Down-regulation of CK2α correlates with decreased expression levels of DNA replication minichromosome maintenance protein complex (MCM) genes

Protein kinase CK2 is a serine/threonine kinase composed of two catalytic subunits (CK2α and/or CK2α’) and two regulatory subunits (CK2β). It is implicated in every stage of the cell cycle and in the regulation of various intracellular pathways associated with health and disease states. The catalytic subunits have similar biochemical activity, however, their functions may differ significantly in cells and in vivo. In this regard, homozygous deletion of CK2α leads to embryonic lethality in mid-gestation potentially due to severely impaired cell proliferation. To determine the CK2α-dependent molecular mechanisms that control cell proliferation, we established a myoblast-derived cell line with inducible silencing of CK2α and carried out a comprehensive RNA-Seq analysis of gene expression. We report evidence that CK2α depletion causes delayed cell cycle progression through the S-phase and defective response to replication stress. Differential gene expression analysis revealed that the down-regulated genes were enriched in pathways implicated in cell cycle regulation, DNA replication and DNA damage repair. Interestingly, the genes coding for the minichromosome maintenance proteins (MCMs), which constitute the core of the replication origin recognition complex, were among the most significantly down-regulated genes. These findings were validated in cells and whole mouse embryos. Taken together, our study provides new evidence for a critical role of protein kinase CK2 in controlling DNA replication initiation and the expression levels of replicative DNA helicases, which ensure maintenance of proliferative potential and genome integrity in eukaryotic cells.

In order to systematically examine the role of CK2α in the control of proliferation in non-cancerous cells, we created a myoblast cell line derived from H9c-2 cells with inducible down-regulation of CK2α. Cultured myoblasts are one of the models chosen for studying biological processes in vitro. The H9c-2 myoblast cell line isolated from ventricular tissue, is currently used as a mimetic for skeletal muscle but it also has the ability to differentiate towards a cardiac-like phenotype under appropriate experimental conditions responding similarly to neonatal cardiomyocytes to several stimuli 25 . We transduced H9c-2 myoblast cells with a tGFP-expressing lentiviral-based vector designed to express a short-hairpin RNA (shRNA) targeting rat CK2α under the control of doxycycline (Fig. 1a). Because a myoblast cell line with inducible down-regulation of CK2α had not been previously described, we first characterized biochemically the newly established cell line (hereafter referred to as H9c2-CK2α-44). To determine the timing and extent of transduction, H9c2-CK2α-44 cells were analyzed for tGFP expression following addition of doxycycline. Cells were harvested at various intervals for up to six days and green fluorescence emission was determined by flow cytometry. As indicated in Fig. 1b,c, virtually all the cells were able to express tGFP and showed increasing fluorescence signal in a time-dependent fashion indicating the successful stable transduction of the target cells. Levels of expression of CK2α were determined by Western blot. Results shown in Fig. 1d revealed high intracellular levels of CK2α than CK2α' in the absence of doxycycline. Incubation with doxycycline for up to six days resulted in nearly complete disappearance of CK2α protein, a slight increase in the expression of CK2α' and decreased expression levels of CK2β (Fig. 1e). To support the molecular effects of CK2α disappearance on a known intracellular CK2 target protein, we analyzed the phosphorylation status of PTEN at S380/T382/383 26 . Western blot analysis on whole lysate from cells treated as indicated in Fig. 1f revealed that the levels of phosphorylation of PTEN were decreased in cells with reduced expression of the individual CK2 catalytic isoforms as compared to control experiment (Fig. 1f, lanes 2 and 3 vs lane 1). PTEN phosphorylation further decreased when CK2α and CK2α' were simultaneously down-regulated (Fig. 1f, lane 4) suggesting that both isoforms contribute to PTEN phosphorylation.

Down-regulation of CK2α interferes with cell cycle progression and cell proliferation.
