Functional impact of Aurora A-mediated phosphorylation of HP1γ at serine 83 during cell cycle progression
© Grzenda et al.; licensee BioMed Central Ltd. 2013
Received: 14 February 2013
Accepted: 14 June 2013
Published: 5 July 2013
Previous elegant studies performed in the fission yeast Schizosaccharomyces pombe have identified a requirement for heterochromatin protein 1 (HP1) for spindle pole formation and appropriate cell division. In mammalian cells, HP1γ has been implicated in both somatic and germ cell proliferation. High levels of HP1γ protein associate with enhanced cell proliferation and oncogenesis, while its genetic inactivation results in meiotic and mitotic failure. However, the regulation of HP1γ by kinases, critical for supporting mitotic progression, remains to be fully characterized.
We report for the first time that during mitotic cell division, HP1γ colocalizes and is phosphorylated at serine 83 (Ser83) in G2/M phase by Aurora A. Since Aurora A regulates both cell proliferation and mitotic aberrations, we evaluated the role of HP1γ in the regulation of these phenomena using siRNA-mediated knockdown, as well as phosphomimetic and nonphosphorylatable site-directed mutants. We found that genetic downregulation of HP1γ, which decreases the levels of phosphorylation of HP1γ at Ser83 (P-Ser83-HP1γ), results in mitotic aberrations that can be rescued by reintroducing wild type HP1γ, but not the nonphosphorylatable S83A-HP1γ mutant. In addition, proliferation assays showed that the phosphomimetic S83D-HP1γ increases 5-ethynyl-2´-deoxyuridine (EdU) incorporation, whereas the nonphosphorylatable S83A-HP1γ mutant abrogates this effect. Genome-wide expression profiling revealed that the effects of these mutants on mitotic functions are congruently reflected in G2/M gene expression networks in a manner that mimics the on and off states for P-Ser83-HP1γ.
This is the first description of a mitotic Aurora A-HP1γ pathway, whose integrity is necessary for the execution of proper somatic cell division, providing insight into specific types of posttranslational modifications that associate to distinct functional outcomes of this important chromatin protein.
KeywordsHeterochromatin protein 1 (HP1) Mitosis Aurora kinase Epigenetics Spindle pole Centrosome
Heterochromatin protein 1 (HP1), the reader of histone H3 lysine 9 methylation (H3K9me), was originally discovered through studies in Drosophila melanogaster of mosaic gene silencing, known as position effect variegation[1, 2]. In human and other mammalian cells, the three mammalian HP1 isoforms, HP1α, HP1β and HP1γ, have been well-studied for their localization, as well as their roles within the heterochromatic regions that associate with gene silencing. However, subsequent investigations have made it increasingly unmistakable that HP1 proteins not only localize to heterochromatic regions but also euchromatic regions[3, 4]. These proteins are involved in diverse cellular processes, ranging from chromatin modification and epigenetic gene silencing to replication and DNA repair to nuclear architecture and chromosomal stability[3, 4]. Moreover, HP1 proteins respond to a diversity of signaling pathways and acquire various posttranslational modifications, which impact on their function[5–9]. We have previously reported that, during interphase, phosphorylation of HP1γ at serine 83 (P-Ser83-HP1γ) via the cAMP-protein kinase A (PKA) pathway upon activation of cell surface receptors relocates this protein to euchromatin, where it plays a role in transcriptional elongation. Thus, it is essential to define HP1-mediated pathways to map useful networks of membrane-to-chromatin signaling cascades for better understanding of the regulation of important cellular processes.
Ample evidence indicates that HP1γ is important during both somatic and germ cell proliferation. Indeed, high levels of HP1γ protein associate with enhanced somatic and meiotic cell proliferation. Genetic inactivation of HP1γ results in both meiotic and mitotic failure[11, 12]. Studies in primordial germ cells demonstrate that loss of HP1γ also reduces their cell number through impaired cell cycle progression. However, the responsible molecular mechanisms that link this vital biological process to the functional regulation of HP1γ remain unknown.
