be an alternative function of SHPRH in addition to the DNA damage response and repair. For this function of SHPRH, a ubiquitylating activity is also required, as the RING domain mutation of SHPRH no longer recruited RNA polymerase I to the rDNA promoter, although the RING domain itself was not required for interaction (Fig. 6D and Fig. S8 B and C). Thus, the regulation of pre-rRNA transcription by SHPRH requires protein–protein interaction, as well as ubiquitylation activity of SHPRH.

Histone codes, also known as epigenetic codes, have begun to emerge as an additional important signature affecting gene expression. The loci of rDNA are also modified with diverse histone methylations and acetylation (45, 58). Regulation of pre-rRNA transcription by SHPRH through the interaction between the PHD of SHPRH and the histone codes suggests rRNA expression is also tightly regulated via different histone codes and their interacting proteins. Lysine 4 trimethylation of histone H3, which is typically coded for active transcription by RNA polymerase II, is differently recognized in rRNA transcription as a poised epigenetic marker. The CHD4/NuRD is associated with such poised rRNA genes and protects the methylation-dependent inactivation of rDNA (16). The NuRD complex also has activities of ATP-dependent nucleosome disruption and histone deacetylation activities that usually result in the deactivation of transcriptional activity (59, 60). Starvation turns off rRNA transcription by dissociating SHPRH together with the RNA polymerase I complex from the rDNA promoter. Indeed, when cells are differentiated or in serum depletion, the level of H3 K4me3 in rDNA is increased (16). Although CHD4 establishes the poised rDNA, a depletion of CHD4 causes the up-regulation of H4 acetylation and rDNA methylation, which result in the down-regulation of rDNA transcription (16). Therefore, CHD4 depletion might make rDNA devoid of rRNA transcription machinery despite histone acetylation, which leads to the dissociation of SHPRH from the rDNA promoter (Fig. 7A). Moreover, in CHD4-depleted cells, the H3K4me2 was increased at a nonpromoter rDNA region, including H8, and decreased at rDNA promoter H42.9 (Fig. S9 A and B), which can drive the redistribution of SHPRH. Therefore, the role of SHPRH at the rDNA promoter for rRNA transcription can be suppressed through the establishment of constitutive heterochromatins in association with CHD4 depletion.

NuRD complexes and CHD4 are important during embryonic development, carcinogenesis, and starvation. In addition, theses complexes are recruited to damaged DNA and appear to regulate DNA damage responses (61⇓–63). On the basis of such diverse effects of the CHD4 and NuRD complexes in modulating histone signatures, it is possible that the methylation-dependent localization of SHPRH would not be restricted to rRNA gene expression. SHPRH might affect gene regulation in DNA that has differential histone signatures caused by DNA damage, gene silencing, development, or differentiation. Indeed, SHPRH is widely distributed in the nucleus, and most of the SHPRH was detected in the chromatin bound fraction (Fig. 1A and Fig. S3) and localized at the promoters of several genes, although the significance of such localization is not understood at the present time (Fig. S5 J and K).

Starvation or rapamycin treatment induced nucleolar SHPRH foci (Fig. 1 B and C). The foci represent SHPRH relocalization and accumulation outside of the rDNA promoter because SHPRH dissociates from the rDNA promoter either on starvation or on treatment of rapamycin without changing the level of SHPRH protein (Fig. 2C and Fig. S4 G and H). Depletion of SHPRH increased the frequency of MMS-induced chromosomal breaks (22, 26). In yeast, the number of rDNA repeats is regulated during aging (64). Thus, it is possible that accumulation of SHPRH on starvation could suppress genomic instability in repeated rDNA through recombination. Although it is not clear whether the number of rDNA repeats in mammals are changed during aging, it is possible that starvation-induced SHPRH foci could reflect a structure of antirecombination in the nucleoli. It is an intriguing idea, as such regulatory mechanisms could indirectly connect to the longevity regulation of the mTOR pathway (65). Alternatively, starvation-induced SHPRH foci could be a simple reservoir of unnecessary proteins before its degradation. Indeed, during starvation or autophagy, nonessential proteins lose their activity and are degraded (66). Consistent with this notion, on 24 h-starvation, the level of SHPRH protein was slightly decreased, suggesting it is being degraded.

An abnormality of ribosome biogenesis is linked with many genetic diseases, including Diamond-Blackfan anemia and 5q− syndrome (8, 67). Mutations in genes encoding ribosomal proteins would directly affect ribosomal functions in protein translation and could result in anemia. However, the causative mutations of a large portion of these anemic diseases have not been identified (68). Therefore, it is possible that mutations in genes regulating rRNA transcription or expression of ribosomal protein would be causative mutations affecting the remaining anemic patients whose mutations have not been identified. Although there was no clear anemic phenotype in Shprh−/− mice under normal growing conditions, it is possible that stress or nutrient challenge could induce an anemic condition or nonanemic diseases in Shprh−/− mice; for example, Treacher Collins syndrome. In the hematopoietic system, differential methylation on H3 lysine 4 also marks active or poised hematopoietic genes and determines the lineage of hematopoietic development (69). Thus, it is possible that SHPRH could function in hematopoietic systems as well.

