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. Author manuscript; available in PMC: 2013 Sep 1.
Published in final edited form as: Cancer Discov. 2012 Jun 29;2(9):812–825. doi: 10.1158/2159-8290.CD-12-0116

DDX5 Regulates DNA Replication And Is Required For Cell Proliferation In A Subset Of Breast Cancer Cells

Anthony Mazurek 1, Weijun Luo 1,*, Alexander Krasnitz 1, James Hicks 1, R Scott Powers 1, Bruce Stillman 1
PMCID: PMC3440546  NIHMSID: NIHMS399630  PMID: 22750847

Abstract

Understanding factors required for DNA replication will enrich our knowledge of this important process and potentially identify vulnerabilities that can be exploited in cancer therapy. We applied an assay that measures the stability of maintenance of an episomal plasmid in human tissue culture cells to screen for new DNA replication factors. We identify an important role for DDX5 in G1-to-S phase progression where it directly regulates DNA replication factor expression by promoting the recruitment of RNA Polymerase II to E2F-regulated gene promoters. We find that the DDX5 locus is frequently amplified in breast cancer and that breast cancer derived cells with amplification of DDX5 are much more sensitive to its depletion than breast cancer cells and a breast epithelial cell line that lack DDX5 amplification. Our results demonstrate a novel role for DDX5 in cancer cell proliferation and suggest DDX5 as a therapeutic target in breast cancer treatment.

Keywords: DDX5, DNA replication, transcription, breast cancer, ERBB2

INTRODUCTION

Defects in the control of cell proliferation are a hallmark of cancer and DNA replication is a key process for cell proliferation. Understanding how DNA replication is regulated in human cells can provide insight into cancer development and may reveal vulnerabilities that can be exploited therapeutically. Indeed, a number of agents currently used in cancer treatment are known to target DNA synthesis. We applied an assay that is loosely based upon the minichromosome maintenance screen performed in budding yeast (1) in which we measured the stability of maintenance of an episomal plasmid in human tissue culture cells to screen for new factors required for DNA replication. The plasmid encodes both the EBNA1 gene and OriP, a cis-acting element derived from the Epstein-Barr virus (EBV) genome that functions as an origin of DNA replication in plasmids and enables them to tether to chromosomes during mitosis thereby allowing the plasmids to replicate and segregate to daughter cells without the need to integrate into host chromosomes (2). To be maintained in cells OriP plasmids require both the EBNA1 protein, which is encoded by a gene also derived from the EBV genome, and host cell DNA replication factors (3, 4). Importantly, OriP plasmids replicate once-per-S phase and are thus licensed to replicate similar to human chromosomes (5). Our rationale for using this episomal plasmid for the screen is that stable maintenance of a plasmid whose duplication in cells is dependent upon a single origin of replication should be more sensitive to reduced expression of DNA replication factors than chromosomes that encode many origins of replication. We identified 6 genes that had no appreciated role in DNA replication but whose expression is necessary for stable plasmid maintenance in cells and present evidence that one of these, the DEAD-box protein DDX5, regulates the expression of DNA replication genes and is required for cancer cell proliferation.

DDX5 is an ATP dependent RNA helicase that was first identified as a protein that cross-reacted with an antibody against SV40 large T-antigen (6, 7). It exhibits considerable sequence identity in its helicase core with the DEAD-box protein DDX17 and these two proteins interact in cells (8). DDX5, and in fewer cases DDX17, can function as transcriptional co-regulators with estrogen receptor alpha, p53, MyoD, and Runx2 (9). A role for DDX5 and DDX17 in miRNA maturation, ribosome biogenesis, mRNA splicing, and insulator function have also been described (10-16). DDX5 is frequently over-expressed in colon, prostate, and breast cancer (17-20) and RNAi of both DDX5 and DDX17 together impairs cancer cell proliferation (13, 19). However, the activity of DDX5 in promoting cell proliferation is poorly understood.

We find that in certain cancer cells DDX5 is required for plasmid stability and cell proliferation where it promotes G1-to-S phase progression and the expression of essential DNA replication genes. DDX5 localizes to the E2F-regulated promoters of genes encoding DNA replication factors and is required for RNA Polymerase II loading. Consistent with its positive regulation of genes required for cell division and the high frequency of DDX5 overexpression previously reported in breast cancer we find that the DDX5 locus is frequently amplified in breast cancer and is often co-amplified along with ERBB2. Surprisingly, breast cancer cell lines with amplification of the DDX5 locus are considerably more sensitive to its knockdown than breast cancer cell lines and a normal breast epithelial cell line lacking this amplification. Thus the dependence of breast cancer cells on DDX5 expression varies and is correlated with DDX5 gene copy number. These results show that breast cancer cells acquire dependence upon DDX5 suggesting a vulnerability that may be targeted by therapy.

RESULTS

Plasmid stability assay to identify genes required for DNA replication in human cells

An assay that measures stability of maintenance of an episomal plasmid in human tissue culture cells was applied to the human colorectal tumor cell line HCT116 to identify new human DNA replication factors (Figure 1A, see Materials and Methods). Cells were infected with retrovirus encoding puromycin resistance and either an shRNA targeting expression of a gene of interest or a control shRNA targeting expression of Ff-luciferase. Infected cells were selected with puromycin then transfected with the OriP plasmid encoding hygromycin resistance. Cells were then seeded at a clonogenic density in growth media containing either puromycin alone (proliferation assay) or puromycin and hygromycin (plasmid stability assay) and allowed time to expand into colonies after which the colonies were stained with crystal violet. We developed a plasmid stability ratio score to measure the ability of shRNAs targeting genes of interest to selectively affect plasmid stability as opposed to a non-selective effect on proliferation (see Materials and Methods). A ratio score of less than one indicates that the experimental shRNA impairs plasmid stability stronger than cell proliferation.

Figure 1.