Earlier investigations have found that HP1γ is phosphorylated throughout the cell cycle and, in particular, hyperphosphorylated in mitosis. In the current study, we report a novel pathway, whereby HP1γ is regulated by mitotic kinases, in particular, Aurora kinase A, a master regulator of mitotic transitions. We demonstrate that HP1γ is phosphorylated at serine 83 (Ser83) in G2/M where it colocalizes with Aurora A kinase, and its mitotic targets, cyclin B1, cyclin B2 and cyclin-dependent kinase 1 (CDK1) during cell division. HP1γ is phosphorylated at Ser83 by Aurora A in vitro and in cells. In addition, siRNA-mediated knockdown of HP1γ leads to a decrease of P-Ser83-HP1γ accompanied by mitotic aberrations. Notably, reintroduction of wild type HP1γ rescues, to a significant extent, these abnormal mitotic effects, while the nonphosphorylatable S83A-HP1γ mutant is unable to rescue this consequence of HP1γ knockdown. Congruent with these functions, phosphomimetic S83D-HP1γ results in an increase of cell proliferation, whereas the nonphosphorylatable S83A-HP1γ mutant abrogates this effect. In addition, overexpression of either the S83A-HP1γ or S83D-HP1γ mutant supports this effect in resultant cell cycle-related gene expression networks. Thus, together, these results reveal that a novel Aurora A-HP1γ pathway targeting Ser83 phosphorylation is necessary for the proper execution of cell division, thereby extending our knowledge of the biochemical and cell biological function of this important chromatin protein.
HP1γ is phosphorylated at the G2/M phase of the cell cycle
HP1γ is phosphorylated at G2/M by Aurora A
P-Ser83-HP1γ is required for normal mitotic function
P-Ser83-HP1γ status affects cell proliferation and mitotic gene expression networks
We next investigated whether the changes observed in EdU incorporation by both phosphomimetic and nonphosphorylatable Ser83-HP1γ mutants were accompanied by changes in other biochemical surrogates for cell cycle progression, such as known mitotic gene networks. For this purpose, we performed a genome-wide query using Affymetrix (Santa Clara, CA, USA) profiles as transcriptional readouts of their effects. Hierarchical clustering of targets significantly altered by HP1γ (526 targets), S83A-HP1γ (492 targets) or S83D-HP1γ (1,727 targets) overexpression demonstrated that gene networks modulated by HP1γ experienced deregulation in the presence of the Ser83 mutation, indicating dependence of these processes on regulation of Ser83 phosphorylation (Figure 6B). Based on Euclidean distance calculation and the resulting dendrogram, both control and nonphosphorylatable S83A-HP1γ mutant samples were statistically the most similar (Figure 6B). The fact that the EV and the S83A-HP1γ mutant possessed the closest relationship suggested that the latter worked predominantly as either an inactive or dominant negative mutant. However, the phosphomimetic S83D-HP1γ mutant, for the most part, reversed the effect of the S83A-HP1γ mutant, suggesting that it likely worked in a constitutively active manner thereby mimicking Aurora A-mediated Ser83 phosphorylation. Pathway-specific RT-PCR was used to validate a subset of significant targets (Additional file2: Table S1). These experiments revealed that HP1γ and its phosphorylated form have the ability to change the levels of transcripts related to mitosis.
Gene Ontology (GO) ANOVA analysis was utilized to probe for differentially expressed functional groupings of genes (Figure 6C). Overall, HP1γ overexpression resulted in significant enrichment of targets related to regulation of cellular proliferation, cell division, and mitosis (P <0.05). S83A-HP1γ mutant overexpression yielded differential expression in targets related to protein localization to the chromosome, regulation of the S phase of the mitotic cell cycle and regulation of the G2/M transition of mitotic cell cycle. S83D-HP1γ mutant overexpression showed significant alteration in genes related to the regulation of the mitotic cell cycle, regulation of the G2/M anaphase-promoting complex, maintenance of centrosome location and spindle pole structure, among others. Consequently, from these data, we conclude that disruption of phosphorylation status of HP1γ has diverse effects on multiple aspects of the mitotic cell cycle, which is congruent with its cell cycle-associated phosphorylation pattern (Figures 1 and2) indicating a pervasive role of the regulation of HP1γ in cell division.
Interestingly, previous studies have shown that depletion of HP1γ in primordial germ cells reduces their number as a result of impaired cell cycle progression. Comparison of our expression dataset with a published dataset in primordial germ cells revealed that the expression of the nonphosphorylatable S83A-HP1γ mutant displayed a highly similar pattern as HP1γ depletion, including targets related to cell cycle, proliferation and growth. This ability of the S83A-HP1γ mutation to mimic conditions of absolute HP1γ depletion at the level of gene expression networks, combined with the inability of the S83A-HP1γ mutant to rescue the mitotic defects observed with HP1γ knockdown, indicates that posttranslational modification of this residue is needed for proper progression through mitosis. Furthermore, it may be concluded from our genome-wide analysis that HP1γ participates in the regulation of processes, which support proper cell division and proliferation through phosphorylation-dependent and phosphorylation-independent mechanisms.