Taken together, we demonstrate a regulatory function of SHPRH in 47S ribosomal RNA transcription through the specific recognition of histone codes in the rDNA promoter, with the cooperation of the chromosomal remodeler, CHD4.

Materials and Methods

Cell Culture, Reagents, Proteins, and Antibodies.

HeLa cells were maintained in DMEM supplemented with 10% FBS. To induce starvation conditions, cells were incubated in HBSS, including Ca2+ and Mg2+ (Invitrogen). The sources of antibodies and reagents are provided in SI Materials and Methods.

In Vitro Peptide-Binding Assay.

Twenty nanograms of purified GST and GST-tagged PHD of SHPRH was incubated in pulldown buffer (50 mM Tris⋅HCl at pH 7.5, 150 mM NaCl, 0.1% Tween-20, protease inhibitor mixture) for 1 h at 4 °C with rotation with 0.5 μg biotinylated synthetic H3 peptides (Upstate or Invitrogen; SI Materials and Methods) that were differentially methylated or mutated, followed by a further 30-min incubation with 10 μL Streptavidin Sepharose (GE Healthcare). Precipitates were washed 3 times with pulldown buffer and subjected to Western blot analysis.

Chromatin Immunoprecipitation.

Chromatin immunoprecipitation was performed as previously described (70). Cells were cross-linked with 1% formaldehyde at room temperature for 10 min, rinsed with PBS twice, and collected into collection buffer (100 mM Tris⋅HCl at pH 9.4, 10 mM DTT). Cells were incubated for 15 min at 30 °C and collected by centrifugation for 5 min at 2,000 × g. Pellets were washed sequentially with PBS, buffer I (10 mM Hepes at pH 6.5, 10 mM EDTA, 0.5 mM EGTA, 0.25% Triton X-100) and buffer II (10 mM Hepes at pH 6.5, 200 mM NaCl, 1 mM EDTA, 0.5 mM EGTA). Pellets were then lysed in lysis buffer (50 mM Tris⋅HCl at pH 8.1, 10 mM EDTA, 1% SDS, protease inhibitor mixture) by sonication. Cell debris was removed by centrifugation of lysates for 10 min at 13,000 × g, and supernatants were diluted in 10 volume dilution buffer (20 mM Tris⋅HCl at pH 8.1, 150 mM NaCl, 2 mM EDTA, 1% Triton X-100, protease inhibitor mixture). Cross-linked DNA and protein complexes were immunoprecipitated by incubation with 2 μg specific antibody overnight after immunoclearing with 2 μg salmon sperm DNA, 2 μg control IgG, and Protein G Sepharose for 2 h at 4 °C. Soluble chromatins were further incubated with Protein G Sepharose, 2 μg salmon sperm DNA for 1 h. Immunocomplexes were washed sequentially in TSE I (20 mM Tris⋅HCl at pH 8.1, 150 mM NaCl, 2 mM EDTA, 1% Triton X-100, 0.1% SDS), TSE II (20 mM Tris⋅HCl at pH 8.1, 500 mM NaCl, 2 mM EDTA, 1% Triton X-100, 0.1% SDS), and buffer III (10 mM Tris⋅HCl at pH 8.1, 250 mM LiCl, 1 mM EDTA, 1% Nonidet P-40, 1% deoxycholate), and three times in TE buffer. DNA–protein complexes were eluted two times with elution buffer (0.1 M NaHCO3, 1% SDS) and reverse-cross-linked by incubating at 65 °C for 6 h. DNA fragments were purified by ethanol precipitation and analyzed by quantitative PCR, using Platinum SYBR Green qPCR SuperMix-UDG with ROX (Invitrogen). Primer sequences for qPCR are referred from ref. 71.

Nascent rRNA Staining.

Staining of Nascent rRNA was performed as described previously (44). HeLa cells were washed with PBS and with permeabilization buffer (20 mM Tris⋅HCl at pH 7.5, 5 mM MgCl2, 0.5 mM EGTA, 0.5 mM PMSF). Cells were treated with permeabilization buffer containing 0.5% Triton X-100 for 5 min and washed with permeabilization buffer. Synthesis of rRNA was achieved by incubation of permeabilized cells in synthesis buffer (50 mM Tris⋅HCl at pH 7.5, 100 mM KCl, 5 mM MgCl2, 0.5 mM EGTA, 25 U/mL RNasin, 1 mM PMSF, 0.5 mM ATP, CTP and GTP, 0.2 mM Br-UTP) for 30 min at 37 °C. The cells were then washed three times with PBS containing 25 U/mL RNasin and fixed with 10% formaldehyde in PBS containing 25 U/mL RNasin and 0.1% BSA for 20 min. After washing twice with PBS containing 0.1% BSA for 5 min each, nascent rRNA in the cells were stained with anti-BrdU (BU1/75) antibody (Abcam) and Alexa488-conjugated anti-rat secondary antibody. Images were analyzed using ImageJ (NIH).

