![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||
T. Geraldine Guerin-Peyrou,2,
Girdhar G. Sharma,1,
Changwon Park,1
Manjula Agarwal,1
Ramesh K. Ganju,3
Shruti Pandita,1
Kyunghee Choi,1
Saraswati Sukumar,4
Raj K. Pandita,1
Thomas Ludwig,2* and
Tej K. Pandita1*
Washington University School of Medicine, Saint Louis, Missouri 63108,1 Institute of Cancer Genetics, Columbia University, New York, New York 10032,2 Harvard Medical School, Boston, Massachusetts 02115,3 John Hopkins School of Medicine, Baltimore, Maryland 212314
Received 13 June 2007/ Returned for modification 16 July 2007/ Accepted 11 October 2007
 |
ABSTRACT |
|---|
 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Acetylation at K16 of histone H4 (H4K16ac) is a prevalent and reversible posttranslational chromatin modification in eukaryotes, and recent studies have highlighted its significance. Shogren-Knaak and coworkers have found that a single histone H4K16ac modification modulates both higher-order chromatin structure and functional interactions between a nonhistone protein and the chromatin fiber (39). Shia and coworkers have demonstrated that the presence of H4K16ac and H2A.Z synergistically prevent the ectopic propagation of heterochromatin in the subtelomeric regions of yeast (36). Furthermore, it is well understood that H4K16ac disrupts higher-order chromatin structure, changes the functional interactions between chromatin-associated proteins (39), and serves as a switch for altering chromatin from a repressive to a transcriptionally active state in yeast and humans (36). Interestingly, Dou and coworkers reported that MOF-mediated histone acetyltransferase activity, specific for H4K16, is required for optimal transcription activation on a chromatin template in vitro and in vivo (6). The loss of monoacetylation at lysine 16 of histone H4 specifically at nucleosome repetitive sequences is a common hallmark of tumor cell lines and human cancer where it is progressively lost from early preneoplastic stages to the most malignant stage (8). Furthermore, inhibition of SIRT1 deacetylase in breast and colon cancer cells has been shown to cause increased acetylation of H4K16 at two specific endogenous promoters that correlated with decreased proliferation (29). In contrast, mouse embryonic fibroblasts (MEFs) deficient for SIRT1, have dramatically increased resistance to replicative senescence (5). These results demand further evaluation of H4K16ac in tumor and normal cells.
Vigorous cellular proliferation is common and essential during both embryogenesis and oncogenesis. Therefore, these two processes may potentially have common chromatin modification signatures characteristic of their proliferation status. Recent studies have demonstrated that hMOF depletion, resulting in the loss of H4K16 acetylation correlates with a decrease in DNA damage-induced activation of ATM and prevents ATM from phosphorylating downstream effectors, such as p53 and CHK2 (12, 45). hMOF physically interacts with ATM and p53 (6, 12), and in vitro, hMOF binds to and acts synergistically with p53 to increase histone H4K16ac and target gene transcription (6). While H4K16ac is known to alter higher-order chromatin structure into a relaxed conformation (39), it is important to determine whether there is a correlation between this modification and cellular growth both during embryonic development and in the process of tumorigenesis.
In this study, we determined whether MOF, which specifically acetylates H4K16, is required for cell growth and proliferation during either embryogenesis or tumorigenesis and whether proliferating tumors and tumor-derived cell lines demonstrated loss of MOF and H4K16 acetylation. Specifically, we analyzed the impact of mammalian MOF expression on embryonic development and oncogenic transformation, two processes that rely on cellular proliferation. We examined MOF and H4K16ac levels in single-cell zygotes through blastocyst-stage embryos and found that ablation of MOF correlated with loss of histone H4K16ac and paralleled embryonic lethality and cell death. Conversely, MOF overexpression increased H4K16ac levels, which correlated with oncogenic transformation and tumor growth. All tumor cell lines examined expressed hMOF and H4K16ac. Strikingly, primary human cells immortalized with the catalytic unit of telomerase (hTERT) have higher levels of hMOF and H4K16ac than primary cells do.
|
MATERIALS AND METHODS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
|
flox/+/Rosa26creERT2/+ ES cells. Electroporation of these cells with the conditional targeting vector then gave rise to mMofflox/+/Rosa26creERT2/+ and mMofflox/flox/Rosa26creERT2/+ ES cell clones.
|
Generation of various mouse genotypes. All mice in this study have SV129 background. The Atm and p53 heterozygous mice have been described previously (33, 50). Atm–/– mice are infertile. Atm–/–/Mof+/– mice were generated by crossing male and female Atm+/–/Mof+/– genotypes. Offspring were genotyped at age 21 days by PCR-based assays using mouse tail DNA (50).