We examined the effect of CK2α silencing on the proliferation of H9c2-CK2α-44 cells essentially by three complementary approaches: i.e. FACS analysis, Western blot and BrdU-based assay. Incubation with doxycycline for three and six days, respectively, resulted in marked differences in the cell density as compared to control cells (Fig. 2a). Flow cytometry analysis of H9c2-CK2α-44 cells was carried out at various time points after the addition of doxycycline to determine whether the reported differences resulted from cell cycle perturbation and/or enhanced cell death (Fig. 2b). Analysis of DNA content revealed a reproducible and significant slightly increased G1 population for up to six days of incubation time with doxycycline as compared to control experiments. The fraction of cells in sub-G1, which provides an indicative measurement of cell death, was, however, negligible and could not explain the significant decrease in cell density observed in cells with lowered expression of CK2α (Fig. 2a). Next, we synchronized cells by serum starvation and looked at their ability to resume the cell cycle after adding complete growth medium. As shown in Fig. 2c, cells expressing CK2α resumed proliferation and reached the S phase after 12 h from serum deprivation as also confirmed by the analysis of the expression of cyclin E which is considered a critical regulator of the G1-S transition (Fig. 2d, 27,28 ). Conversely, cells lacking CK2α did not resume the cell cycle at the same pace showing, instead, a delayed progression from G1 to S phase (Fig. 2c,d).
In support of these results, cellular BrdU incorporation assay carried out with H9c2-CK2α-44 cells showed fewer cells in S-phase three days after addition of doxycycline suggesting that down-regulation of CK2α resulted in either less efficient or delayed entry into S-phase (Fig. 2e). Analysis of the parental cell line did not show any difference on BrdU incorporation under the same experimental conditions indicating that these perturbations of the cell cycle could account for the observed slow proliferation rate in cells lacking CK2α.
CK2α silencing results in enhanced sensitivity to replication stress. We hypothesized that delayed entry into S-phase could be caused by defective DNA replication initiation in cells with reduced levels of CK2α. If so, we anticipated that these cells would become highly sensitive to replication stress induced in the presence Results from one representative experiment are shown. (d) H9c2-CK2α-44 cells were treated with vehicle (i.e. dd water) or with 1 µg/ml doxycycline (Dox) for increasing amounts of time. Whole cell lysates were analyzed by Western blot employing a mouse monoclonal antibody against CK2α and CK2α' . β-actin detection served as loading control. (e) H9c2-CK2α-44 cells and the parental cell line were harvested after 0, 2 and 3 days of treatment with vehicle (−) or 1 µg/ml doxycycline (+) and whole cell lysates were analyzed by Western blot of DNA replication inhibitors. To test this hypothesis, we studied cell cycle progression in response to mild replication stress in the absence or presence of 0.1 μM aphidicolin for increasing amounts of times. As shown in Fig. 3a, treatment with aphidicolin and doxycycline significantly impaired the proliferation of H9c2-CK2α-44 cells as compared to control cells or cells treated with either compounds alone. Conversely, the proliferation rate of the parental cell line was not significantly affected by the treatment with the compounds used either alone or in combination (Fig. 3b). Analysis of DNA content by flow cytometry of cells treated with doxycycline revealed a slightly but reproducible higher percentage of cells in G1-phase (Fig. 3c). Cells treated with aphidicolin showed marginally increased accumulation of cells at the G1-S border, which was expected considering the low concentration of aphidicolin employed to induce mild replication stress. Finally, incubation with both compounds resulted in accumulation of cells in both G1 and S phases (Fig. 3c). Hence, their accumulation in the early phases of the cell cycle at the expense of G2/M may explain lack of proliferation observed following induction of mild replication stress.

Global gene expression profiling uncovers a novel role of CK2α in DnA replication initiation.