Other materials and methods are provided in SI Materials and Methods.

SI Materials and Methods

Reagents and Antibodies.

Rapamycin and doxycycline were purchased from Sigma and Clontech, respectively.

Anti-SHPRH (3F8) monoclonal antibody was obtained from OriGene Technologies. Anti-RPA194 (Santa Cruz), anti-UBF (Santa Cruz), anti-Fibrillarin (Cell Signaling), anti-myc 9E10 (Sigma), anti-6xHis (Abcam), anti-Histone H3 (Millipore), anti-GST (Amersham Bioscience), anti-H3 K4me2 and anti-H3 K4me3 (Abcam), anti-phospho-RPA32 S4/S8, anti-phospho-RPA32 S33 (Bethyl Laboratory), anti-γH2AX (Upstate), anti-pol η (Abcam), anti-pol κ (Abcam), anti-PCNA (SantaCruz), anti-phospho-CHK1 (Cell Signaling), anti-53BP1 (Abcam), anti-XPA, and anti-BRCA1 (SantaCruz) were purchased and used as recommended by the manufacturer for immunofluorescence, immunoprecipitation, and Western blot assay.

Plasmids, siRNAs, and Transfections.

Plasmid DNAs for SHPRH PHD mutants (E660A, E660Y) were generated by site-directed mutagenesis (Stratagene) with a wild-type full-length SHPRH-expressing plasmid (pcDNA3.1/SHPRH-myc-His, PKJM704) as a template (22), according to the manufacturer’s instructions. Primer sequences for mutagenesis are 5′-AACACCTCTGATTACCGCTTTGCATGTATATGTGGTGAACTTGAT-3′ (E660A sense; PRKJM2931), 5′-ATCAAGTTCACCACATATACATGCAAAGCGGTAATCAGAGGTGTT-3′ (E660A antisense; PRKJM2932), 5′-AACACCTCTGATTACCGCTTTTATTGTATATGTGGTGAACTTGAT-3′ (E660Y sense; PRKJM2933) and 5′-ATCAAGTTCACCACATATACAATAAAAGCGGTAATCAGAGGTGTT-3′ (E660Y antisense; PRKJM2934). The sequences resulting in the E660A or E660Y substitutions incorporated during site directed mutagenesis are italicized.

All siRNAs were purchased or synthesized from GE Healthcare Dharmacon Inc. siRNA transfection was performed using RNAiMAX (Invitrogen). ON-TARGETplus nontargeting pool and ON-TARGETplus SMART pool siRNAs for SHPRH and CHD4 were purchased. To target the 3′ UTR of SHPRH, siRNAs were synthesized from GE Healthcare Dharmacon Inc.: GAGCUAUGUUUUAGAGAAAUU (#1 sense), UUUCUCUAAAACAUAGCUCUU (#1 antisense), GUAAAGUGGUGUUAGUGUAUU (#3 sense) and UACACUAACACCACUUUACUU (#3 antisense).

Plasmid DNAs were transfected using Attractene (Qiagen) or Lipofectamine 2000 (Invitrogen) to HeLa or 293 cells, respectively, according to manufacturer’s instructions, and transfected cells were incubated for 24 h before further analysis. Transfection of siRNA was performed using RNAiMAX (Invitrogen), and cells were incubated for 48 h before further analysis.

Purification of Full-Length SHPRH and GST-PHD, and in Vitro Binding Assay.

Full-length SHPRH-myc-His was expressed in Sf21 cells and purified through an immobilized metal ion affinity chromatography by scientists at ProteinOne LLC. To investigate the interaction between SHPRH and histones, 2 μg purified SHPRH-myc-His was incubated with 1–10 µg calf thymus total histones (Worthington Biochemical Co.) in binding buffer [50 mM Tris⋅HCl at pH 7.5, 1 M NaCl, 1% Nonidet P-40, 0.5 mM EDTA, 1 mM PMSF plus protease inhibitor mixture (Roche)] at 4 °C for 4 h, followed by an additional 1 h incubation with Ni-NTA Sepharose bead (Thermo Fisher Scientific) and extensive washing.

The PHD of SHPRH (amino acids 656–712) was subcloned in pGEX-6P-1 vector (GE Healthcare; named pKJM1345), and GST-fused PHD, as well as GST, were purified according to the manufacturer’s instructions, using Glutathione Sepharose 4B (GE Healthcare). Wild-type and methylated histone H3 peptides were purchased from Upstate, and mutant H3 peptides were synthesized by Invitrogen. The sequence of histone H3 peptides used in pull-down experiments was ARTKQTARKSTGGKAPRKQLA-GGK-biotin, and each lysine 4 (K4) or K9 is methylated for modifications or substituted with alanine.

Immunocytochemistry, Nascent rRNA Staining, and Microscopy.