Cell lines. The cell lines used were human lymphoblastoid (GM07349, GM10860, GM10854, GM10863, GM10851, GM10850, and GM10847), human leukemia (HL60, K562, and Jurkat), mesothelial cancer lines (REN, LRK, MS1022H, H266, H2052, and H2452), lymphoma (Raji, U937, and HT), breast (MCF-7 and MCF-10A), colon (HCT-116 and RKO), lung (H1299), osteosarcoma (MG63) lymphoblastoid cell lines, and 293 and HeLa cells. All cells were maintained as described previously (26, 40). Primary fibroblast cells lines (GM5823, HFF, and BJ) and the establishment of their hTERT-derived counterparts were described previously (48). Mouse fibroblasts 435 (with ATM) and 743 (without ATM) cells have been described previously (10).
Survival assay. Cells were plated into 60-mm dishes in 5.0 ml of medium, incubated for 6 h, and subsequently exposed to ionizing radiation (IR) as described previously (25). The number of cells plated per dish was selected to ensure that about 50 colonies would survive treatment with a particular radiation dose. Cells were exposed to IR in a dose range of 0 to 6 Gy at room temperature. Cells were incubated for 12 or more days and then fixed in methanol-acetic acid (3:1) prior to staining with crystal violet. Only colonies containing at least 50 cells were counted.
Western blot analysis. Cell lysates for Western blot analysis were prepared as previously described (27). Antibodies used to detect MOF, histone H4, and acetylated H4K16 were previously described (12). Immunoblotting and detection of MOF, histone H4, and acetylated H4K16 were done according to previously described procedures (12, 27, 35).
Primary cell culture, establishment of cell lines, and growth assays. MEFs from mMof heterozygous intercrosses were isolated from embryonic day 13.5 (E13.5) embryos by standard procedures (15, 25). Cells were maintained in Dulbecco's modified Eagle's medium (DMEM) with 10% fetal bovine serum (FBS) and frozen at different passages for storage. Continuous passaging was utilized to allow for emergence of spontaneously immortalized clones from both wild-type and mMof heterozygous MEFs. These clones were positive for telomerase activity and were passaged for at least 100 population doublings.
For growth assays, MEFs were plated in 60-mm tissue culture dishes, and four dishes were counted on the indicated days. Four different passages (passages 1, 5, 12, and 19) were utilized in order to determine the effect of passage number on growth rate. To determine plating efficiency, MEFs were seeded at various densities, and after 15 days in culture, the colonies were fixed, stained, and counted.
Analysis of micronuclei and ratio of normochromatic to polychromatic erythrocytes. Micronucleus formation and the ratio of normochromatic to polychromatic erythrocytes were determined by previously described procedures (15, 23). Briefly, bone marrow smears from the mice were prepared, and the stained smears were examined to determine the incidence of micronucleated cells in 1,800 polychromatic erythrocytes and the ratio of normochromatic to polychromatic erythrocytes for each animal.
Transformation assay. The transformation assay was performed as described previously (9, 15). Exponentially growing subconfluent cells were trypsinized and plated 48 h prior to exposure to IR at a dose of 1 Gy. Immediately after irradiation, the cells were trypsinized and replated into 10-cm-diameter culture dishes at cell numbers estimated to result in either 300 viable cells per dish for the assay of neoplastic transformation or 30 viable cells for the cell survival assay. For the assay of neoplastic transformation, cells were grown in Eagle's basal medium supplemented with 10% heat-inactivated FBS, and the culture medium was changed at 12-day intervals during the 6- to 8-week incubation. Cells plated for cell survival determination were incubated as described above. At the end of 12 or 42 days of incubation, cells were fixed in formalin and stained with Giemsa. Cell survival was determined by the colony assay as described above, while neoplastic transformed foci II and III were identified according to the criteria of Reznikoff et al. (30, 31).
Reverse transcriptase PCR of hMOF mRNA. To compare hMOF mRNA levels in paired tumor and normal tissue samples, reverse transcriptase PCR was performed. mRNAs isolated from paired tumor and normal tissue samples were obtained from the Alvin J. Siteman Cancer Center of the Washington University School of Medicine, St. Louis, MO; the Sidney Kimmel Cancer Center at John Hopkins, Baltimore, MD; and the M.D. Anderson Cancer Center, Houston, TX. The isolated RNAs from tumor and normal tissues were treated with RNase-free DNase (Roche Diagnostic Corporation, Indianapolis, IN) (1 mg/ml) for 2 h at 37°C, followed by heat inactivation at 65°C for 10 min. The reverse transcriptase reaction mixture contained 1 mg/ml DNase-treated RNA, 0.25 mg/ml pdN6 random primers (Roche Diagnostic Corporation), first-strand buffer (GIBCO-BRL), 0.5 mM deoxynucleoside triphosphate (Roche Diagnostic Corporation, Indianapolis, IN), and 200 units Moloney murine leukemia virus reverse transcriptase (GIBCO-BRL), and the mixtures were incubated for 1 h at 37°C. PCR was performed by using gene-specific primers along with the primers for β-actin. cDNAs of paired samples (tumor and normal tissue) were analyzed with ABI Prism 7700 (Applied Biosystems) with SYBR green ROX mix (Abgene, United Kingdom). Each cDNA was analyzed in triplicate, and dihydrofolate reductase (DHFR) was used as an internal control for comparison of the expression status. The relative quantification of hMOF mRNA (compared to DHFR mRNA) was determined as described previously (34, 35). The primer oligonucleotide sequences used for hMOF and DHFR quantification are available upon request. The products of the reverse transcriptase PCRs were analyzed by gel electrophoresis, and they gave single bands of the appropriate amplicon size.