To shed light on the mechanisms by which CK2α regulates cell proliferation in myoblasts, we performed a global gene expression analysis with RNA isolated from H9c2-CK2α-44 cells left untreated or incubated with doxycycline for three days. The transcriptome analysis was also carried out with the parental cell line following siRNA-mediated down-regulation of CK2α to exclude off-target effects resulting from particular sites of integration of lentiviral-based constructs into the genome of the host cells. We performed two independent experiments in duplicate obtaining, therefore, four independent replicates for each cell line and condition using the Illumina platform. From the 2 × 100 bp paired end Illumina run and after trimming of the raw reads, we generated between 131.422.420 and 173.796.784 million reads for each of the four conditions. The reads were then mapped and aligned to the rat genome version rnor6.0 using STAR version 2.5.0c. On average for the four conditions a total of 95% of the reads were mapped and around 70% of the reads were mapped uniquely (Fig. 4a). Furthermore, the quality of the data was also confirmed from proper assessment of the fastQC files, which fulfilled all quality controls. Gene expression analysis was performed using DEseq2. By comparing the log2 fold-changes of gene expression in the two cell lines (i.e. H9c2-CK2α-44 and parental cell lines), we obtained a high correlation between the generated data (Fig. 4b) indicating that gene expression modifications induced by silencing of CK2α in H9c2-CK2α-44 cells was not influenced by the integration site of the lentiviral-based construct. Interestingly, we found that the down-regulated genes were remarkably more represented within specific cellular pathways than the up-regulated genes. Of the 15542 genes identified following analysis of the H9c2-CK2α-44 cells, 1318 genes were found to be either up-regulated or down-regulated (8.5%, padj <0.05). In the case of the parental cell line, of the 16435 identified genes, 1435 were found differentially expressed (8.7%, padj <0.05). H9c2-CK2α-44 and H9c-2 cell lines shared 95 up-regulated and 339 down-regulated genes, respectively, following silencing of CK2α. From the analysis of H9c2-CK2α-44 cells, 548 genes were found up-regulated while 770 resulted down-regulated (Fig. 4c).
To identify the cellular pathways significantly modified in H9c2-CK2α-44 cells and in the parental cell line we performed KEGG pathway analysis on the significantly down-regulated genes (Fig. 5). Interestingly, the transcriptome analysis revealed that loss of CK2α had a substantial negative effect on the expression of genes controlling cell cycle (114/115 genes), DNA replication (33 genes) and DNA damage repair [mismatch repair (21 genes), FA pathway (46 genes), homologous recombination (25 genes), nucleotide excision repair (42 genes) and base excision repair (32 genes), Fig. 5 and Table S1]. Within the list of differentially expressed genes we found down-regulation of proliferating cell nuclear antigen (PCNA) gene, a number of cyclin-coding genes (Ccna2, Ccnab1, Ccne1, Ccne2 and Ccnd2), E2F transcription factor 1 (E2f1) gene and the subunits 1 and 6 of the origin recognition complex (ORC1, ORC6) genes (Tables 1 and S2). Interestingly, among the most significantly down-regulated transcripts we found the minichromosome maintenance protein complex (MCM2-7) genes and those coding for DNA-directed polymerase epsilon, delta 1, alpha 2, delta 2, epsilon 2 (Pole, Pold1, Pola2, Pold2, Pole2, Table 1). Finally and as predicted, CK2α transcripts were among those most significantly down-regulated (log2 Fold-change −1.67791, padj 4.90981E-39) while CK2α' and CK2β transcripts, were not found significantly changed (results not shown). To the best of our knowledge, a cross-talk between CK2 and MCM proteins has never been reported before. Numerous studies have shown that these proteins form pre-replication (pre-RCs) complexes, also known as "origin licensing" that allow binding of DNA polymerases and other factors to chromatin to start DNA replication during the G1-phase [29][30][31][32] . Since their reduced expression levels in cells confers hypersensitivity to replication stress 33 , we decided to further investigate them.

CK2α down-regulation correlates with decreased McMs levels in vitro and in vivo.