Cells were fixed with 4% paraformaldehyde in PBS for 10 min and washed 3 times with PBS. Cells were permeabilized by treatment with 0.5% Triton X-100 in PBS for 10 min and further incubated in PBS supplemented with 10% donkey serum and 3% BSA for 1 h. Primary antibodies were incubated for 1 h and washed 3 times with 0.05% Triton X-100 in PBS. Secondary antibodies conjugated with Alexa Fluor 488 or 568 (Invitrogen) were incubated for 50 min and washed 3 times. Cells were mounted using Prolong Gold Antifade reagent supplemented with DAPI (Invitrogen). Images were taken with Axio Imager.D2, Axio Observer.D1, or LSM780NLO (Zeiss)

Immunoprecipitation.

Immunoprecipitation was performed as described (28). Cells were washed twice with PBS and harvested with 3× IP buffer (10 mM Tris⋅HCl at pH 7.5, 400 mM NaCl, 1 mM EDTA, 15% glycerol, 0.5% Nonidet P-40, protease inhibitor mixture). After sonication, lysates were diluted with two volumes of dilution buffer (10 mM Tris⋅HCl at pH 7.5, 1 mM EDTA, 15% glycerol, 0.5% Nonidet P-40, protease inhibitor mixture) and clarified by centrifugation. Nonspecific binding proteins to Protein G Sepharose were precleared by incubating cell lysates with Protein G Sepharose for 2 h. After removing nonspecific binding proteins attached in Protein G Sepharose, the precleared lysates were incubated with primary antibody for 2 h, followed by incubation with Protein G Sepharose for 1 h. Precipitated proteins were analyzed by Western blotting. To eliminate DNA-dependent interactions, 100 μg/mL EtBr was added into the IP buffer and maintained during the washing step (72).

Preparation of RNA and Measurement of RNA Level.

Total cellular RNA was prepared by using RNeasy Plus Kit (Qiagen), according to the manufacturer’s protocol. One microgram total RNA was used for reverse transcription by using SuperScript III platinum kit (Invitrogen) or QuantiTect Reverse Transcription Kit (Qiagen). The RNA level was measured using Platinum SYBR Green qPCR SuperMix-UDG with ROX (Invitrogen). PCR primers used for RT-PCR were 5′-GCTCGTCGTCGACAACGGCTC-3′, 5′-CAAACATGATCTGGGTCATCTTCTC-3′ (human β-actin; PRKJM1076 and PRKJM1077), 5′-TGGTGGAGTCATCCTTCAGTGGT-3′, 5′- AGTTTTGCTAGGCCAGCTTCCAAA-3′ (human SHPRH; PRKJM2867 and PRKJM2868). For the human pre-rRNA, either of two primer sets was used: 5′-CCTGCTGTTCTCTCGCGCGTCCGAG-3′ and 5′-AACGCCTGACACGCACGGCACGGAG-3′ [human pre-rRNA (71)] or 5′-GAACGG TGGTGTGTCGTTC-3′ and 5′-GCGTCTCGTCTCGTCTCACT-3′ (73).

Primer lists for amplifications of nonrDNA loci.

View this table:

Primer lists for RT-qPCR of nonrDNA loci.

View this table:

Cell Fractionation.

Total cell extracts were prepared by resuspending cells in 1× Laemmli sample buffer, followed by boiling for 10 min. The chromatin-bound fraction was prepared as described previously (74). Briefly, cells were washed with PBS, resuspended in buffer A (10 mM Hepes at pH 7.9, 10 mM KCl, 1.5 mM MgCl2, 0.34 M sucrose, 10% glycerol, 1 mM DTT, 10 mM NaF, 0.1% Triton X-100, and protease inhibitor mixture) and incubated on ice for 5 min. The cytoplasmic fraction was separated by centrifugation (1,300 × g, 5 min, 4 °C). The nuclear fraction was washed with buffer A, resuspended in buffer B (3 mM EDTA, 0.2 mM EGTA, 1 mM DTT, and protease inhibitor mixture) and incubated on ice for 10 min. The chromatin-bound fraction was separated from the soluble nuclear fraction by centrifugation (1,700 × g, 5 min, 4 °C).

Acknowledgments

We thank Dr. D. Bodine (National Human Genome Research Institute), Dr. J. Fox (National Center for Advancing Translational Sciences), and members of the K.M. laboratory for helpful discussions and comments on the manuscript; K.M. especially thanks E. Cho. This research was supported by the intramural Research Programs of the National Human Genome Research Institute (HG200376-01) (to K.M.) and of the National Institute of Child Health and Human Development (R.W.). This work was also supported by the Institute for Basic Science (IBS-R022-D1-2015) (to K.M.). D.L. was supported in part by the National Research Foundation of Korea Ministry of Science, ICT, and Future Planning (MSIP) Grant 2010-0028684.

Footnotes

  • ↵1Present address: Medytox Inc., Bundang-gu, Seongnam-si, Gyeonggi-do, 463-400, Korea.

  • ↵2Present address: Department of Biological Sciences, Chonbuk National University, Jeonju, Chonbuk, 561-756, Korea.

  • ↵3Present address: Department of Life Sciences, National Cheng Kung University, Taiwan City, Taiwan 70101.