Mice and tumor irradiation. Eight-week-old NMRI nu/nu male mice were maintained in a pathogen-free mouse colony for the duration of experiments. Mice were randomly distributed four per cage, and each mouse was labeled with a code on the ear. Two million exponentially growing RKO cells with and without hMOF overexpression in a volume of 20 µl were injected (intramuscularly) in the right thigh flank, and the mice were checked daily for tumor appearance as described previously (32). Animals with a tumor of 8 mm in mean diameter were locally irradiated with a single dose of 25 Gy using a 8-MeV linear particle accelerator (dose rate of 1 Gy/min). For the regrowth of tumors, the previously described protocol was followed (32).
Chemotaxis assay. Migration assays were done in 24-well cell culture chambers with 8-µm-pore membranes. The lower surface of the membrane was precoated with fibronectin (25 µg/ml). Cells were resuspended in chemotaxis buffer (DMEM, 0.1% bovine serum albumin, 12 mM HEPES) at 2.5 x 106 cells/ml, and 150 µl of cells from each sample was loaded into the upper chamber. Chemotaxis buffer (0.6 ml) containing DMEM, 10% FBS, epidermal growth factor (EGF) (20 ng/ml), and insulin growth factor (IGF) (20 ng/ml) was dispensed into the lower chamber. The plates were incubated for 24 h at 37°C in 5% CO2. After incubation, the inserts were removed carefully, and the cells were fixed and stained using the HEMA 3 staining kit (Fisher Scientific Company, MI). Migrated cells were counted in a minimum of five high-power fields (magnification of x100).
|
RESULTS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
58 kDa, a molecular mass similar to that of human MOF protein (Fig. 1C) (12, 42). To determine the physiological significance of mammalian MOF, we disrupted the mMof gene by homologous recombination in embryonic stem cells (Fig. 2A) and generated germ line-transmitting chimeras. Heterozygous (mMof+/–) intercrosses yielded wild-type (mMof+/+) and heterozygous (mMof+/–) offspring pups but no homozygous mMof null mutant pups (Fig. 2B). The complete absence of homozygous Mof null mutants in all litters examined implied that Mof deficiency is lethal to embryos (Fig. 2B). We then determined the stage of lethality by analyzing embryos from mMof+/– intercrosses at different times postcoitum. In the litters of mice at E7.5 and E8.5 that we dissected, we found approximately 25% empty or necrotic deciduae (Fig. 2C), and all the morphological normal embryos were either wild type or heterozygous (Fig. 2D and data not shown). At E7.5, all the normal embryos (wild type and heterozygotes) had gastrulated, while the Mof null mutant deciduae contained clusters of cells within a small sac or some giant cells only (Fig. 2C), indicating that Mof-deficient embryos are capable of implantation but die prior to the onset of gastrulation.
The observation that no viable mMof–/– embryos are found at the E7.5 stage of development led us to investigate the expression and localization of MOF protein and H4K16ac status during early embryogenesis (single-cell zygote to preimplantation blastocyst) (Fig. 3A). PCR-based genotyping revealed that about 25% of all zygotes did not have the mMof gene (Fig. 3B). MOF protein localized throughout the entire zygotic cell (whether mMof+/+, mMof+/–, or mMof–/–), including the cytoplasmic and chromatin regions, indicating a strong maternal contribution of MOF. In the single-cell zygotes, both male and female pronuclei chromatin in the fertilized egg stained equally strongly for acetylated histone H4K16 (H4K16ac) (Fig. 3C, top panel). All zygotes were positive for MOF and H4K16ac staining; however, about 25% of the zygotes did not have the mMof gene (Fig. 3B). Through the first few cleavages, until the 16-cell stage embryo is reached, neither MOF nor H4K16ac immunofluorescence distinguishes mMof–/– embryos from the mMof+/+ or mMof+/– embryos (Fig. 3C and D). This is most likely due to the abundance of maternal MOF protein present in mMof–/– oocytes, which probably allowed for continued cleavage during early embryonic development and normal levels of H4K16ac in the chromatin. At the 32-cell stage, mMof+/+ or mMof+/– embryos have increasingly distinct nuclear signals for MOF (Fig. 3D, top panel). H4K16ac levels in Mof null mutants at this stage remained unaffected compared to the levels in heterozygous or wild-type embryos.
|
Inactivation of ATM and p53 does not rescue lethality of mMof deletion.