Figure 1

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Plasmid maintenance screen. (A) Schematic of plasmid stability assay used for screen. (B) Quantitative western blot analysis of MCM3 and PCNA (loading control) in whole cell extracts obtained from cells transduced with the indicated shRNAs (V2HS_262054 is an shRNA that targets MCM3 expression). The Luciferasemi203 whole cell extract is loaded such that there is either equal total protein loaded as the V2HS_262054 whole cell extracts (lane indicated as “1.00” on the blot) or 25% or 10% total protein loaded as the V2HS_262054 whole cell extracts (lanes indicated as “0.25” and “0.10” respectively). (C) Colony formation assay results for triplicate HCT116 cultures transduced with either V2HS_262054, EBNA1mi1666, or Luciferasemi203 shRNAs that were grown in either growth media + puromycin (proliferation assay) or growth media + puromycin and hygromycin (plasmid stability assay). (D) Plot of plasmid stability ratios calculated from results shown in (C). (E) Plasmid stability ratios for shRNAs tested in screen. Blue bars on plot correspond to plasmid stability ratios for HCT116 cultures transduced with shRNAs that target expression of known replication factors.

We tested the hypothesis that an episomal plasmid containing a single origin-of-replication would be more sensitive to reduced expression of an essential replication factor than multi-origin chromosomes by transducing cells with an shRNA targeting the expression of MCM3, a subunit of the MCM complex that is required for DNA replication. MCM complexes are known to be loaded onto chromatin in excess of the amount necessary for chromosome duplication (21-23) and proliferation of human cancer cell lines is relatively resistant to knockdown of MCM3 in the absence of replication inhibitors (24, 25). Despite a ten-fold reduction of MCM3 protein (Figure 1B) proliferation of HCT116 cells was not impaired compared to cells transduced with either the EBNA1 or Ff-luciferase shRNAs that did not target genes endogenous in HCT116 cells (Figure 1C - left plate). In contrast, MCM3 knockdown strongly impaired plasmid stability (Figure 1C - right plate). The low plasmid stability ratio for cells transduced with the EBNA1 shRNA and the MCM3 shRNA demonstrated that the plasmid stability ratio was a useful measure for shRNAs that target plasmid maintenance while having little effect on cell proliferation. (Figure 1D).

We tested the effect of 230 different shRNAs on cell proliferation and plasmid stability (Supplemental Table 1) that included 44 shRNAs targeting the expression of 16 known DNA replication factors. The remaining 186 shRNAs targeted 55 genes of poorly characterized function that are either up-regulated in expression during G1 and S phases of the cell cycle (26) or encode proteins found to interact with human ORC (S. Prasanth and B. Stillman, unpublished data). Selection of shRNAs tested in the screen was not based upon prior experimental data demonstrating efficacy and so variable potency of gene knockdown was obtained with these reagents ranging from poor to strong knockdown of gene expression. Twenty six shRNAs targeting the expression of 14 known replication factors impaired plasmid stability with ratios of 0.45 or less, consistent with plasmid stability being a sensitized reporter of DNA replication (Figure 1E). Western blots revealed that shRNAs yielding plasmid stability ratios greater than 0.45 are less potent at knocking down their gene targets compared to shRNAs that target the same gene but yield plasmid stability ratios of less than 0.45 (Supplemental Figure 1A-D).

We established the criteria for consideration as a hit in the screen as any gene in which two or more shRNAs targeting its expression yielded plasmid stability ratios of 0.45 or less. Six of the 55 genes we tested met this criterion (Supplemental Table 2). One gene, AND-1, was a gene of unknown function at the time it was tested but has since been shown to have an essential role in DNA replication where it interacts with MCM10 and is required for loading of DNA polymerase alpha onto origins-of-replication (27). This validated the utility of this assay in identifying new DNA replication factors. Other hits from the screen included the DEAD-box protein DDX5 (p68), the condensin II subunit NCAP-G2, the PLK1 interacting protein PICH, and two proteins of unknown function C14ORF130 and KDELC1. For DDX5, AND1, and NCAP-G2 knockdown correlated well with the observed plasmid stability ratios where the more potent shRNAs targeting the expression of these genes resulted in the greatest impairment of plasmid stability (Supplemental Figure 2A-C).

DDX5 is known to interact with the related DDX17 and in some cases, but not all, they share overlapping activities in cells. shRNAs targeting DDX17 were included in the screen and despite ten-fold knockdown of DDX17 none of them impaired plasmid stability (Supplemental Figure 3A-B). This was in contrast to DDX5 shRNAs that demonstrated knockdown of DDX5 protein level four-fold and strongly impaired plasmid stability (Supplemental Figure 2B). This result suggested that the activity of DDX5 that underlies its role in plasmid stability was separable from its overlapping activities with DDX17 in the cell.

DDX5 is required for both plasmid stability and cancer cell proliferation

Additional shRNAs targeting DDX5 expression were tested to verify the requirement for DDX5 in plasmid stability (Figure 2A). All the DDX5 shRNAs strongly impaired plasmid stability compared to cells transduced with the Ff-luciferase shRNA (Figure 2B). The most potent DDX5 shRNAs: DDX5mi2053, DDX5mi1314, and DDX5mi2008 also strongly impaired HCT116 proliferation (Figure 2C) demonstrating a requirement for DDX5 in proliferation in this cancer cell line. Interestingly, DDX5 is overexpressed in HCT116 cells compared to non-tumor colon cell lines (19). Despite the sensitivity of HCT116 cells to DDX5 depletion we noted that not all cell lines required DDX5 to proliferate (see below) and so we investigated why HCT116 cells were dependent upon DDX5.

Figure 2.

Figure 2

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DDX5 is required for plasmid stability and cell proliferation. (A) Quantitative western blot analysis of DDX5 and Beta-actin (loading control) in whole cell extracts obtained from cells transduced with the indicated shRNAs (V2HS_26045 and V2HS_24063 are shRNAs used in the screen that target DDX5 expression whereas the other DDX5 shRNAs shown are new shRNAs not tested in the screen). (B) Colony formation results from the plasmid stability assay for either duplicate (each DDX5 shRNA) or quadruplicate (Luciferasemi203 shRNA) transduced cultures. (C) Colony formation results from the proliferation assay. Quantification of colony formation results are plotted in (B) and (C).