  • ↵4To whom correspondence should be addressed. Email: kmyung{at}ibs.re.kr.

  • Author contributions: D.L. and K.M. designed research; D.L., J.A., Y.U.P., H.L., and J.H.P. performed research; R.W. and K.M. contributed new reagents/analytic tools; D.L., J.A., Y.U.P., H.L., J.H.P., and K.M. analyzed data; D.L., R.W., and K.M. wrote the paper.

  • The authors declare no conflict of interest.

  • This article is a PNAS Direct Submission. K.C. is a Guest Editor invited by the Editorial Board.

  • This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1701978114/-/DCSupplemental.

References

  1. ↵
    1. Gonzalez IL,
    2. Sylvester JE
    (1995) Complete sequence of the 43-kb human ribosomal DNA repeat: Analysis of the intergenic spacer. Genomics 27:320–328.
    .
    OpenUrlCrossRefPubMed
  2. ↵
    1. Boisvert FM,
    2. van Koningsbruggen S,
    3. Navascués J,
    4. Lamond AI
    (2007) The multifunctional nucleolus. Nat Rev Mol Cell Biol 8:574–585.
    .
    OpenUrlCrossRefPubMed
  3. ↵
    1. Grummt I,
    2. Pikaard CS
    (2003) Epigenetic silencing of RNA polymerase I transcription. Nat Rev Mol Cell Biol 4:641–649.
    .
    OpenUrlCrossRefPubMed
  4. ↵
    1. Granneman S,
    2. Baserga SJ
    (2005) Crosstalk in gene expression: Coupling and co-regulation of rDNA transcription, pre-ribosome assembly and pre-rRNA processing. Curr Opin Cell Biol 17:281–286.
    .
    OpenUrlCrossRefPubMed
  5. ↵
    1. Beau I,
    2. Esclatine A,
    3. Codogno P
    (2008) Lost to translation: When autophagy targets mature ribosomes. Trends Cell Biol 18:311–314.
    .
    OpenUrlCrossRefPubMed
  6. ↵
    1. Chen FW,
    2. Ioannou YA
    (1999) Ribosomal proteins in cell proliferation and apoptosis. Int Rev Immunol 18:429–448.
    .
    OpenUrlCrossRefPubMed
  7. ↵
    1. Volarevic S, et al.
    (2000) Proliferation, but not growth, blocked by conditional deletion of 40S ribosomal protein S6. Science 288:2045–2047.
    .
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Burwick N,
    2. Coats SA,
    3. Nakamura T,
    4. Shimamura A
    (2012) Impaired ribosomal subunit association in Shwachman-Diamond syndrome. Blood 120:5143–5152.
    .
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Hannan KM,
    2. Sanij E,
    3. Rothblum LI,
    4. Hannan RD,
    5. Pearson RB
    (2013) Dysregulation of RNA polymerase I transcription during disease. Biochim Biophys Acta 1829:342–360.
    .
    OpenUrlCrossRefPubMed
  10. ↵
    1. Liu JM,
    2. Ellis SR
    (2006) Ribosomes and marrow failure: Coincidental association or molecular paradigm? Blood 107:4583–4588.
    .
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Sonenberg N,
    2. Hinnebusch AG
    (2009) Regulation of translation initiation in eukaryotes: Mechanisms and biological targets. Cell 136:731–745.
    .
    OpenUrlCrossRefPubMed
  12. ↵
    1. van Riggelen J,
    2. Yetil A,
    3. Felsher DW
    (2010) MYC as a regulator of ribosome biogenesis and protein synthesis. Nat Rev Cancer 10:301–309.
    .
    OpenUrlCrossRefPubMed
  13. ↵
    1. Drygin D,
    2. Rice WG,
    3. Grummt I
    (2010) The RNA polymerase I transcription machinery: An emerging target for the treatment of cancer. Annu Rev Pharmacol Toxicol 50:131–156.
    .
    OpenUrlCrossRefPubMed
  14. ↵
    1. Murayama A, et al.
    (2008) Epigenetic control of rDNA loci in response to intracellular energy status. Cell 133:627–639.
    .
    OpenUrlCrossRefPubMed
  15. ↵
    1. Santoro R,
    2. Li J,
    3. Grummt I
    (2002) The nucleolar remodeling complex NoRC mediates heterochromatin formation and silencing of ribosomal gene transcription. Nat Genet 32:393–396.
    .
    OpenUrlCrossRefPubMed
  16. ↵
    1. Xie W, et al.
    (2012) The chromatin remodeling complex NuRD establishes the poised state of rRNA genes characterized by bivalent histone modifications and altered nucleosome positions. Proc Natl Acad Sci USA 109:8161–8166.
    .
    OpenUrlAbstract/FREE Full Text
  17. ↵
    1. Mayer C,
    2. Grummt I
    (2006) Ribosome biogenesis and cell growth: mTOR coordinates transcription by all three classes of nuclear RNA polymerases. Oncogene 25:6384–6391.