Using qPCR we validated the expression of eight genes (i.e. MCM2-7, Polε and Polδ1, Fig. 6) in cells untreated or incubated employing the indicated antibodies. β-actin detection served as loading control. (f) H9c2-CK2α-44 cells were incubated with 1 µg/ml doxycycline for three days, transfected with scr-siRNA and CK2α'-siRNA for three days, respectively, as indicated in the figure. Last lane refers to cells treated with doxycycline and transfected with CK2α'-siRNA for three days. Whole cell lysates were analyzed by Western blot employing the indicated antibodies. All experiments were performed three times obtaining similar results; one Western blot experiment of three is shown. Abbreviations: LTR: 5' Long Terminal Repeat; Ψ: Psi packaging sequence; RRE: Rev response element; P TRE3G : Inducible promoter with tetracycline response elements; P mCMV : SMARTchoice promoter; Puro R : Puromycin resistance; 2a: Self-cleaving peptide; Tet-On 3G: Doxycycline-regulated transactivator protein; WPRE: Woodchuck hepatitis post-transcriptional regulatory element; 3'SIN LTR: 3' Self-inactivating long terminal repeat. (2019) 9:14581 | https://doi.org/10.1038/s41598-019-51056-5 www.nature.com/scientificreports www.nature.com/scientificreports/ www.nature.com/scientificreports www.nature.com/scientificreports/ with doxycycline for three days (shRNA qPCR1). This analysis confirmed down-regulation of the aforementioned genes. qPCR analysis in cells incubated with doxycycline for six days (shRNA qPCR2) showed reduced down-regulation of six (i.e. MCM2, 3,4,6,7 and Polε) of the eight genes analyzed suggesting that some adaptation might have occurred. Results of one representative experiment are shown +/− STDEV, *P < 0.05, **P < 0.005. (b) Comparison between H9c2-CK2α-44 and H9c-2 cells with respect to proliferation efficiency. Cells were treated essentially as described in (a) for the indicated times. Mean values+/− STDEV of one representative experiment out of three is shown. *P < 0.05, **P < 0.0005 with respect to CT, # P < 0.01. (c) H9c2-CK2α-44 cells left untreated or treated with 1 µg/ml doxycycline for three days were co-treated with 0.1% DMSO or 0.1 µM aphidicolin for additional five days. Cells were analyzed by flow cytometry following propidium iodide staining (PI) and events were quantified and expressed in percentage.
www.nature.com/scientificreports www.nature.com/scientificreports/ Next, we determined whether down-regulation of the MCM genes correlated with decreased expression levels of their coded proteins. For this, we analyzed whole lysates from cells treated with vehicle or doxycycline for three days. Results shown in Fig. 7a confirmed that silencing of CK2α results in decreased expression of MCM proteins with respect to control experiments.
In vivo validation of gene expression variability of two members of the MCM protein family, namely MCM3 and MCM4, was investigated in tissues of WT and CK2α-knockout embryos i.e.: the hearts (Fig. 7b,c) and somites at E10.5 (Fig. 7d,e). It was reported that a variety of defects in the heart at this developmental stage are probably responsible for the embryonic lethality observed in mice lacking CK2α 12 , therefore, we included the cardiac tissue in the analysis. In hearts, MCM3 and MCM4 specific signal was strongly detected in the cell nucleus. Negative controls for MCM4 showed some cytoplasmic background signal both in the myocardial wall and the trabeculae (Fig. S1a). We only analyzed the myocardial cells in non-trabecular myocardium, as the trabecular myocardium in CK2α−/− embryos is less developed than in the WT heart. Since MCM3 and MCM4 staining showed heterogeneous levels of fluorescence in WT hearts, we quantified separately cells emitting high and low fluorescence levels. Bar-graphs show a statistically significant lower number of cells emitting high intensity fluorescence in CK2α−/− E10.5 myocardium as compared to WT myocardium (Fig. 7b,c). In somites, MCM3 and MCM4 specific signal was strongly detected in the cell nucleus (Fig. 7d,e) while negative controls showed little background signal (Fig. S1b). Significant differences in the expression of MCM3 and MCM4 could be detected in WT and KO somites, respectively, suggesting that loss of CK2α affects MCM protein levels in vivo during early mouse development. www.nature.com/scientificreports www.nature.com/scientificreports/

Discussion
There is ample evidence showing that CK2α plays an important role in the regulation of cell division. In this study, we aimed at identifying potential novel molecular mechanisms controlling cell proliferation in non-cancerous cells mediated by CK2α. We show here for the first time that down-regulation of CK2α leads to a significant reduction in the expression levels of components of the MCM complex suggesting that MCM proteins could be www.nature.com/scientificreports www.nature.com/scientificreports/ responsible, at least in part, for the lowered proliferation rate observed in cells and in vivo. The minichromosome maintenance complex is a family of structurally related proteins with replicative helicase activity highly conserved from yeast to man that are required for cell proliferation, migration and invasion [34][35][36] . Mutations in individual MCMs have been shown to play a critical role in DNA replication initiation and result in embryonic death, growth retardation and limited fetal erythropoiesis [37][38][39][40] .