DDX5 is required for entry into S-phase

We were interested in whether DDX5 contributes toward DNA replication so the effect of depletion of DDX5 on cell cycle progression was examined. siRNAs were used to knockdown DDX5 since their effects on cells were more acute. The DDX5si2008 and DDX5si2053 siRNAs were selected since they were the most potent at depleting DDX5 when embedded within the Mir30-based shRNA backbone (Figure 2A). The EBNA1si1666 siRNA was used as the negative control since EBNA1 is not an endogenous protein in HCT116 cells and since the EBNA1mi1666 shRNA did not impair cell proliferation (Figure 1C). Within 48hrs after siRNA transfection DDX5 was knocked down 90% or greater by both DDX5 siRNAs, with the DDX5si2008 siRNA being more potent than the DDX5si2053 siRNA (Figure 3A). Knockdown of DDX5 resulted in an increased fraction of cells in G1 phase and reduced fraction of cells in S-phase (Figure 3B, Supplemental Figure 4A). Moreover, the efficiency of DNA replication in S-phase cells is reduced upon DDX5 depletion (Supplemental Figure 4B).

Figure 3.

Figure 3

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DDX5 is required for G1-S phase progression. (A) Quantitative western blot analysis of DDX5 and Beta-actin (loading control) in whole cell extracts obtained from cultures 48hrs after transfection with the indicated siRNAs. (B) Flow cytometry analysis of cell cycle in 48hr post-transfection cells with either DDX5si2008 (left panel) or EBNA1si1666 (right panel). (C) Progression into S-phase following serum addition to serum starved cultures previously transfected with either DDX5si2008 (circles) or EBNA1si1666 (squares) siRNAs. The fraction of cells in S-phase at each time point after serum addition was determined using flow cytometry analysis of propidium iodide incorporation and gating cells with greater than 2C but less than 4C DNA content. (D) Western blot analysis of RB, G1/S cyclin expression, and DNA replication factor expression in G1 and early S-phase whole cell extracts obtained from cells at increasing time following addition of serum to serum-starved cells previously transfected with the indicated siRNAs. (E) Western blot analysis of replication factors in chromatin fractions obtained from siRNA transfected S-phase cells over time after serum addition. Note the upper band detected on the CDC45 blots and marked with an asterisk in (D) and (E) is a non-specific cross-hybridizing protein recognized by the antibody.

Next we investigated whether DDX5 depletion impaired cell cycle entry and progression following addition of serum to serum-starved cells. Cells were transfected for 24hrs with the indicated siRNAs then they were incubated in serum-free media for another 48hrs. Serum was then added to the media and cell cycle progression monitored using flow cytometry and western blot analysis. Following serum addition, cells with DDX5 knockdown progressed slower into S-phase than cells transfected with the negative control siRNA (Figure 3C). DDX5 depletion did not block the ability of cells to re-enter the cell cycle and progress through G1 phase since RB hyper-phosphorylation and expression of G1/S cyclins were unaffected (Figure 3D). We also analyzed both the expression and loading onto chromatin of different subunits of the CMG complex, which is required for initiation of DNA replication and hence cell cycle progression into S phase where it functions to unwind duplex DNA ahead of DNA synthesis (28). The expression of different subunits of the CMG complex including MCM2, MCM5, and CDC45 were impaired (Figure 3D). Moreover, the loading of these subunits onto chromatin at time points corresponding to when cells entered and progressed through S-phase was impaired by DDX5 depletion (Figure 3E). We conclude that DDX5 contributes toward initiation of replication and thus S-phase entry where it promotes DNA replication pre-initiation complex assembly on chromatin. Since both episomal plasmid stability and cell proliferation were dependent on MCM5 and CDC45 expression (Supplemental Figure 1D and Supplemental Table 1) the reduced expression of both these proteins in cells with DDX5 knockdown likely contributes toward the requirement of DDX5 for plasmid stability and cell proliferation. Since DDX5 depletion resulted in reduced abundance of multiple different CMG proteins in whole cell extracts this suggests that DDX5 indirectly contributes toward DNA replication by regulating the expression of DNA replication genes.

We were interested in performing RNAi rescue experiments with wild-type and various mutant DDX5 transgenes in HCT116 cells to identify key amino acids in DDX5 that are required for its activity in cell cycle progression. While we could detect overexpression of the transcript encoding the RNAi resistant DDX5 transgene (Supplemental Figure 5A-B) we were unable to obtain a sufficient level of RNAi resistant DDX5 protein expression in either HCT116 cells or other cell lines for RNAi rescue experiments (Supplemental Figure 5C). This was true regardless of whether we used retroviral transfer or transient transfection methods for expression of either tagged or untagged DDX5 transgenes. These results suggest that expression of DDX5 is tightly regulated in cells and we are currently investigating this regulation to develop a method to achieve sufficient RNAi resistant DDX5 transgene expression for these studies.

DDX5 contributes toward DNA replication gene expression

DDX5 is involved in gene transcription (9). The observation that MCM2, MCM5, and CDC45 proteins are all down-regulated in DDX5 depleted cells released from serum starvation suggests that DDX5 may be regulating their expression. Indeed knockdown of DDX5 in asynchronous HCT116 cultures resulted in the down-regulation of a number of DNA replication proteins including components of the pre-replication complex (ORC1, ORC6, and CDC6), CMG subunits (MCM2, MCM5, and CDC45), and MCM10 (Figure 4A). This down-regulation also occurred with their transcripts at both 24hrs and 48hrs post-siRNA transfection (Figure 4B). Importantly, down-regulation of transcripts for DNA replication factors was observed within 24hrs after DDX5 siRNA transfection largely preceding the accumulation of cells in G1 phase that occurred by 48hrs post-siRNA transfection (Supplemental Figure 6A and 6B). Moreover RB hyper-phosphorylation was unaffected by DDX5 knockdown at 24hrs post-siRNA transfection indicating that the reduced expression of DNA replication genes at this time point was not due to suppression of their expression by hypo-phosphorylated RB (Supplemental Figure 6C). We performed microarray analysis on transcripts isolated from 24hrs post-siRNA transfected cells to determine how DDX5 knockdown affects the expression of other DNA replication genes and found that the expression of many genes annotated in the DNA replication gene ontology group were down-regulated in expression in DDX5 depleted cells (Figure 4C). Gene set analysis performed using GAGE (29) confirmed that the down-regulation of the DNA replication gene set in DDX5 depleted samples was significant (FDR = 3.85 × 10−17). Thus changes in replication gene expression arising after DDX5 knockdown was occurring prior to changes in cell cycle progression supporting that these gene expression changes were contributing toward this phenotype.

Figure 4.