Prior studies have reported that depletion of hMOF resulted in a G2/M arrest (46) that could be partly rescued by inactivation of specific cell cycle checkpoint gene products. Therefore, we examined whether the delayed developmental progression and lethality of mMof–/– embryos could be rescued by inactivating genes such as ATM or p53, which are involved in cell cycle control (20, 22, 27, 38). Although knockdown of hMOF in human cells results in a G2/M checkpoint arrest (46), inactivation of ATM or p53 could not rescue or ameliorate the early embryonic lethality of mMof nullizygous embryo (data not shown). We determined the influence on growth due to mMof haploinsufficiency in Atm heterozygous and Atm null backgrounds by monitoring the health and survival of mMof+/+/Atm+/+ (n = 34), mMof+/–/Atm+/+ (n = 45), mMof+/+/Atm+/– (n = 30), mMof+/–/Atm+/– (n = 38), mMof+/+/Atm–/– (n = 24), and mMof+/–/Atm–/– (n = 19) mice over a period of 1 year (Fig. 4A). In contrast to 92% of mMof+/+/Atm+/+, mMof+/+/Atm+/–, or mMof+/–/Atm+/+ mice, only 76% mice with the mMof+/–/Atm+/– genotype remained healthy and survived the first year of life. mMof heterozygosity in Atm null background (mMof+/–/Atm–/–) resulted in a modest weight reduction and earlier death compared to Atm null mice (Fig. 4A and B). Although the differences in survival are modest, they are statistically significant (P < 0.05). Interestingly, mMof heterozygosity in a p53-deficient background (mMof+/–/p53–/–) resulted in death at an earlier age compared to p53 null mice (Fig. 4C and data not shown).
|
Thus, we designed a conditional mMof allele (Mofflox) by flanking exons 1 to 3 with loxP sites (Fig. 5A) and generated mMofflox/+ ES cells. To inactivate the conditional allele by cre-mediated recombination, we introduced the tamoxifen-inducible creERT2 fusion gene into the ROSA26 locus (ROSA26creERT2/+) using a knock-in strategy and subsequently generated mMofflox/+/Rosa26creERT2/+ and mMofflox/flox/Rosa26creERT2/+ ES cells. The phenotypic consequences of MOF deficiency were then determined by tamoxifen-induced expression of Cre recombinase and inactivation of the conditional Mofflox allele (Fig. 5A and B). Deletion of mMof resulted in the subsequent loss of MOF protein, loss of acetylation of histone H4 at K16 (Fig. 5C), and decreased cellular proliferation (Fig. 5D). Heterozygous mMof (mMof+/–) MEFs also had reduced levels of MOF (Fig. 6A) and demonstrated a modest increase in population doubling time (Fig. 6B), decreased IR-induced 
-H2AX focus formation (Fig. 6C), and a slight decrease in survival after IR exposure (Fig. 6D). The decreased survival in mMof+/– MEFs could be due to defective repair of IR-induced single- and double-strand DNA breaks. MOF, a chromatin-modifying factor, may have a critical role in the repair of DNA cross-links. We also compared the effect of mitomycin C-induced DNA cross-links on mMof +/+ and mMof +/– MEFs. Cell killing after mitomycin C treatment was higher in mMof+/– MEFs than in mMof+/+ MEFs (Fig. 6E).
|
-H2AX focus formation was also observed in heterozygous mMof cells compared to wild-type mMof cells. Interestingly, reduced levels of mMof increased genomic instability and decreased survival after IR or mitomycin C exposure, while total loss resulted in cell lethality.
|
|
mMof [Fig. 7A and unpublished data]) or wild-type mMof for IR-induced cell killing and oncogenic transformation. Cells expressing ectopic mMof have higher levels of mMof and H4K16ac than cells expressing mMof do (Fig. 7B). Cells expressing mMof had decreased survival (Fig. 7C) and reduced spontaneous and IR-induced oncogenic transformation rates compared to cells overexpressing mMof (Fig. 7D) as determined by focus formation described previously (15). We also determined the levels of mMof and H4K16ac in mouse tumors and normal tissues and found that tumor tissues have increased levels of MOF and H4K16ac in comparison to normal tissues (Fig. 7E).
|
|
MOF levels correlate with tumor growth.