Reduced levels of CK2α resulted in decreased proliferation rate in H9c2-CK2α-44 cells (Fig. 2). Similar results were obtained when the mitotic index in CK2α−/− embryo hearts was compared to the one in WT embryos (Suppl. Fig. S2). Our data on WT embryonic heart proliferation fits well with previous reports showing that in WT embryos, OFT myocardium has the lowest proliferation rate of whole heart myocardium, and that proliferation rates diminish from E9.5 to E10 in the different heart regions 41,42 . Reduced proliferation was, however, not accompanied by a significant increase in cell death both in vitro (Fig. 2b) and in vivo (Table S3) suggesting that decreased proliferation rate may explain, at least in part, the growth defects observed in the CK2α−/− embryo heart. Impaired expression of CK2α caused a reproducible slight increase in the G1/S cell population (Fig. 2). We show here for the first time that DNA replication and DNA damage pathways are significantly down-regulated in CK2α-silenced myoblasts. Specifically, the expression of MCMs was reduced at both mRNA and protein levels. Ibarra    www.nature.com/scientificreports www.nature.com/scientificreports/ proliferating at a slower pace, accumulating in G1 phase and becoming hypersensitive to replication stress 33 . Accordingly, it is plausible that the increase in the G1/S phase population in CK2α-silenced myoblasts accompanied by enhanced sensitivity to replication stress is due to decreased MCMs expression levels.
It has been shown that mice hemizygous for the individual MCM helicases show compromised stability of the entire hexameric complex, cell proliferation defects and elevated micronuclei frequencies associated with genomic instability (GIN 43 ,). The analysis of CK2α−/− embryo hearts revealed a drastic impairment of cell proliferation, however, it did not show increased GIN (data not shown). It is possible that a certain threshold level of expression of the MCM helicases might be necessary to cause GIN and/or that the CK2α−/− embryos do not live long enough to accumulate chromosomal aberrations.
A strength of this work is that we used four biological replicates for the RNA-Seq analysis, and we obtained reproducible data on proliferation rates, cell cycle aberrations and MCM expression levels both in vivo and with cell lines. Part of the work has been carried-out with a clonal cell line for which the risk of clonal selectivity cannot be underestimated. However, results obtained with knockout mice (Fig. 7) and siRNA-treated cells (results not shown) corroborate the findings obtained with the H9c2-CK2α-44 cells.
We observed a slight increase in CK2α' expression levels in myoblasts depleted of CK2α (Fig. 1d). It is conceivable that the residual CK2 activity, due to the expression of CK2α' , can make up for the absence of CK2α. A compensatory mechanism is plausible since CK2α may compensate for the lack of CK2α' in mice 11 . This increase in CK2α' is similar to what has been reported in studies with mouse embryo fibroblasts (MEFs 44 ), and in contrast to the lack of increase seen in the CK2α−/− mice 13 . This suggests that a compensatory mechanism might occur in primary cells and cell lines to preserve a certain level of CK2 kinase activity necessary for the in vitro survival of the cells which grow in the absence of the endogenous extracellular matrix to which cells do attach in vivo.
In line to what we and others have previously observed in small interfering RNA and gene knock-out studies 13,19,22,44 cells expressing reduced amounts of CK2α also displayed reduced levels of CK2β suggesting that the contribution of the latter to the reported effects needs to be addressed in the future. Based on this, we also anticipate that a complete absence of CK2 kinase activity (i.e. double CK2α-and CK2α'-ablated mice) will have a more profound effect on cell proliferation, and experiments to test this hypothesis are under way.
Taking all these together, our data uncover a novel role of CK2α in the regulation of chromosomal DNA replication initiation, and propose that MCM helicases deregulation could be an important contributor to the observed reduction in cell proliferation in CK2α-deficient cells. We cannot, however, exclude that post-translational modifications of CK2α target proteins and/or additional genes identified in our genome wide expression analysis could contribute to the regulation of cell division.