Figure 4

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DDX5 is required for expression of DNA replication genes. (A) Quantitative western blot analysis of DNA replication factors in whole cell extracts obtained from asynchronous cell cultures 48hrs post-siRNA transfection with the indicated siRNAs. The asterisk shown in the MCM2, MCM10, and CDC45 blots indicate the position of non-specific proteins detected by these antibodies. (B) QPCR analysis of DNA replication factor transcript abundance in cells either 24hrs or 48hrs after transfection with either DDX5si2008 (red bar), EBNA1si1666 (blue bar) or mock transfected (no siRNA – green bar). Results for each transcript are normalized to the abundance of the indicated transcript in cells transfected with the EBNA1si1666 siRNA. Error bars indicate standard deviations calculated from 3 independent experiments. (C) Heat map showing row-wise standardized expression level for DNA replication genes 24hrs after transfection of cells with the indicated siRNAs.

DDX5 is required for RNA Polymerase II recruitment to promoters of DNA replication genes

We analyzed promoters of genes down-regulated as a result of DDX5 knockdown for common transcription factor binding motifs and found that that the E2F binding motif was frequently present (FDR = 4.55 × 10−6) (Supplementary Figure 7). Thus we tested whether DDX5 interacted with the transcription factor E2F1 and indeed found an interaction between these two proteins (Figure 5A). This interaction is direct since purified GST-E2F1 interacted with in vitro transcribed and translated DDX5 (Figure 5A). We also tested whether DDX5 localized to promoters of the CDC6, CDC45, and MCM5 genes that encode E2F binding sites where we confirmed that CDC6 expression is E2F1-dependent in this cell line (Supplemental Figure 8). We observed enrichment for DDX5 at these promoters compared to a non-E2F regulated promoter (Figure 5B), however, this interaction was not required for E2F1 to localize to these promoters since knockdown of DDX5 did not impair E2F1 localization (Figure 5C). These results suggested a direct role for DDX5 in DNA replication factor gene expression.

Figure 5.

Figure 5

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DDX5 localizes to promoters of DNA replication genes and functions in the loading of RNA Polymerase II onto these promoters. (A) Western blot analysis of E2F1 and DDX5 in DDX5 and IgG immunoprecipitation samples from nuclear extracts prepared from HCT116 cultures (top) and analysis of the interaction of in vitro transcribed and translated radiolabeled DDX5 with GST-E2F1 purified from bacteria and immobilized on glutathione beads (bottom). (B) Q-PCR analysis of DDX5 ChIP samples at the indicated promoters. Cells were serum starved, then re-stimulated with serum for 2 hrs and then harvested for ChIP. Note that the CD4 promoter is used as negative control. Error bars show the standard deviations for duplicate experiments. The green bars correspond to results for the DDX5 ChIPs whereas the black bars correspond to results for the IgG control ChIPs. All primer pairs used for Q-PCR amplify within 200bp of the transcription start sites of the indicated genes. Q-PCR analysis of E2F1 ChIP (C), Acetylated Histone H3 ChIP (D), RNA Polymerase II ChIP (E), and TFIIB ChIP (F) at the indicated promoters in asynchronous cells transfected with either DDX5si2008 (blue bars) or EBNA1si1666 (red bars) siRNAs. Error bars show the standard deviations for duplicate experiments. Neither the GAPDH nor CD4 transcripts are down-regulated by DDX5 knockdown and therefore their promoters are used as negative controls in for the ChIP experiments presented in (D), (E), and (F).

We investigated histone modifications and the abundance of proteins required for gene expression at DNA replication gene promoters. DDX5 knockdown did not reduce the abundance of either acetylated Histone H3 or acetylated Histone H4 at these promoters (Figure 5D, data not shown). In contrast, DDX5 knockdown significantly reduced the abundance of both RNA Polymerase II and TFIIB at the DNA replication gene promoters (Figure 5E and 5F). These results were consistent with the requirement of DDX5 for expression of these genes and indicated that DDX5 directly functions in transcriptional pre-initiation complex assembly at their promoters.

The DDX5 locus is amplified in breast cancer

The gene expression data revealed a significant overlap between genes down-regulated by DDX5 knockdown and genes described as a proliferation cluster in breast cancer (FDR = 8.3 × 10−17) (30). Proliferation cluster genes are those whose expression are positively correlated with mitotic index of a given breast tumor sample. Most of the genes shared between these two sets of expression data were down-regulated in DDX5 depleted cells, supporting the idea that DDX5 was required for expression of genes that promoted cell proliferation and suggested a connection between DDX5 and breast cancer cell proliferation. We also extracted expression data for 533 breast tumors from The Cancer Genome Atlas (TCGA) database (31) and analyzed this to identify gene pathways that were correlated with DDX5 expression and found that genes annotated to the S-phase pathway in Reactome were significantly correlated with DDX5 expression (FDR=0.08; with an odds ratio of 2.12). Combined with results from a recent study demonstrating that the DDX5 protein is frequently overexpressed in breast cancer (20), these observations prompted us to investigate whether the DDX5 gene was amplified in breast cancer. Our analysis included genome copy number data obtained from a previous study where representational oligonucleotide microarray analysis (ROMA) was applied to breast cancer samples obtained from the Karolinska Institute, Sweden and the Oslo Micrometastasis Study, Oslo, Norway (32). We found that the DDX5 locus was amplified in 63 of 255 breast cancer genomes (Figure 6A – red bar). We also analyzed breast tumor data available in the TCGA database for DDX5 copy number changes and observed DDX5 to be amplified in 26 of 93 tumors where DDX5 copy number was correlated with expression (z-score = 0.64, p = 6.5 × 10−12). This was consistent with our analysis of DDX5 copy number vs. mRNA levels in 33 established breast cancer cell lines where we found a Pearson correlation coefficient of 0.59 further demonstrating that DDX5 was often overexpressed in cells where it was amplified (Figure 6B).

Figure 6.