To determine whether hMOF levels influence tumor growth, we used a standard nude mouse xenograft assay to examine tumor growth and its response to IR exposure. Knockdown of hMOF in colorectal carcinoma (RKO) cells (Fig. 9A) inhibited cell growth. Conversely, overexpression of hMOF increased H4K16ac (Fig. 9B) and accelerated proliferation (Fig. 9C). RKO cells, with or without overexpression of hMOF, were injected into mice, and when the tumors reached 8 mm in size, the tumors were locally irradiated with a single 25-Gy dose. Untreated tumors arising from injected RKO cells overexpressing hMOF grew rapidly compared to control cells (Fig. 9D). Tumors from RKO cells, with and without hMOF overexpression, grew at relatively constant rates, whereas radiation treatment (single dose of 25 Gy) caused temporary shrinkage of the tumors, followed by regrowth in most of the tumors. Interestingly, tumors from RKO cells with hMOF overexpression regrew faster than control cells. Two months after irradiation, 20% of the tumors from control cells were below detectable levels compared to only 8% tumors derived from cells overexpressing hMOF. An immediate relapse of tumor growth was much more prominent in cells expressing hMOF. These results suggest that hMOF promotes tumor growth.
|
|
DISCUSSION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The loss of cell proliferation could be due to cell cycle arrest specifically induced by p53 and by ATM as suggested by a previous report of G2/M arrest in hMOF knockdown cells (46). In addition, hMOF has been reported to acetylate p53 at lysine 120, and this acetylation may help distinguish between the cell cycle arrest and apoptotic functions of p53 (45). If the mMof depletion-dependent cell cycle arrest were the cause of embryonic lethality or cellular death, then depletion of p53 or ATM should have rescued or at least partially ameliorated the observed embryonic lethality. However, our results do not support this hypothesis.
Inactivation of the mMof gene in ES cells caused depletion of Mof protein accompanied with loss of acetylated H4K16 and resulted in reduced proliferation and ultimately cell death. Taken together with the lack of success in deriving homozygous Mof mutant ES cells, these experiments strongly suggest that Mof function is indispensable for cellular proliferation and/or viability. These results also raised the question of whether tumor cells can survive in the absence of MOF and the associated loss of H4K16 acetylation.
Multiple assays were utilized previously to quantitate H4K16 monoacetylation at the global level and genome region/locus-specific level, and the data suggested that loss of monoacetylation at H4K16 occurs in tumors and tumor-derived cell lines (8). Our results, however, demonstrate that tumors and tumor-derived cell lines have similar or higher levels of hMOF or H4K16ac compared to those of normal control cells (Fig. 7E and 8 and data not shown). Similar results were found in multiple different transformed cell lines (Fig. 8 and data not shown). Moreover, we found that primary cells have lower levels of hMOF and H4K16ac than the corresponding isogenic hTERT immortalized cells do (Fig. 8A). These in vitro results were also supported by the in vivo hMOF expression analysis of 300 different tumors, which demonstrated no loss of hMOF expression. In addition, tumor cells with higher levels of hMOF had faster initial growth and immediate regrowth (recurrence) of tumors after irradiation exposure (Fig. 9D), suggesting that hMOF promotes cellular proliferation. Therefore, after examination of a large number of tumor-derived cell lines and tumor samples, we propose that hMOF and H4K16ac may be required for the process of transformation, either in vitro or in vivo.
Intriguingly, MOF overexpression was associated with both increased K16H4 acetylation and increased cell proliferation (12, 46; also the present study), while cells treated with SIRT1 inhibitors had increased K16H4 acetylation levels but a reduction in cell proliferation (29). Thus, although MOF seems to be closely associated with cell proliferation, the relationship between the H4K16 acetylation status and cell proliferation requires further study.
The loss of MOF and H4K16ac correlates with increased genomic instability (Tables 1 and 2 and Fig. 6) (12, 46), which is considered an important step in cancer development because it can greatly accelerate the genetic changes leading to tumor cell progression (14). However, induction of genomic instability and increased radiosensitivity caused by depletion of MOF result in decreased oncogenic transformation, whereas overexpression of MOF resulted in increased oncogenic transformation (Fig. 7C and D); this suggests that the loss of MOF is probably not a marker of oncogenesis.
Collectively, our data suggest that the requirement for MOF during tumor cell proliferation is similar to that during embryogenesis prior to tissue differentiation. The role of MOF in cellular proliferation is supported by the following. (i) There is a strong correlation observed between the ectopic expression of the catalytic unit of telomerase (hTERT) and the levels of hMOF and H4K16ac seen in isogenic cell systems (Fig. 8A). (ii) Depletion of MOF results in cellular lethality. (iii) Overexpression of MOF results in enhanced oncogenic transformation and faster relapse of tumor growth after radiation treatment (Fig. 7D and 9D and E). Thus, the results suggest that expression of hMOF and acetylation of H4K16 are linked with proliferation status and is supported by the fact that all the tumor-derived cell lines and immortalized cell lines examined have both hMOF and acetylated H4K16. These novel observations presented here provide a foundation for future molecular studies on the significance of epigenetic modifications in varied contexts of embryonic development, tumorigenesis, and DNA metabolism.
|
ACKNOWLEDGMENTS |
|---|
This investigation was supported by funds from NIH (CA123232 and CA10445), Department of Defense, and the Department of Radiation Oncology of the Washington University School of Medicine to T.K.P., and NCI grant PO1 CA97403 (Project 5) to T.L.
|
FOOTNOTES |
|---|
Published ahead of print on 29 October 2007.