The down-regulation of CK2α was achieved with the transduction of shRNA constructs and the transfection of small interfering RNAs, respectively. We found that the overlap of the up-or down-regulated genes in both systems was not significantly high, however, irrespective of the strategy pursued, we observed that a large number of significantly down-regulated transcripts were in common between the two systems. In this respect, although the goal is the same (i.e. the down-regulation of CK2α), the way this result is obtained differs with the two approaches. The siRNA-mediated approach involves transfection of a certain number of cells and is dependent on the variable efficiency of siRNA up-take. The lentivirus shRNA-knockdown system should, theoretically, affect all cells with respect to the silencing of a specific gene product. In the siRNA-based method, a certain amount of cells did not take up the siRNA molecules consequently; CK2α was not down-regulated in all cells. Because of this, one could not expect a perfect overlapping of the results obtained with the two systems emphasizing the importance to apply different approaches and validate significant results as we have performed in this study. Our global analysis of gene expression identified 770 significantly down-regulated genes (Fig. 4) including not only genes coding for proteins regulating DNA synthesis or DNA replication initiation but also for proteins directly regulating cell cycle phase transitions such as cyclin-dependent kinases and cyclin proteins. Furthermore, we cannot rule out that the decreased proliferation rate seen in cells depleted of CK2α might result from induction of the differentiation program. In this respect, Kankeu et al., carried out a proteomic analysis monitoring the changes in protein expression upon differentiation of cardiac myoblasts into cardiomyocyte-like cells and reported evidence that the proteins forming the MCM complex are significantly down-regulated in the differentiated cells 45 .
Future experiments should address the impact of miss-regulation or destabilization of the MCM complex and the mechanisms of reduced expression levels of MCMs in cells lacking CK2α. This is important, since mutations in MCMs are responsible for a number of diseases, including cancer, and malformations 29,46,47 . It will also be essential to perform rescue experiments with combinations of the MCM genes to address the importance of MCM proteins with respect to cell proliferation in vivo in the context of CK2α deregulation. Finally, it will also be important to generate tissue-specific CK2α knockout mice to further elucidate the function of this enzyme in different organs and define the pathways that lead to MCM deregulation. This knowledge will be pivotal for progressing our understanding of the functional specialization of protein kinase CK2α during embryonic development and in adulthood. www.nature.com/scientificreports www.nature.com/scientificreports/ supplemented with 10% fetal bovine serum (FBS, Biochrom AG, Berlin, Germany). H9c-2 cells were passaged before reaching confluence following guidelines to prevent their differentiation. Cells were treated with aphidicolin and doxycycline as indicated in the figure legends (both reagents from Sigma-Aldrich, Brøndby, Denmark). Down-regulation of protein expression was carried out by RNA interference as previously described 24 . Sets of four small interfering RNA duplexes (ON-TARGET plus SMART pools, Dharmacon, Lafayette, CO, USA) directed against CK2α and CK2α', respectively, were used. Synchronization of cells at G0/G1 was obtained by growing them in the presence of 0.1% serum for 48 h. After that, the cell cycle was resumed by adding complete growth medium and cells were harvested at different time points as indicated in the figure.

Methods
Gene expression silencing in H9c-2 myoblasts. The establishment of a cell line expressing shRNA for the inducible down-regulation of CK2α under the control of doxycycline was carried out with the Thermo Scientific SMARTchoice inducible shRNA kit following the manufacturer's instructions (Thermo Scientific, Rockford, IL, USA). The SMARTchoice inducible shRNA vector consists of an inducible promoter with tetracycline response element (P TRE3G ), activated by the Tet-On 3G protein in the presence of doxycycline (Dox). The SMARTvector contains a turbo-GFP (tGFP) reporter gene and a universal scaffold carrying shRNA directed against the CK2α sequence GAA.TTA.GAT.CCA.CGT.TTCA. Initially, general transduction conditions and the functional titer of viral particles were optimized, according to the manufacturer's recommendations. Cells were treated with optimal viral titer in transduction medium containing 10% serum and 10 µg/ml polybrene for 20 h followed by addition of normal growth medium for further 24 h. Transduced cells were selected with 0.3 µg/ml puromycin for three days, re-seeded as single cells and allowed to grow until each clone contained 30 to 40 cells. The clones were analyzed by Western blot for the efficient down-regulation of CK2α following treatment with 1 µg/ml doxycycline (Dox). Analysis of green fluorescence-emitting cells was done on a FACSCalibur flow cytometer and data acquisition was carried out with Cell Quest Pro Analysis software (BD Biosciences, Franklin Lakes, New Jersey, USA). Cell pictures were taken with a Leica DMRBE fluorescence microscope equipped with a DFC 420 C camera and Leica Application Suite V 3.3.0 software (Leica Microsystem, Wetzler, Germany).