    .
    OpenUrlCrossRefPubMed
  18. ↵
    1. Sarbassov DD,
    2. Ali SM,
    3. Sabatini DM
    (2005) Growing roles for the mTOR pathway. Curr Opin Cell Biol 17:596–603.
    .
    OpenUrlCrossRefPubMed
  19. ↵
    1. Iadevaia V,
    2. Zhang Z,
    3. Jan E,
    4. Proud CG
    (2012) mTOR signaling regulates the processing of pre-rRNA in human cells. Nucleic Acids Res 40:2527–2539.
    .
    OpenUrlAbstract/FREE Full Text
  20. ↵
    1. Tsang CK,
    2. Liu H,
    3. Zheng XF
    (2010) mTOR binds to the promoters of RNA polymerase I- and III-transcribed genes. Cell Cycle 9:953–957.
    .
    OpenUrlCrossRefPubMed
  21. ↵
    1. Vazquez-Martin A,
    2. Cufí S,
    3. Oliveras-Ferraros C,
    4. Menendez JA
    (2011) Raptor, a positive regulatory subunit of mTOR complex 1, is a novel phosphoprotein of the rDNA transcription machinery in nucleoli and chromosomal nucleolus organizer regions (NORs). Cell Cycle 10:3140–3152.
    .
    OpenUrlCrossRefPubMed
  22. ↵
    1. Motegi A, et al.
    (2006) Human SHPRH suppresses genomic instability through proliferating cell nuclear antigen polyubiquitination. J Cell Biol 175:703–708.
    .
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Unk I, et al.
    (2006) Human SHPRH is a ubiquitin ligase for Mms2-Ubc13-dependent polyubiquitylation of proliferating cell nuclear antigen. Proc Natl Acad Sci USA 103:18107–18112.
    .
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Fox JT,
    2. Lee KY,
    3. Myung K
    (2011) Dynamic regulation of PCNA ubiquitylation/deubiquitylation. FEBS Lett 585:2780–2785.
    .
    OpenUrlCrossRefPubMed
  25. ↵
    1. Unk I,
    2. Hajdú I,
    3. Blastyák A,
    4. Haracska L
    (2010) Role of yeast Rad5 and its human orthologs, HLTF and SHPRH in DNA damage tolerance. DNA Repair (Amst) 9:257–267.
    .
    OpenUrlCrossRefPubMed
  26. ↵
    1. Lin JR,
    2. Zeman MK,
    3. Chen JY,
    4. Yee MC,
    5. Cimprich KA
    (2011) SHPRH and HLTF act in a damage-specific manner to coordinate different forms of postreplication repair and prevent mutagenesis. Mol Cell 42:237–249.
    .
    OpenUrlCrossRefPubMed
  27. ↵
    1. Motegi A, et al.
    (2008) Polyubiquitination of proliferating cell nuclear antigen by HLTF and SHPRH prevents genomic instability from stalled replication forks. Proc Natl Acad Sci USA 105:12411–12416.
    .
    OpenUrlAbstract/FREE Full Text
  28. ↵
    1. Unk I, et al.
    (2008) Human HLTF functions as a ubiquitin ligase for proliferating cell nuclear antigen polyubiquitination. Proc Natl Acad Sci USA 105:3768–3773.
    .
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Hendel A, et al.
    (2011) PCNA ubiquitination is important, but not essential for translesion DNA synthesis in mammalian cells. PLoS Genet 7:e1002262.
    .
    OpenUrlCrossRefPubMed
  30. ↵
    1. Krijger PH, et al.
    (2011) HLTF and SHPRH are not essential for PCNA polyubiquitination, survival and somatic hypermutation: Existence of an alternative E3 ligase. DNA Repair (Amst) 10:438–444.
    .
    OpenUrlCrossRefPubMed
  31. ↵
    1. Tomi NS, et al.
    (2014) Analysis of SHPRH functions in DNA repair and immunoglobulin diversification. DNA Repair (Amst) 24:63–72.
    .
    OpenUrl
  32. ↵
    1. Horáková AH, et al.
    (2010) SUV39h-independent association of HP1 beta with fibrillarin-positive nucleolar regions. Chromosoma 119:227–241.
    .
    OpenUrlCrossRefPubMed
  33. ↵
    1. Proud CG
    (2002) Regulation of mammalian translation factors by nutrients. Eur J Biochem 269:5338–5349.
    .
    OpenUrlPubMed
  34. ↵
    1. Scheer U,
    2. Hock R
    (1999) Structure and function of the nucleolus. Curr Opin Cell Biol 11:385–390.
    .
    OpenUrlCrossRefPubMed
  35. ↵
    1. Warner JR
    (1990) The nucleolus and ribosome formation. Curr Opin Cell Biol 2:521–527.
    .
    OpenUrlCrossRefPubMed
  36. ↵
    1. O’Sullivan AC,
    2. Sullivan GJ,
    3. McStay B
    (2002) UBF binding in vivo is not restricted to regulatory sequences within the vertebrate ribosomal DNA repeat. Mol Cell Biol 22:657–668.