These authors contributed equally.
|
REFERENCES |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
2. Bird, A. W., D. Y. Yu, M. G. Pray-Grant, Q. Qiu, K. E. Harmon, P. C. Megee, P. A. Grant, M. M. Smith, and M. F. Christman. 2002. Acetylation of histone H4 by Esa1 is required for DNA double-strand break repair. Nature 419:411-415.[CrossRef][Medline]
3. Bodnar, A. G., M. Ouellette, M. Frolkis, S. E. Holt, C. P. Chiu, G. B. Morin, C. B. Harley, J. W. Shay, S. Lichtsteiner, and W. E. Wright. 1998. Extension of life-span by introduction of telomerase into normal human cells. Science 279:349-352.
4. Bone, J. R., J. Lavender, R. Richman, M. J. Palmer, B. M. Turner, and M. I. Kuroda. 1994. Acetylated histone H4 on the male X chromosome is associated with dosage compensation in Drosophila. Genes Dev. 8:96-104.
5. Chua, K. F., R. Mostoslavsky, D. B. Lombard, W. W. Pang, S. Saito, S. Franco, D. Kaushal, H. L. Cheng, M. R. Fischer, N. Stokes, M. M. Murphy, E. Appella, and F. W. Alt. 2005. Mammalian SIRT1 limits replicative life span in response to chronic genotoxic stress. Cell Metab. 2:67-76.[CrossRef][Medline]
6. Dou, Y., T. A. Milne, A. J. Tackett, E. R. Smith, A. Fukuda, J. Wysocka, C. D. Allis, B. T. Chait, J. L. Hess, and R. G. Roeder. 2005. Physical association and coordinate function of the H3 K4 methyltransferase MLL1 and the H4 K16 acetyltransferase MOF. Cell 121:873-885.[CrossRef][Medline]
7. Feil, R., J. Wagner, D. Metzger, and P. Chambon. 1997. Regulation of Cre recombinase activity by mutated estrogen receptor ligand-binding domains. Biochem. Biophys. Res. Commun. 237:752-757.[CrossRef][Medline]
8. Fraga, M. F., E. Ballestar, A. Villar-Garea, M. Boix-Chornet, J. Espada, G. Schotta, T. Bonaldi, C. Haydon, S. Ropero, K. Petrie, N. G. Iyer, A. Perez-Rosado, E. Calvo, J. A. Lopez, A. Cano, M. J. Calasanz, D. Colomer, M. A. Piris, N. Ahn, A. Imhof, C. Caldas, T. Jenuwein, and M. Esteller. 2005. Loss of acetylation at Lys16 and trimethylation at Lys20 of histone H4 is a common hallmark of human cancer. Nat. Genet. 37:391-400.[CrossRef][Medline]
9. Freyer, G. A., D. A. Palmer, Y. Yu, R. C. Miller, and T. K. Pandita. 1996. Neoplastic transformation of mouse C3H10T1/2 cells following exposure to neutrons does not involve mutation of ras gene as analyzed by SSCP and cycle sequencing. Mutat. Res. 357:237-244.[Medline]
10. Gage, B. M., D. Alroy, C. Y. Shin, O. N. Ponomareva, S. Dhar, G. G. Sharma, T. K. Pandita, M. J. Thayer, and M. S. Turker. 2001. Spontaneously immortalized cell lines obtained from adult Atm null mice retain sensitivity to ionizing radiation and exhibit a mutational pattern suggestive of oxidative stress. Oncogene 20:4291-4297.[CrossRef][Medline]
11. Guo, K., J. E. McMinn, T. Ludwig, Y. H. Yu, G. Yang, L. Chen, D. Loh, C. Li, S. Chua, Jr., and Y. Zhang. 2007. Disruption of peripheral leptin signaling in mice results in hyperleptinemia without associated metabolic abnormalities. Endocrinology 148:3987-3997.
12. Gupta, A., G. G. Sharma, C. S. H. Young, M. Agarwal, E. R. Smith, T. T. Paull, J. C. Lucchesi, K. K. Khanna, T. Ludwig, and T. K. Pandita. 2005. Involvement of human MOF in ATM function. Mol. Cell. Biol. 25:5292-5305.
13. Hilfiker, A., D. Hilfiker-Kleiner, A. Pannuti, and J. C. Lucchesi. 1997. mof, a putative acetyl transferase gene related to the Tip60 and MOZ human genes and to the SAS genes of yeast, is required for dosage compensation in Drosophila. EMBO J. 16:2054-2060.[CrossRef][Medline]
14. Hoeijmakers, J. H. 2001. Genome maintenance mechanisms for preventing cancer. Nature 411:366-374.[CrossRef][Medline]
15. Hunt, C. R., D. J. Dix, G. G. Sharma, R. K. Pandita, A. Gupta, M. Funk, and T. K. Pandita. 2004. Genomic instability and enhanced radiosensitivity in Hsp70.1- and Hsp70.3-deficient mice. Mol. Cell. Biol. 24:899-911.