Figure 6

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The DDX5 locus in amplified in breast cancer. (A) Representative plots of genome copy number variation across chromosome 17 in selected breast cancer specimens. The green vertical bar indicates the position of the ERBB2 locus and the red vertical bar indicates the position of the DDX5 locus. (B) Analysis of DDX5 gene copy number and mRNA abundance in 33 established breast cancer cell lines. (C) Frequency of DDX5 amplification in the different breast cancer subtypes. (D) Results of colony formation assays performed with MDA-MB-453 cells transduced with either the DDX5mi2008 shRNA (red line) or empty vector (no shRNA - blue line) then expanded in media containing increasing concentrations of trastuzumab. Percent inhibition was calculated using the equation, (1 – AN / A0) × 100 where AN is the measured absorbance for either DDX5mi2008 or empty vector transduced cell cultures grown in media supplemented with a specific concentration of trastuzumab “N” and A0 is the measured absorbance of either the DDX5mi2008 or empty vector transduced cell cultures grown in media without trastuzumab. Note that the curve for the DDX5mi2008 transduced cells is shifted to the left indicating greater sensitivity of these cells to trastuzumab than the empty vector transduced cells. Error bars show standard deviations for duplicate cell cultures.

While analyzing the genomic data obtained from the Scandinavian breast tumors we found that the DDX5 and ERBB2 loci were frequently co-amplified since 37 of 63 DDX5 amplified breast cancer samples also had amplification of the ERBB2 locus (Figure 6A – green bar). The p-value from Fisher’s exact test for this degree of coincidence is 3.9 × 10−15. Although the ERBB2 and DDX5 genes are both present on chromosome 17 we see many instances where they were amplified as separate, independent amplicons (e.g. Figure 6A). This was consistent with the correlation between DDX5 overexpression and ERBB2 expression previously reported in breast cancer (20). Subtype information was available for the subset of 104 tumors collected from the Oslo Micrometastasis Study and we found DDX5 amplification in each subtype with the exception of the “normal” unclassified subtype (Figure 6C). Because DDX5 is frequently amplified in ERBB2 positive breast tumors we tested whether DDX5 knockdown affected the sensitivity of ERBB2 positive breast cancer cells to trastuzumab and indeed we found that DDX5 knockdown increased the sensitivity of these cells to inhibition of proliferation by trastuzumab (Figure 6D, Supplemental Figure 9A-C).

Breast cancer cells with amplification of DDX5 are more sensitive to its depletion than breast cancer cells lacking amplification of DDX5

We were interested in testing whether there were differences in the sensitivity of breast cancer cells to DDX5 depletion for DDX5 amplified vs. non-amplified cell lines and so we identified breast cancer cell lines with or without amplification of the DDX5 locus (Supplemental Figure 10A-B). The sensitivity of four different breast cancer cell lines with amplification of the DDX5 locus to DDX5 depletion by 3 different potent shRNAs was tested and significant inhibition of proliferation was observed (Figure 7A). Knockdown efficiency of DDX5 in the different cell lines varied. The strongest knockdown of DDX5 was achieved in the MDA-MB-453 and SK-BR-3 cell lines where 90% knockdown of DDX5 protein was observed, resulting in 5-to-10 fold inhibition of cell proliferation. In contrast, when we transduced three different breast cancer cell lines lacking amplification of the DDX5 locus and the breast epithelial cell line, MCF10A, with the DDX5 shRNAs none of the DDX5 shRNAs impaired their proliferation 2-fold or greater despite achieving 90% or more knockdown of DDX5 in each of these cell lines (Figure 7B). We concluded that breast cancer cells having amplification of the DDX5 gene were more dependent upon its expression to proliferate than breast cancer cells lacking this amplification.

Figure 7.

Figure 7

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Differential sensitivity of breast cancer cells to DDX5-depletion. (A) Quantitative western blot analysis of DDX5 and Beta-actin and colony formation assay results for different breast cancer cell lines with amplification of the DDX5 locus following transduction of the indicated DDX5 shRNAs or EBNA1 shRNA (negative control). Representative colony formation assay cultures are shown. Plots reflect quantitated results for the colonies formation assays. Error bars show standard deviations for triplicate cultures (B) Same as in “A” except breast cancer cell lines lacking DDX5 amplification are tested. (C) Western blot analysis of DNA replication factors in the indicated cell lines transduced with either DDX5mi2008 (“D5”) or EBNA1mi1666 (“E”). The asterisk indicates a non-specific cross-hybridizing band detected by the CDC45 antibody in the SK-BR-3 and MCF10A whole cell extracts. (D) QPCR analysis of the indicated transcripts in SK-BR-3 cells transduced with either DDX5mi2008 (red bars) or EBNA1mi1666 (blue bars). Results are normalized to the relative abundance of the indicated transcripts in the EBNA1si1666 transfected cultures. Error bars indicate the standard deviations from duplicate experiments. (E) Q-PCR analysis of RNA Polymerase II ChIP samples obtained from SK-BR-3 cells transduced with either DDX5mi2008 (red bars) or EBNA1mi1666 (blue bars). Results presented are normalized to the relative abundance of the indicated promoters in ChIP samples obtained from EBNA1si1666 transfected cultures. Error bars indicate the standard deviations from duplicate experiments.

Since DDX5 promoted expression of DNA replication factors in HCT116 cells we tested how DDX5 depletion affected the expression of DNA replication genes in the DDX5-amplified breast cancer cell line SK-BR-3 vs. the DDX5-nonamplified cell lines HCC1143 and MCF10A and observed that DDX5 depletion led to down-regulation of DNA replication proteins only in SK-BR-3 cells (Figure 7C). In SK-BR-3 cells DDX5 knockdown resulted in the down-regulation of DNA replication gene transcripts (Figure 7D) and reduced RNA Polymerase II loading onto the gene promoters (Figure 7E). These results were analogous to observations in HCT116 cells and support the conclusion that DDX5 was playing an important role in the initiation of transcription of DNA replication genes in breast cancer cell lines with DDX5 amplification. Moreover we tested whether DDX5 interacted with E2F1 in either MDA-MB-453, SK-BR-3 or MCF10A cells and found a co-IP interaction between DDX5 and E2F1 in MDA-MB-453 and SK-BR-3 cells but not MCF10A cells (Supplemental Figure 11) indicating that this interaction did not occur in all cell lines but rather was acquired by MDA-MB-453 and SK-BR-3 cells.