    .
    OpenUrlAbstract/FREE Full Text
  37. ↵
    1. Baker LA,
    2. Allis CD,
    3. Wang GG
    (2008) PHD fingers in human diseases: Disorders arising from misinterpreting epigenetic marks. Mutat Res 647:3–12.
    .
    OpenUrlCrossRefPubMed
  38. ↵
    1. Musselman CA,
    2. Kutateladze TG
    (2009) PHD fingers: Epigenetic effectors and potential drug targets. Mol Interv 9:314–323.
    .
    OpenUrlCrossRefPubMed
  39. ↵
    1. Sanchez R,
    2. Zhou MM
    (2011) The PHD finger: A versatile epigenome reader. Trends Biochem Sci 36:364–372.
    .
    OpenUrlPubMed
  40. ↵
    1. Lan F, et al.
    (2007) Recognition of unmethylated histone H3 lysine 4 links BHC80 to LSD1-mediated gene repression. Nature 448:718–722.
    .
    OpenUrlCrossRefPubMed
  41. ↵
    1. Mansfield RE, et al.
    (2011) Plant homeodomain (PHD) fingers of CHD4 are histone H3-binding modules with preference for unmodified H3K4 and methylated H3K9. J Biol Chem 286:11779–11791.
    .
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Hamperl S, et al.
    (2013) Chromatin states at ribosomal DNA loci. Biochim Biophys Acta 1829:405–417.
    .
    OpenUrlCrossRef
  43. ↵
    1. Zentner GE,
    2. Balow SA,
    3. Scacheri PC
    (2014) Genomic characterization of the mouse ribosomal DNA locus. G3 (Bethesda) 4:243–254.
    .
    OpenUrl
  44. ↵
    1. Ko YG,
    2. Kang YS,
    3. Kim EK,
    4. Park SG,
    5. Kim S
    (2000) Nucleolar localization of human methionyl-tRNA synthetase and its role in ribosomal RNA synthesis. J Cell Biol 149:567–574.
    .
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Zentner GE,
    2. Saiakhova A,
    3. Manaenkov P,
    4. Adams MD,
    5. Scacheri PC
    (2011) Integrative genomic analysis of human ribosomal DNA. Nucleic Acids Res 39:4949–4960.
    .
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Khapre RV,
    2. Samsa WE,
    3. Kondratov RV
    (2010) Circadian regulation of cell cycle: Molecular connections between aging and the circadian clock. Ann Med 42:404–415.
    .
    OpenUrlCrossRefPubMed
  47. ↵
    1. You Z,
    2. Bailis JM
    (2010) DNA damage and decisions: CtIP coordinates DNA repair and cell cycle checkpoints. Trends Cell Biol 20:402–409.
    .
    OpenUrlCrossRefPubMed
  48. ↵
    1. Jackson SP,
    2. Durocher D
    (2013) Regulation of DNA damage responses by ubiquitin and SUMO. Mol Cell 49:795–807.
    .
    OpenUrlCrossRefPubMed
  49. ↵
    1. Polo SE,
    2. Jackson SP
    (2011) Dynamics of DNA damage response proteins at DNA breaks: A focus on protein modifications. Genes Dev 25:409–433.
    .
    OpenUrlAbstract/FREE Full Text
  50. ↵
    1. Gao Y, et al.
    (1998) A critical role for DNA end-joining proteins in both lymphogenesis and neurogenesis. Cell 95:891–902.
    .
    OpenUrlCrossRefPubMed
  51. ↵
    1. Soulas-Sprauel P, et al.
    (2007) V(D)J and immunoglobulin class switch recombinations: A paradigm to study the regulation of DNA end-joining. Oncogene 26:7780–7791.
    .
    OpenUrlCrossRefPubMed
  52. ↵
    1. Le May N, et al.
    (2010) NER factors are recruited to active promoters and facilitate chromatin modification for transcription in the absence of exogenous genotoxic attack. Mol Cell 38:54–66.
    .
    OpenUrlCrossRefPubMed
  53. ↵
    1. Zhu J,
    2. Petersen S,
    3. Tessarollo L,
    4. Nussenzweig A
    (2001) Targeted disruption of the Nijmegen breakage syndrome gene NBS1 leads to early embryonic lethality in mice. Curr Biol 11:105–109.
    .
    OpenUrlCrossRefPubMed
  54. ↵
    1. Lee KY, et al.
    (2010) Human ELG1 regulates the level of ubiquitinated proliferating cell nuclear antigen (PCNA) through Its interactions with PCNA and USP1. J Biol Chem 285:10362–10369.
    .
    OpenUrlAbstract/FREE Full Text
  55. ↵
    1. Lee KY,
    2. Fu H,
    3. Aladjem MI,
    4. Myung K
    (2013) ATAD5 regulates the lifespan of DNA replication factories by modulating PCNA level on the chromatin. J Cell Biol 200:31–44.
    .
    OpenUrlAbstract/FREE Full Text
  56. ↵
    1. Sikdar N, et al.