Based on the current study, our demonstration that HP1γ, a well-known epigenetic regulator, undergoes robust phosphorylation at Ser83 in G2/M has significant biological relevance and deserves careful consideration. Previous studies demonstrating that HP1 proteins are ejected from chromosomes during mitosis[28, 29] led to the assumption that this protein is not involved in the regulation of this process, even though it is highly express in rapidly dividing cancer cells. In this regard, the current study reveals that, during G2/M, an extrachromosomal subpopulation of HP1γ, P-Ser83-HP1γ, localizes with γ-tubulin, Aurora A kinase and other mitotic targets, including cyclin B1, cyclin B2 and CDK1, at the spindle poles. Thus, this data demonstrates for the first time that, in spite of its ejection from chromosomes, HP1γ does not disappear during mitosis, but rather relocates to organelles, known for enrichment in cell cycle regulators, where it undergoes G2/M-specific phosphorylation at Ser83 by Aurora A. In addition, the colocalization and coupling of Aurora A to HP1γ in cell cycle regulation is reconstituted in time and space in each cell cycle.
Examination of the effect of the related kinase, Aurora B, demonstrates that this enzyme can phosphorylate the Ser83 site in vitro. However, siRNA and dominant negative experiments demonstrate that Aurora B was not as robust as Aurora A on modulating levels of P-Ser83-HP1γ in cells. Treatment of cells with the Aurora B inhibitor, hesperidin, does not impair P-Ser83-HP1γ and, more importantly, Aurora B does not localize with P-Ser83-HP1γ in mitotic cells. These results reveal a significant level of specificity for these kinases in the phosphorylation of HP1 proteins.
We found that HP1γ, though ejected from chromosomes by the previously described Aurora-mediated P-Ser10-H3[28, 29], remains tightly associated to a mitotic organelle which is rich in cell cycle regulators. This reveals the existence of coupled mechanisms of ejection and relocalization of HP1γ, which ultimately has significant consequences for the regulation of cell division. Both steps involved in this process, H3 and HP1γ phosphorylation, are mediated by Aurora kinases. Thus, it is most likely that one function of Auroras has evolved, in part, to secure that epigenetic regulators are turned on and off during cell division in a highly synchronized manner, to achieve the proper transfer of genetic-epigenetic material through generations. Interestingly, although HP1 proteins themselves have not been previously observed at the centrosome/spindle pole, several HP1-interacting proteins are known to reside in this cell compartment. For example, a subpopulation of origin recognition complex 2 (Orc2) protein has been localized to centrosomes. However, contrary to the Aurora A-cyclin B-CDK1 pathway, which links the phosphorylation of HP1γ at the spindle during G2/M transition, Orc2 associates with HP1 only in the population that is tightly bound to heterochromatin in G1 and early S phase. In addition, immunoprecipitation of Orc2 shows specific interaction with HP1α and HP1β, but not HP1γ, the HP1 protein studied here. Since posttranslational modifications of HP1 were not considered in the Orc2 experiments, it remains possible that subpopulations of distinct posttranslationally modified HP1 proteins, such as P-Ser83-HP1γ, which cannot be detected with pan-HP1 antibodies, also interact with Orc2. It is not likely, however, that Orc2 is responsible for recruitment of HP1γ to this cell compartment, given that Orc2 is localized there throughout the entire cell cycle. Nevertheless, our results demonstrate a high degree of selectivity for HP1γ to work with certain regulatory enzymes (kinases) to maintain mitotic functions.