DISCUSSION

We found that DDX5 is required for plasmid stability and G1-to-S phase progression in a colorectal cancer cell line that overexpresses DDX5 where it regulates DNA replication by directly promoting the expression of DNA replication genes. DDX5 depletion impairs DNA replication factor gene expression in the absence of an effect on either RB phosphorylation or E2F1 localization to their promoters, indicating a role for DDX5 downstream of RB and E2F to elevate DNA replication factor gene expression and enhance cell proliferation. DDX5 is required for the general transcription factor TFIIB and RNA Polymerase II to localize to the promoters of genes encoding DNA replication factors, demonstrating an activity in the early steps of transcription initiation at these promoters. This mirrors the observations of others who studied transcriptional activation of promoters regulated by estrogen receptor alpha and MyoD (33, 34). In the former study DDX5 localized to the pS2 promoter in response to estradiol treatment and prior to TFIIB and RNA Polymerase II recruitment. In the later study DDX5 was required for the localization of the chromatin remodeler, BRG1, as well as the general transcription factor, TBP, and RNA Polymerase II to both the MHCIIB and TNNC2 promoters following induction of C2C12 cell differentiation. Consistent with this latter study we do not observe an effect of DDX5 knockdown on histone acetylation at DDX5-dependent promoters indicating that DDX5 is not playing an essential role in HAT recruitment.

DDX5 was initially identified as a host protein that cross-reacted against an antibody raised against SV40 Large T antigen (7). This similarity between DDX5 and Large T may mark a conserved activity since Large T has also been shown to function in transcriptional pre-initiation complex assembly at gene promoters (35, 36). Our results suggest that relief of RB suppression may alone not be sufficient to up-regulate E2F-dependent expression of DNA replication factor genes and hence cell proliferation. Consistent with this, HCT116 cells that overexpress DDX5 also lack p16INK4a expression, which antagonizes RB phosphorylation by Cyclin D – CDK4, and our analysis of gene expression data for 533 breast tumors available from the TCGA database revealed DDX5 expression in breast cancer is correlated with Cyclin D1 expression (p = 2.38 × 10−6) and is inversely correlated with p16INK4a expression (p = 5.67 × 10−4). This indicates that the proliferative advantage conferred by DDX5 overexpression to tumors occurs in the presence of mutations that impair the RB pathway. We suggest that amplification or overexpression of DDX5 leads to increased transcription via a new mechanism that is downstream of RB regulation. Such deregulation most likely requires DDX5 interacting with the gene promoters because we have localized it to these gene regulatory elements and it is required for efficient loading of TFIIB and RNA polymerase II. Recently, an essential role for ncRNAs in the regulation of E2F-dependent gene expression has been demonstrated (37). We propose that DDX5 may couple this ncRNA activity or the activity of another ncRNA at E2F promoters with RNA Polymerase II recruitment to promote gene expression in the absence of repression by RB.

The activity of DDX5 in promoting DNA replication gene expression is cell context dependent since it contributes toward the expression of these genes only in cell lines where it is either overexpressed (HCT116) and/or amplified (SK-BR-3). DDX5 knockdown did not affect the expression of DNA replication factors in either the HCC1143 or the MCF10A cell lines in which the DDX5 gene is not amplified. It is possible that the activity of DDX5 at DNA replication promoters is redundant with another protein where this redundancy is lost in cancer cells with DDX5 overexpression/ gene amplification. Alternatively, DDX5 may not normally function in DNA replication gene expression but this activity may be acquired during tumorigenesis, perhaps oncogene stimulated. Consistent with the latter, DDX5 interacts with E2F1 in the DDX5 amplification cell lines MDA-MB-453 and SK-BR-3 cells but not in MCF10A cells. These results point toward the positive role for DDX5 in DNA replication factor gene expression being acquired during cancer development, where it up-regulates expression of these genes and confers a significant advantage to cancer cell proliferation. Importantly, our data suggests that cancer cells then become dependent on DDX5 expression, a vulnerability that may be exploited in therapy. We are currently investigating whether upstream oncogenic signaling promotes DDX5 involvement in DNA replication factor gene expression. The observations that DDX5 is amplified in 25% of the breast cancer genomes we analyzed, 28% of the breast tumors in the TCGA database, that its expression in breast tumors is correlated with S-phase gene expression, and that DDX5 is required for DNA replication factor expression in amplified breast cancer cells supports a broad dependence of breast cancers on DDX5 activity and underscores the need to understand whether this activity can be targeted by pharmacologic inhibitors. Moreover, since depletion of DDX5 enhances the sensitivity of ERBB2 positive breast cancer cells to trastuzumab, an inhibitor against DDX5 may enhance the efficacy of trastuzumab in breast cancer therapy. Cyclin D is essential for neu driven tumorigenesis in mouse models and for proliferation of ERBB2 positive breast cancer cells (38, 39). Combination treatment of ERBB2 amplified breast cancer cells with flavopiridol, a cyclin dependent kinase inhibitor, and trastuzumab results in synergistic inhibition of proliferation (40). Cyclin D-CDK4 hyper-phosphorylates RB to de-repress E2F-dependent gene expression and inhibition of Cyclin D-CDK4 should restore RB-mediated repression of these genes. We propose the synergistic inhibition of cell proliferation resulting from DDX5 knockdown and trastuzumab treatment follows a similar mechanism as Cyclin D inhibition where DDX5 depletion impairs RNA Polymerase II recruitment to E2F-regulated promoters and thus antagonizes E2F-dependent gene expression.

We observed frequent amplification of DDX5 in luminal subtype breast cancers consistent with the previously described activity of DDX5 as a transcriptional co-activator of estrogen receptor alpha dependent gene expression (41). We also observed frequent co-amplification of the ERBB2 and DDX5 genes. This agrees with the significant correlation reported for ERBB2 and DDX5 expression in a panel of estrogen receptor alpha positive breast tumors (20). However, our analysis of the ERBB2/DDX5 double positive breast cancers did not reveal a correlation with estrogen receptor expression and thus suggests an estrogen receptor independent activity for DDX5 in breast cancer. Indeed, the SK-BR-3 and MDA-MB-453 breast cancer cell lines we found to be dependent upon DDX5 to proliferate are negative for estrogen receptor expression. Interestingly, in addition to identifying a significant correlation between DDX5 and ERBB2 expression in their panel of estrogen receptor alpha positive breast cancers the aforementioned study also reported a significant correlation between DDX5 and AIB1 (aka NCOA3) expression. NCOA3 has been demonstrated to be a transcriptional co-activator of E2F-regulated genes (42, 43). In light of our results, we suggest that DDX5 and NCOA3 may cooperate in breast cancer to up-regulate the expression of DNA replication genes and thus promote cancer cell proliferation.