16. Kim, N. W., M. A. Piatyszek, K. R. Prowse, C. B. Harley, M. D. West, P. L. Ho, G. M. Coviello, W. E. Wright, S. L. Weinrich, and J. W. Shay. 1994. Specific association of human telomerase activity with immortal cells and cancer. Science 266:2011-2015.
17. Kimura, A., K. Matsubara, and M. Horikoshi. 2005. A decade of histone acetylation: marking eukaryotic chromosomes with specific codes. J. Biochem. (Tokyo) 138:647-662.
18. Kusch, T., L. Florens, W. H. Macdonald, S. K. Swanson, R. L. Glaser, J. R. Yates III, S. M. Abmayr, M. P. Washburn, and J. L. Workman. 2004. Acetylation by Tip60 is required for selective histone variant exchange at DNA lesions. Science 306:2084-2087.
19. Lavender, J. S., A. J. Birley, M. J. Palmer, M. I. Kuroda, and B. M. Turner. 1994. Histone H4 acetylated at lysine 16 and proteins of the Drosophila dosage compensation pathway co-localize on the male X chromosome through mitosis. Chromosome Res. 2:398-404.[CrossRef][Medline]
20. Morgan, S. E., and M. B. Kastan. 1997. p53 and ATM: cell cycle, cell death, and cancer. Adv. Cancer Res. 71:1-25.[Medline]
21. O'Neill, L. P., and B. M. Turner. 1995. Histone H4 acetylation distinguishes coding regions of the human genome from heterochromatin in a differentiation-dependent but transcription-independent manner. EMBO J. 14:3946-3957.[Medline]
22. Pandita, T. K. 2003. A multifaceted role for ATM in genome maintenance. Expert Rev. Mol. Med. 5:1-21.[Medline]
23. Pandita, T. K. 1988. Assessment of the mutagenic potential of a fungicide Bavistin using multiple assays. Mutat. Res. 204:627-643.[CrossRef][Medline]
24. Pandita, T. K. 1983. Mutagenic studies on the insecticide Metasystox-R with different genetic systems. Mutat. Res. 124:97-102.[CrossRef][Medline]
25. Pandita, T. K. 2006. Role of mammalian Rad9 in genomic stability and ionizing radiation response. Cell Cycle 5:1289-1291.[Medline]
26. Pandita, T. K., and W. N. Hittelman. 1992. Initial chromosome damage but not DNA damage is greater in ataxia telangiectasia cells. Radiat. Res. 130:94-103.[Medline]
27. Pandita, T. K., H. B. Lieberman, D. S. Lim, S. Dhar, W. Zheng, Y. Taya, and M. B. Kastan. 2000. Ionizing radiation activates the ATM kinase throughout the cell cycle. Oncogene 19:1386-1391.[CrossRef][Medline]
28. Peterson, C. L., and M. A. Laniel. 2004. Histones and histone modifications. Curr. Biol. 14:R546-R551.[CrossRef][Medline]
29. Pruitt, K., R. L. Zinn, J. E. Ohm, K. M. McGarvey, S. H. Kang, D. N. Watkins, J. G. Herman, and S. B. Baylin. 2006. Inhibition of SIRT1 reactivates silenced cancer genes without loss of promoter DNA hypermethylation. PLoS Genet. 2:e40.[CrossRef][Medline]
30. Reznikoff, C. A., J. S. Bertram, D. W. Brankow, and C. Heidelberger. 1973. Quantitative and qualitative studies of chemical transformation of cloned C3H mouse embryo cells sensitive to postconfluence inhibition of cell division. Cancer Res. 33:3239-3249.
31. Reznikoff, C. A., D. W. Brankow, and C. Heidelberger. 1973. Establishment and characterization of a cloned line of C3H mouse embryo cells sensitive to postconfluence inhibition of division. Cancer Res. 33:3231-3238.
32. Sawant, S. G., V. Gregoire, S. Dhar, C. B. Umbricht, S. Cvilic, S. Sukumar, and T. K. Pandita. 1999. Telomerase activity as a measure for monitoring radiocurability of tumor cells. FASEB J. 13:1047-1054.
33. Scherthan, H., M. Jerratsch, S. Dhar, Y. A. Wang, S. P. Goff, and T. K. Pandita. 2000. Meiotic telomere distribution and Sertoli cell nuclear architecture are altered in Atm- and Atm-p53-deficient mice. Mol. Cell. Biol. 20:7773-7783.
34. Sharma, G. G., A. Gupta, H. Wang, H. Scherthan, S. Dhar, V. Gandhi, G. Iliakis, J. W. Shay, C. S. Young, and T. K. Pandita. 2003. hTERT associates with human telomeres and enhances genomic stability and DNA repair. Oncogene 22:131-146.[CrossRef][Medline]
35. Sharma, G. G., K. K. Hwang, R. K. Pandita, A. Gupta, S. Dhar, J. Parenteau, M. Agarwal, H. J. Worman, R. J. Wellinger, and T. K. Pandita. 2003. Human heterochromatin protein 1 isoforms HP1Hs
and HP1Hsβ interfere with hTERT-telomere interactions and correlate with changes in cell growth and response to ionizing radiation. Mol. Cell. Biol. 23:8363-8376.