Preparation of whole cell lysate, Western blot analysis and antibodies. Cells were harvested
and further processed for Western blot analysis as previously described 19 . The following primary antibodies were employed: mouse monoclonal anti-β-actin (Sigma-Aldrich); goat polyclonal anti-MCM3 and goat polyclonal anti-MCM6 (all from Santa Cruz Biotechnology, Heidelberg, Germany); rabbit monoclonal anti-cyclin E1, rabbit monoclonal anti-MCM2, rabbit monoclonal anti-MCM4, rabbit monoclonal anti-MCM7 and rabbit polyclonal anti-P-PTEN (S380/T382/383, all from Cell Signalling Technology, MA, USA); rabbit polyclonal anti-PTEN (Upstate, Lake Placid, NY, USA). Rabbit polyclonal anti-CK2α' was obtained by immunizing rabbits with a specific peptide sequence of human CK2α' (i.e. SQPCADNAVLSSGTAAR). Rabbit polyclonal anti-CK2α was obtained by immunizing rabbits against the human full-length protein sequence. Mouse monoclonal anti-CK2α/α' and mouse monoclonal CK2β were from KinaseDetect Aps, Odense, Denmark.
Cell cycle analysis. Cell cycle analysis and determination of cell death were carried out by propidium iodide staining and flow cytometry essentially as previously described 22 . The analysis of cells was carried out with a FACSCalibur flow cytometer (BD Biosciences). Acquired data were processed by Cell Quest Pro Analysis software (BD Biosciences). For each measurement, 10,000 events were analyzed. Flow cytometry of trypsin-digested mouse embryo cells was performed as described in 48 . Cells were analyzed using a FACScan flow cytometer (BD Biosciences) and data were processed using Cell Quest Pro analysis software (BD Biosciences).
Determination of cell proliferation. Cell proliferation was determined using the BrdU Cell proliferation Assay (Merck-Millipore, Hellerup, Denmark) following the manufacturer's instructions. In brief, cells were incubated with BrdU for 8 h. After fixation and denaturation, cells were labeled with anti-BrdU antibody for 1 h and subsequently with goat anti-mouse IgG HRP-conjugated for 30 min. Conjugates were visualized by adding HRP substrate solution. Absorbance was measured using a spectrofotometric plate reader at dual wavelengths of 450-595 nm. Alternatively, cells were harvested by trypsinization and counted every 24 h for up to 144 h with a Neubauer improved counting chamber.

RNA-Seq library preparation, sequencing and data analysis. Total RNA from cells was extracted
using Isol-RNA lysis reagent (AH Diagnostics, Aarhus, Denmark) and chloroform followed by precipitation in isopropanol. RNA concentration, purity and integrity were analyzed applying Nanodrop and Agilent 2100 Bioanalyzer RNA 6000 nano kit (Agilent Technologies, Inc., Santa Clara, CA, USA). Only RNA with RIN ≥8.0 and a 28 s/18 s ratio of approx. 1.8 was used for sample preparation. Processing of 500 ng RNA samples for library construction was essentially carried out as described in 49 using the TruSeq Total RNA LT Sample Prep kit, Set A (Illumina) and following the manufacturer's instructions (Illumina TruSeq Stranded Total RNA sample preparation guide). Amplified cDNA libraries were validated in regard to size by Agilent 2100 Bioanalyzer using a DNA 1000 kit from Agilent Technologies and concentration by qPCR using the KaPa Library quantification Kits (KaPa Biosystems, Wilmington, MA, USA). 15 pM denatured libraries were loaded on the flow cell for cluster formation by bridge amplification cycles followed by paired-end 100 bp sequencing on an Illumina HiSeq1500. For data analysis, samples were first trimmed to remove TruSeq adapters using cutadapt 50 and, subsequently, mapped to the rat genome assembly version rnor6 using STAR 51 version 2.5.0c with junction annotation from Ensembl ver 84 52 . Subsequently, gene counts were obtained using the same Ensembl reference and HTSeq 53 , and gene expression analyzed with DESeq2 54 using cqn 55 derived normalization factors. Pathway analysis was performed using GAGE 56 .