Our findings that the DDX5 gene is frequently amplified in breast cancer and that breast cancer cell lines harboring this amplification are more sensitive to its depletion than breast cancer cell lines that lack DDX5 amplification suggest that DDX5 may function as an oncogene. Overexpression of DDX5 in murine fibroblasts promotes transformation and tumor formation in nude mice (44). However we have been unable to overexpress a DDX5 transgene in many different human and murine cancer and non-cancer cell lines (see Supplemental Figure 5A-C). This has hampered our efforts to test whether elevated DDX5 expression transforms breast epithelial cells and also to identify mutants in an RNAi resistant DDX5 transgene that do not restore cell proliferation to DDX5-dependent cell lines with endogenous DDX5 knockdown. We speculate that co-expression of DDX5 with another protein and/or ncRNA may enable robust expression of the DDX5 transgene and we are currently investigating this hypothesis.

The data herein suggest that DDX5 is a viable candidate drug target for selective anti-cancer therapy directed at those tumors that have an amplified DDX5 locus. We are currently testing this notion by performing a screen for inhibitors of DDX5 activity. Like treatment with trastuzumab that is linked to tumors harboring amplification of the HER2 gene, cancer treatment targeting DDX5 could be linked to those breast cancers that have this locus amplified.

Materials and Methods

A detailed description of materials and methods are provided in supplementary material.

Antibodies

Western blot analysis: From Bethyl Laboratories - anti-DDX5 cat. # A300-523A, anti-DDX17 cat. # A300-509A, anti-MCM5 cat. # A300-195A, anti-AND1 cat. # A301-141A, and anti-NCAP-G2 cat. # A300-605A; from Sigma, anti-Beta Actin cat. # A5316, anti-PCNA cat. # P8825, and anti-E2F1 cat. # E8901; from Abcam - anti-CDC45L cat. #ab56476; from Cell Signaling - anti-CCND1 cat. # 2922 and anti-CCNE2 cat. # 4132; from Novus - anti-CCNA2 cat. # NB100-91726; from Pharmingen - anti-RB cat. # 554136; and from Proteintech Group - anti-MCM10 cat. #12251-1-AP. In-house antibodies that were used include anti-MCM2 #CS732 (polyclonal), anti-MCM3 #738 (polyclonal), anti-CDC6 #1881 (polyclonal), anti-ORC1 clone PKS 1-40 (monoclonal), anti-ORC6 clone 30 (monoclonal), anti-ORC2 pAB205B (polyclonal), anti-ORC3 PKS16-11 (monoclonal), anti-RB C-36 (monoclonal) and anti-RB X2-55 (monoclonal). The antibody against PSF2 was kindly provided by Dr. Juan Mendez. For immunoprecipitation experiments rabbit anti-DDX5 from Bethyl Laboratories cat. # A300-523A and normal Rabbit anti-IgG from Caltag Laboratories cat. # 10500C were used. For ChIP experiments rabbit anti-DDX5 from Bethyl Laboratories cat. # A300-523A, Mouse anti-E2F1 from Sigma cat. # E8901, Rabbit anti-acetyl-Histone H3 from Millipore cat. # 06-599, Rabbit anti-RNA Polymerase II from Santa Cruz cat. # sc-899, and Rabbit anti-TFIIB from Santa Cruz cat. # sc-225X were used.

Cell Lines and growth media

HCT116 cells were provided by Dr. Richard Boland and propagated in DMEM (Cellgro #10-013-CV) + 10% Fetal Bovine Serum (FBS). SK-BR-3, MDA-MB-453, EVSA-T, ZR-75-1 and HCC1143 cells were provided by Dr. Scott Powers and propagated in either McCoy’s (Cellgro #10-050-CV) + 10% FBS (SK-BR-3), Leibovitz (ATCC #30-2008) + 10% FBS (MDA-MB-453), MEM (ATCC #30-2003) + 2mM L-Glutamine + 10% FBS (EVSA-T), or RPMI 1640 (Cellgro #10-040-CV) + 10% FBS (HCC1143 and ZR-75-1) respectively. EFM-19 and Hs578t cells were provided by Dr. Michael Wigler and were propagated in RPMI 1640 + 10% FBS and DMEM + 10μg/mL insulin + 10% FBS respectively. MCF10A cells were provided by Dr. Senthil Muthuswamy and propagated in DMEM/F12 (Gibco #11330) + 20ng/mL epidermal growth factor + 500ng/mL hydrocortisone + 100ng/mL choleratoxin + 10μg/mL insulin + 5% horse serum. No authentication of the cell lines was performed by the authors.

Plasmids

All shRNAs are designed in a Mir30 backbone and were either obtained from Open Biosystems (indicated as V2HS_xxxxxx in the text) or were self-prepared (indicated as GeneNamemixxxxxx in the text). The p220.2 plasmid encoding OriP, EBNA1, and hygromycin resistance was kindly provided by Dr. Anindya Dutta (4).

Screen

150,000 HCT116 cells were seeded per well in 6-well plates and allowed 24hrs to attach. They were then infected with the 36hr and 48hr viral supernatant (see supplementary material for packaging of retrovirus encoding shRNAs) for a total of two rounds of infection per culture. 24hrs after the second round of infection, cells from each well were suspended and seeded into new wells of 6-well tissue culture (TC) plates in DMEM + 10% FBS + 1.5μg/mL puromycin. Cultures were then selected for 4 days then were suspended by trypsinization and counted using a hemacytometer. 600,000 cells from each suspension were then seeded to separate wells of 6-well plates in 2mL DMEM + 10% FBS per well. Following 24hrs each culture was transfected with 1μg of p220.2 plasmid (encoding OriP, EBNA1, and hygromycin resistance) diluted in Opti-Mem, 4μL Lipofectamine (Invitrogen #18324-012), and 6μL Plus Reagent (Invitrogen #11514-015) using the manufacturer’s protocol. Transfected cultures were then incubated in a cell culture incubator (37 degrees C, 5% CO2) for 4.5hrs. The transfection media was then aspirated, each culture was washed with PBS, then 2mL DMEM + 10% FBS was added to each culture prior to returning to the incubator. 24hrs after p220.2 plasmid transfection, cells were suspended by trypsinization and counted using a hemacytometer. For the proliferation assay, one well of a 12-well TC plate was seeded with 5,000 cells per suspension in 1mL DMEM + 10% FBS + 1.5μg/mL puromycin. For plasmid stability assay, one well of a 12-well TC plate was seeded with 50,000 cells per suspension in 1mL DMEM + 10% FBS + 1.5μg/mL puromycin + 400μg/mL hygromycin. Selection media was replaced every 3rd day. Colonies in proliferation assay cultures were expanded for 8 days and plasmid stability assay cultures for 10 days prior to staining with crystal violet.