36. Shia, W. J., B. Li, and J. L. Workman. 2006. SAS-mediated acetylation of histone H4 Lys 16 is required for H2A.Z incorporation at subtelomeric regions in Saccharomyces cerevisiae. Genes Dev. 20:2507-2512.
37. Shia, W. J., S. G. Pattenden, and J. L. Workman. 2006. Histone H4 lysine 16 acetylation breaks the genome's silence. Genome Biol. 7:217.[CrossRef][Medline]
38. Shiloh, Y. 2003. ATM and related protein kinases: safeguarding genome integrity. Nat. Rev. Cancer 3:155-168.[CrossRef][Medline]
39. Shogren-Knaak, M., H. Ishii, J. M. Sun, M. J. Pazin, J. R. Davie, and C. L. Peterson. 2006. Histone H4-K16 acetylation controls chromatin structure and protein interactions. Science 311:844-847.
40. Smilenov, L. B., S. E. Morgan, W. Mellado, S. G. Sawant, M. B. Kastan, and T. K. Pandita. 1997. Influence of ATM function on telomere metabolism. Oncogene 15:2659-2665.[CrossRef][Medline]
41. Smith, E. R., C. Cayrou, R. Huang, W. S. Lane, J. Cote, and J. C. Lucchesi. 2005. A human protein complex homologous to the Drosophila MSL complex is responsible for the majority of histone H4 acetylation at lysine 16. Mol. Cell. Biol. 25:9175-9188.
42. Smith, E. R., A. Pannuti, W. Gu, A. Steurnagel, R. G. Cook, C. D. Allis, and J. C. Lucchesi. 2000. The Drosophila MSL complex acetylates histone H4 at lysine 16, a chromatin modification linked to dosage compensation. Mol. Cell. Biol. 20:312-318.
43. Sterner, D. E., and S. L. Berger. 2000. Acetylation of histones and transcription-related factors. Microbiol. Mol. Biol. Rev. 64:435-459.
44. Suka, N., K. Luo, and M. Grunstein. 2002. Sir2p and Sas2p opposingly regulate acetylation of yeast histone H4 lysine 16 and spreading of heterochromatin. Nat. Genet. 32:378-383.[CrossRef][Medline]
45. Sykes, S. M., H. S. Mellert, M. A. Holbert, K. Li, R. Marmorstein, W. S. Lane, and S. B. McMahon. 2006. Acetylation of the p53 DNA-binding domain regulates apoptosis induction. Mol. Cell 24:841-851.[CrossRef][Medline]
46. Taipale, M., S. Rea, K. Richter, A. Vilar, P. Lichter, A. Imhof, and A. Akhtar. 2005. hMOF histone acetyltransferase is required for histone H4 lysine 16 acetylation in mammalian cells. Mol. Cell. Biol. 25:6798-6810.
47. Vaziri, H., and S. Benchimol. 1998. Reconstitution of telomerase activity in normal human cells leads to elongation of telomeres and extended replicative life span. Curr. Biol. 8:279-282.[CrossRef][Medline]
48. Wood, L. D., T. L. Halvorsen, S. Dhar, J. A. Baur, R. K. Pandita, W. E. Wright, M. P. Hande, G. Calaf, T. K. Hei, F. Levine, J. W. Shay, J. J. Wang, and T. K. Pandita. 2001. Characterization of ataxia telangiectasia fibroblasts with extended life-span through telomerase expression. Oncogene 20:278-288.[CrossRef][Medline]
49. Wright, P. W., L. C. Bolling, M. E. Calvert, O. F. Sarmento, E. V. Berkeley, M. C. Shea, Z. Hao, F. C. Jayes, L. A. Bush, J. Shetty, A. N. Shore, P. P. Reddi, K. S. Tung, E. Samy, M. M. Allietta, N. E. Sherman, J. C. Herr, and S. A. Coonrod. 2003. ePAD, an oocyte and early embryo-abundant peptidylarginine deiminase-like protein that localizes to egg cytoplasmic sheets. Dev. Biol. 256:73-88.[Medline]
50. Ziv, S., O. Brenner, N. Amariglio, N. I. Smorodinsky, R. Galron, D. V. Carrion, W. Zhang, G. G. Sharma, R. K. Pandita, M. Agarwal, R. Elkon, N. Katzin, I. Bar-Am, T. K. Pandita, R. Kucherlapati, G. Rechavi, Y. Shiloh, and A. Barzilai. 2005. Impaired genomic stability and increased oxidative stress exacerbate different features of ataxia-telangiectasia. Hum. Mol. Genet. 14:2929-2943.
| ||||||||||||