Previous studies have shown that disruption of G9a, one of the histone methyltransferases responsible for the histone mark recognized and bound by HP1, H3 lysine 9, results in chromosome instability along with centrosome abnormalities. In addition to creating the mark to which HP1 binds, G9a localizes with HP1α and HP1γ, which is dependent upon its own automethylation, and HP1γ has been shown to specifically form complexes with G9a in the context of the E2F-6 gene silencing complex. Interestingly, in meiosis cell division during gamete production, HP1γ and G9a are proposed to form an axis that is responsible for retaining centromeric regions of unpaired homologous chromosomes in close alignment, and facilitating progression of their pairing in early meiotic prophase. In fact, HP1γ-deficient mouse spermatocytes undergo meiotic catastrophe. An important observation of our studies is that siRNA-mediated knockdown of HP1γ leads to a decrease of P-Ser83-HP1γ accompanied by mitotic aberrations. While reintroduction of wild type HP1γ rescues, to a significant extent, these abnormal mitotic effects, the nonphosphorylatable S83A-HP1γ mutant is unable to rescue this consequence of HP1γ knockdown, highlighting the importance of Ser83 modification for this function. Moreover, the S83D-HP1γ mutant that mimics Aurora A phosphorylation facilitates cell proliferation, whereas the nonphosphorylatable S83A-HP1γ mutant inhibits this process. Therefore, it is tempting to speculate whether modifications of HP1 influence interactions with G9a and whether these proteins function together in regulating proper cell division. Indeed, additional studies using model organisms support that the function described here for human HP1 proteins is conserved. In Schizosaccharomyces pombe, the HP1 homologue, Swi6, is required to preserve genomic integrity and proper segregation of chromosomes during mitosis. Impaired Swi6 function leads to mitotic alterations that cause severe growth alterations. Furthermore, the HP1-like protein in Dictyostelium discoideum, AX4 chromo domain-containing protein (hcpA), which displays 79% similarity to human HP1γ, colocalizes with electron-dense structures at the nuclear periphery that are compatible with pericentrosomal material. Overexpression of this protein causes growth defects that are accompanied by an increase in the frequency of atypical anaphase bridges. Genetic studies in Drosophila have demonstrated that mutations in the HP1 protein cause defective chromosome segregation[36, 37]. Thus, in combination with this data, the studies described here indicate that HP1 proteins have evolved to support cell division in organisms ranging from fission yeast to humans.
Congruent with our results, previous experiments have defined a role for HP1γ in human diseases that are characterized by abnormal cell proliferation. High levels of HP1γ have been observed in several cancer types, including esophageal, breast, colon, lung and cervical cancer, the cell model used here. In addition, siRNA-mediated knockdown of HP1γ expression inhibits cervical cancer cell proliferation. Of note, Aurora A, the kinase identified in this study as responsible for P-Ser83-HP1γ at G2/M, is amplified and overexpressed in cervical cancer, which induces centrosome amplification, aneuploidy and transformation. Cervical cancer patients with high Aurora A expression correlate with a poorer disease-free survival and overall survival rates than patients with low Aurora A expression, indicating that this protein could be used as a prognostic marker. Based on the current study, the high levels of both HP1γ and Aurora kinases in cervical cancer cells would suggest that there is a resultant increase in P-Ser83-HP1γ. Thus, targeting this pathway would affect P-Ser83-HP1γ-mediated cell proliferation, in addition to other downstream Aurora effectors. In fact, Aurora kinase inhibitors have been shown to suppress proliferation of cervical cancer cells and enhance chemosensitivity[40, 41], suggesting that targeting Aurora in combination with the HP1-histone methyltransferase pathway may be a beneficial therapy in these patients.
In summary, the current study identifies a novel Aurora-HP1γ pathway that involves P-Ser83-HP1γ by Aurora A in G2/M and localization of this HP1γ subpopulation to the spindle pole, which is necessary for proper cell division. Combined, these results constitute robust evidence that P-Ser83-HP1γ plays a role in mitosis and bears importance for understanding impairments, which have been shown to be characterized by abnormally high levels of HP1γ and Aurora kinase activity, including cancer. Our results also suggest a teleological interpretation, namely that certain regulators of chromatin dynamics and transcription, such as HP1γ, may undergo functional pressures (for example Aurora A phosphorylation) to maintain the integrity of cell division so that their own epigenetic inheritance is reproducible from cell generation to cell generation.
Cell lines, reagents and cell treatments
Cell lines were obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA) and maintained according to the manufacturer’s protocol. The human LX2 cell line was obtained as a generous gift from Dr Steve Freeman (Mount Sinai, NY, USA). Roscovitine (Sigma-Aldrich, St Louis, MO, USA) treatment was added at increasing concentrations (0, 5, 10 and 20 μM) for 8 hours, and lysates were harvested. Cells were treated with 3 μg/ml aphidicolin or 2 μg/ml nocodazole (both from EMD Millipore, Billerica, MA, USA) for 16 hours to arrest at G1/S and G2/M, respectively. Control cells were treated with vehicle, dimethyl sulfoxide (DMSO). HeLa cells were synchronized by double thymidine block. Thymidine (2 mM, Sigma-Aldrich) was added to asynchronous cells for 18 hours. Cells were subsequently released for 9 hours in regular growth media prior to the second thymidine (2 mM) block. After 17 hours, cells were released for the thymidine block and lysates were collected at the indicated time points. KT5720 was obtained from EMD Millipore. MLN8237 and hesperidin were purchased from Selleckchem (Houston, TX, USA). For hesperidin treatment, HeLa cells were arrested in mitosis by treatment with nocodazole for 16 hours. Arrested cells were treated with 200 nM hesperidin for the indicated times in the presence of 10 μM of the proteasome inhibitor MG132 (Sigma-Aldrich) to prevent mitotic exit.