    (2009) DNA damage responses by human ELG1 in S phase are important to maintain genomic integrity. Cell Cycle 8:3199–3207.
    .
    OpenUrlCrossRefPubMed
  57. ↵
    1. Sood R, et al.
    (2003) Cloning and characterization of a novel gene, SHPRH, encoding a conserved putative protein with SNF2/helicase and PHD-finger domains from the 6q24 region. Genomics 82:153–161.
    .
    OpenUrlCrossRefPubMed
  58. ↵
    1. Ito T
    (2007) Role of histone modification in chromatin dynamics. J Biochem 141:609–614.
    .
    OpenUrlAbstract/FREE Full Text
  59. ↵
    1. Becker PB,
    2. Hörz W
    (2002) ATP-dependent nucleosome remodeling. Annu Rev Biochem 71:247–273.
    .
    OpenUrlCrossRefPubMed
  60. ↵
    1. Xue Y, et al.
    (1998) NURD, a novel complex with both ATP-dependent chromatin-remodeling and histone deacetylase activities. Mol Cell 2:851–861.
    .
    OpenUrlCrossRefPubMed
  61. ↵
    1. Ho L,
    2. Crabtree GR
    (2010) Chromatin remodelling during development. Nature 463:474–484.
    .
    OpenUrlCrossRefPubMed
  62. ↵
    1. Polo SE,
    2. Kaidi A,
    3. Baskcomb L,
    4. Galanty Y,
    5. Jackson SP
    (2010) Regulation of DNA-damage responses and cell-cycle progression by the chromatin remodelling factor CHD4. EMBO J 29:3130–3139.
    .
    OpenUrlAbstract/FREE Full Text
  63. ↵
    1. Smeenk G, et al.
    (2010) The NuRD chromatin-remodeling complex regulates signaling and repair of DNA damage. J Cell Biol 190:741–749.
    .
    OpenUrlAbstract/FREE Full Text
  64. ↵
    1. Ganley AR,
    2. Ide S,
    3. Saka K,
    4. Kobayashi T
    (2009) The effect of replication initiation on gene amplification in the rDNA and its relationship to aging. Mol Cell 35:683–693.
    .
    OpenUrlCrossRefPubMed
  65. ↵
    1. Harrison DE, et al.
    (2009) Rapamycin fed late in life extends lifespan in genetically heterogeneous mice. Nature 460:392–395.
    .
    OpenUrlCrossRefPubMed
  66. ↵
    1. Klionsky DJ,
    2. Emr SD
    (2000) Autophagy as a regulated pathway of cellular degradation. Science 290:1717–1721.
    .
    OpenUrlAbstract/FREE Full Text
  67. ↵
    1. Ebert BL, et al.
    (2008) Identification of RPS14 as a 5q- syndrome gene by RNA interference screen. Nature 451:335–339.
    .
    OpenUrlCrossRefPubMed
  68. ↵
    1. Gazda HT, et al.
    (2008) Ribosomal protein L5 and L11 mutations are associated with cleft palate and abnormal thumbs in Diamond-Blackfan anemia patients. Am J Hum Genet 83:769–780.
    .
    OpenUrlCrossRefPubMed
  69. ↵
    1. Orford K, et al.
    (2008) Differential H3K4 methylation identifies developmentally poised hematopoietic genes. Dev Cell 14:798–809.
    .
    OpenUrlCrossRefPubMed
  70. ↵
    1. Shang Y,
    2. Hu X,
    3. DiRenzo J,
    4. Lazar MA,
    5. Brown M
    (2000) Cofactor dynamics and sufficiency in estrogen receptor-regulated transcription. Cell 103:843–852.
    .
    OpenUrlCrossRefPubMed
  71. ↵
    1. Grandori C, et al.
    (2005) c-Myc binds to human ribosomal DNA and stimulates transcription of rRNA genes by RNA polymerase I. Nat Cell Biol 7:311–318.
    .
    OpenUrlCrossRefPubMed
  72. ↵
    1. Lai JS,
    2. Herr W
    (1992) Ethidium bromide provides a simple tool for identifying genuine DNA-independent protein associations. Proc Natl Acad Sci USA 89:6958–6962.
    .
    OpenUrlAbstract/FREE Full Text
  73. ↵
    1. Zhai N, et al.
    (2012) Human PIH1 associates with histone H4 to mediate the glucose-dependent enhancement of pre-rRNA synthesis. J Mol Cell Biol 4:231–241.
    .
    OpenUrlAbstract/FREE Full Text
  74. ↵
    1. Kim JE,
    2. McAvoy SA,
    3. Smith DI,
    4. Chen J
    (2005) Human TopBP1 ensures genome integrity during normal S phase. Mol Cell Biol 25:10907–10915.
    .
    OpenUrlAbstract/FREE Full Text
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rRNA transcription by SHPRH
Deokjae Lee, Jungeun An, Young-Un Park, Hungjiun Liaw, Roger Woodgate, Jun Hong Park, Kyungjae Myung
Proceedings of the National Academy of Sciences Apr 2017, 114 (17) E3424-E3433; DOI: 10.1073/pnas.1701978114
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