The plasmid stability ratio for each experimental shRNA was calculated by dividing the plasmid stability result (see colony formation assay description in Supplementary materials and methods) for the culture transduced with the experimental shRNA by the proliferation result for the culture transduced with the same experimental shRNA.

Serum starvation and chromatin fractionation experiments

HCT116 cultures seeded with 125,000 cells per well of 6-well plates and allowed 24hrs to attach were transfected with the indicated siRNAs as described in supplemental material. 24hrs following transfection the media was removed and each culture washed twice with 2mL DMEM (no serum) per culture. The cultures were then incubated 48hrs in DMEM media lacking serum. Following the incubation the serum-free media was removed from each culture and replaced with DMEM + 10% FBS. Chromatin fractionation was performed at the indicated time points after serum addition as described (45).

Quantitative PCR analysis

RNA was prepared from siRNA transfected HCT116 cultures at the indicated time points post-siRNA transfection using the RNeasy Mini Kit (Qiagen cat. # 74104) including on-column DNase digestion (Qiagen cat. # 79254) and eluted in the supplied RNase-free water. RNA from shRNA SK-BR-3 cultures was prepared from cells 8 days post-shRNA transduction. The cDNA used for Q-PCR was prepared from 1μg each RNA sample using TaqMan Reverse Transcription Reagents (Applied Biosystems #N808-0234) with random hexamer priming in a GeneAmp PCR system 9700 thermocycler. Each Q-PCR reaction was prepared using 2μL of 1-to-20 diluted cDNA and 13μL LightCycler 480 SYBR Green I Master Mix (Roche #04887352001) and were performed in 384-well plates using the LightCycler 480 (Roche) as per manufacturer’s instructions. Primer sequences used for analysis of cDNA samples are listed in supplemental material.

Q-PCR analysis of ChIP samples was performed using 2μL of the final ChIP samples that were each diluted in 60μL nuclease-free water following ethanol precipitation. 13μL LightCycler 480 SYBR Green I Master Mix (Roche #04887352001) was added to each ChIP sample and PCR reactions were performed in 384-well plates using a LightCycler 480 (Roche) as per manufacturer’s instructions. Primer sequences are listed in supplemental information.

Microarrays and Analysis

RNA was isolated from HCT116 cultures 24hrs after siRNA transfection. Preparation of cDNA, sample labeling, sample hybridization to microarrays, and microarray scanning were performed by the Microarray Shared Resource Facility at Cold Spring Harbor Laboratory. Samples were hybridized to the Affymetrix GeneChip Human Gene 1.0 ST Array. Raw data from the arrays was then processed using the RMA method with an up-to-date probe set definition (46, 47). The latest gene annotation information was retrieved from the Gene Ontology public database. Microarray data has been deposited in NCBI’s Gene Expression Omnibus and is accessible through GEO series accession number GSE36141 (ref (48), accession number GSE36141). Differential expression, gene set, and transcription factor motif analysis are presented in the supplementary information.

Nuclear extract preparation, immunoprecipitation, GST interaction assay, and chromatin immunoprecipitation assay

Descriptions are presented in the supplementary information.

Analysis of DDX5 copy number in breast tumors and copy number vs. expression in breast cancer cell lines

Genomic DNA was prepared from 33 breast cancer cell lines and copy number analysis was performed in these cell lines relative to a normal reference cell line using ROMA as described (32). Gene expression profiling was performed using NimbleGen Gene Expression arrays using conditions recommended by the manufacturer. Quantile normalization intensities obtained from the arrays were converted to log2 ratios by comparing results obtained from the breast cancer cell lines to the universal reference RNA.

Segmented DNA copy number profiles of chromosome 17 were generated from publicly available normalized copy number ratio data for the MicMa set (125 profiles) and for the WZ set (141 profiles) of breast tumor tissues (ref. (48) accession number GSE19425) (32, 49). The segmentation was performed as described (32).

Supplementary Material

1
2
3
4
5

Significance.

DDX5 is required for cell proliferation by controlling the transcription of genes expressing DNA replication proteins in cancer cells in which the DDX5 locus is amplified and this has uncovered a dependence on DDX5 for cell proliferation. Given the high frequency of DDX5 amplification in breast cancer our results highlight DDX5 as a promising candidate for targeted therapy of breast tumors with DDX5 amplification and indeed we demonstrate that DDX5 inhibition sensitizes a subset of breast cancer cells to trastuzumab (Herceptin®).

Acknowledgements

We thank Chris Johns and Sohail Khan of the CSHL Microarray Shared Resource for their contributions to the microarray experiment. We also thank Uli Bialucha, Lars Zender, Johannes Zuber, Wen Xue, Jackie Lees and members of the Stillman

Funding: for Mazurek and Stillman (NCI CA13016 and the Goldring Family Foundation), for Luo (NCI (NCI CA45508), for Powers (NCI CA124648), for Krasnitz and Hicks (DoD W81XWH-09-1-0591and the Breast Cancer Research Foundation).

Footnotes

Present Address: College of Computing and Informatics, University of North Carolina, Charlotte NC 28223

No financial conflicts of interest.

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Supplementary Materials

1
NIHMS399630-supplement-1.pptx (53.5MB, pptx)
2
NIHMS399630-supplement-2.docx (17.6KB, docx)
3
NIHMS399630-supplement-3.doc (76.5KB, doc)
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NIHMS399630-supplement-4.xlsx (71.1KB, xlsx)
5
NIHMS399630-supplement-5.pptx (57.7KB, pptx)

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