Plasmids, siRNA and recombinant adenovirus
Standard molecular biology techniques were used to clone HP1γ into the pGEX and Ad5CMV vectors. For HP1γ-specific transient shRNA-mediated knockdown, complementary oligonucleotides were synthesized for the target sequence (GCAAATCAAAGAAGAAAAG), annealed and ligated into the pCMS3 vector (kindly provided by Dr Daniel Billadeau, Mayo Clinic, Rochester, MN, USA). For stable shRNA-mediated HP1γ knockdown, control or HP1γ-specific shRNA lentiviral particles (Santa Cruz Biotechnology, Inc, Santa Cruz, CA, USA) were used to infect cells according to the manufacturer’s protocol, followed by puromycin selection (2 μg/ml). Myc-tagged wild type and dominant negative constructs for Aurora A and Aurora B were a kind gift from Dr Paolp Sassone-Corsi. S83A-HP1γ and S83D-HP1γ mutations were obtained using the QuickChange Site-Directed Mutagenesis Kit, as suggested by the manufacturer (Agilent Technologies, Inc, Santa Clara, CA, USA). All constructs were verified by sequencing at the Molecular Biology Core at Mayo Clinic, Rochester, MN, USA. Aurora A (AURKA) and Aurora B (AURKB) Silencer validated siRNAs were purchased from Ambion-Life Technologies (Carlsbad, CA, USA). Epitope-tagged (6xHis-Xpress) HP1γ, S83A-HP1γ and S83D-HP1γ, as well as EV (Ad5CMV), were generated as recombinant adenovirus in collaboration with the Gene Transfer Vector Core at the University of Iowa, IA, USA.
Western blot analysis
Samples were run on 4 to 20% gradient SDS-PAGE gels (Lonza, Walkersville, MD, USA) or 12% SDS-PAGE gels and electroblotted onto polyvinylidene difluoride (PVDF) membranes (EMD Millipore). The membranes were blocked in 5% BSA in tris-buffered saline Tween-20 (TBST) for 1 hour at room temperature. The blots were incubated for 2 hours at room temperature or overnight at 4°C with primary antibody (P-Ser83-HP1γ, 1:1,000; HP1γ, 1:1,000; and P-Ser10-H3, 1:5,000 (all from EMD Millipore); Aurora A, 1:1,000 (BD Biosciences Pharmingen, San Diego, CA, USA); Aurora B, 1:500; cyclin B1, 1:1,000; cyclin B2, 1:1,000; and CDK1, 1:1,000 (Abcam, Cambridge, MA, USA); β-actin, 1:1,000; and α-tubulin, 1:1,000 (Sigma-Aldrich); c-Myc (9E10) for Myc-tagged proteins, 1:1,000 (Thermo Scientific, Rockford, IL, USA); and OMNI D8 for His-tagged proteins, 1:1,000 (Santa Cruz Biotechnology)). After repeated washes in TBST, horseradish peroxidase (HRP)-conjugated anti-rabbit or mouse IgG secondary antibody (1:5,000) was added for 1 hour at room temperature. Blots were developed by Pierce ECL chemiluminescent substrate (Thermo Scientific).
Immunofluorescence and confocal microscopy
Immunofluorescence and confocal microscopy were performed as previously described. The primary antibodies were used at the following dilutions: P-Ser83-HP1γ, 1:200; and γ-tubulin, 1:500 (Sigma-Aldrich); Aurora A, 1:50; and Aurora B, 1:50 (BD Biosciences Pharmingen); cyclin B1, 1:500; cyclin B2, 1:100; and CDK1, 1:40 (Abcam); and cyclin D3, 1:200 (Cell Signaling Technology, Danvers, MA, USA). For localization of P-Ser83-HP1γ during S-phase, EdU incorporation was combined with immunofluorescence. Prior to fixation, cells were incubated for 30 minutes in media containing 10 uM EdU. Subsequently, cells were processed for immunofluorescence, followed by EdU labeling using the Click-iT EdU Imaging Assay Kit (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol. For mitotic aberrations, spindle poles were labeled by immunofluorescence with γ-tubulin and counterstained with 4',6-diamidino-2-phenylindole (DAPI) containing mounting media (Vector Laboratories, Burlingame, CA, USA). For each condition, at least 200 mitotic cells were analyzed to quantify mitotic aberrations.
GST fusion protein purification and in vitro kinase assays
GST fusion protein purification was done as previously described. For Aurora A and Aurora B in vitro kinase assays, HP1 fusion proteins (10 μg) were incubated with recombinant kinases (EMD Millipore) and 10 mM ATP (Sigma-Aldrich) for 10 minutes at 30°C, in either the supplied buffer (Aurora A) or buffer containing 50 mM Tris pH 7.5, 0.1 mM ethylene glycol tetraacetic acid (EGTA), and 15 mM dithiothreitol (DTT, Aurora B). Kinase reactions were terminated by the addition of SDS loading dye and then resolved by western blot as described above.
Cell proliferation assay
Cell proliferation was measured by EdU incorporation using both fluorescence-activated cell sorting (FACS) and microscopy. Cells were infected with adenovirus carrying control, HP1γ, S83A-HP1γ or S83D-HP1γ vectors. Forty-eight hours post-plating, cells were pulsed with 10 μM EdU (Invitrogen) for 1 hour. Subsequently, cells were processed using the Click-iT EdU Flow Cytometry or Imaging Assay Kits (Invitrogen) according to the manufacturer’s protocols. EdU incorporation was measured by FACS analysis at the Mayo Flow Cytometry Research Core Facility, Rochester, MN, USA, or confocal microscopy. Each experiment was performed at least five different times in triplicate, expressed as means with standard error of mean (SEM) and statistical analyses were performed using unpaired t-test.
Gene expression profiling, microarray analysis
Global gene expression profiling was carried out at the Microarrays Facility of the Research Center of Laval University, CRCHUL, QC, Canada, utilizing the Affymetrix Human Gene 1.0 ST arrays (28,869 well-annotated genes and 764,885 distinct probes). Intensity files were generated by Affymetrix GCS 3000 7G and the GeneChip Operating Software (Affymetrix, Santa Clara, CA, USA). Data analysis, background subtraction and intensity normalization was performed using robust multi-array analysis (RMA). Genes that were differentially expressed along with false discovery rate were estimated from t-test (>0.005) and corrected using Bayesian approach[44, 45]. Data analysis, hierarchical clustering and ontology were performed with the oneChannelGUI to extend affylmGUI graphical interface capabilities, and Partek Genomics Suite, version 6.5 (Partek Inc, St Louis, MO, USA) with ANOVA analysis. Final fold changes were calculated as x = 2^log2value. Probes with P value <0.05 and fold change ± 2.2 among HP1γ versus EV, S83A-HP1γ versus EV, and S83D-HP1γ versus EV were selected for further analysis. For GO ANOVA, a minimum threshold of three genes and P <0.05 was used to identify significant functional groups. To validate the Affymetrix microarray, targets with significant alteration (P <0.05) were compared to the real-time data using an arbitrary cutoff of ± 2.2 fold change compared to EV control.
analysis of variance
American Type Culture Collection
bovine serum albumin
cyclin-dependent kinase 1
Chinese hamster ovary
Centre de Recherche du Centre Hospitalier de l'Université Laval
ethylene glycol tetraacetic acid
fluorescence-activated cell sorting
graphical user interface
histone H3 lysine 9 methylation
Dictyostelium discoideum, AX4 chromo domain-containing protein
heterochromatin protein 1
origin recognition complex subunit 2
protein kinase A
phosphorylation of histone H3 at serine 10
phosphorylation of HP1γ at serine 83
phosphorylation of Aurora A at threonine 288
robust multi-array analysis
reverse transcriptase polymerase chain reaction
standard error of mean
shRNA knockdown of HP1γ
short hairpin RNA
small interfering RNA
tris-buffered saline Tween-20.
This work was supported by funding from the Fraternal Order of Eagles and a Career Development Award from the Mayo Clinic SPORE in Pancreatic Cancer (P50 CA102701, both to GL), as well as the National Institutes of Health (grant DK52913 to RU and T32CA148073 to AG), the Mayo Clinic Center for Cell Signaling in Gastroenterology (P30DK084567), and the Mayo Foundation. The authors would like to sincerely thank Dr Debora Bensi for technical assistance during the early development of this work, as well as Holger Dormann and Dr C David Allis for their generous contributions and helpful insights for the hesperidin P-Ser83-HP1γ experiments.
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