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1 Department of Developmental Genetics, Center for Translational and Advanced Animal Research, Graduate School of Medicine, Tohoku University, 2-1 Seiryo, Aoba-ku, Sendai 980-8575, Japan
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CREST, Japan Science and Technology Agency, Kawaguchi, Saitama 332-0012, Japan
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Department of Molecular and Cellular Biology, Medical Institute of Bioregulation, Kyushu University, Fukuoka 812-8582, Japan
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Abstract |
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 Top Abstract IntroductionResultsDiscussionExperimental proceduresReferences |
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Introduction |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
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Geminin was originally discovered in a screen for proteins that were selectively degraded by extracts of mitotic Xenopus eggs (but not by those of eggs in interphase) (McGarry & Kirschner 1998). Geminin contains a destruction box that is recognized by the anaphase-promoting complex/cyclosome (APC/C) ubiquitin ligase and is responsible for the ubiquitin-dependent proteolysis of geminin. The expression of geminin is thus suppressed, whereas pre-RCs are formed at replication origins during the G1 phase. Geminin subsequently accumulates in the nucleus at the beginning of S phase. The central portion of geminin contains five heptad amino acid repeats that are predicted to form a coiled-coil domain, a structure commonly associated with protein dimerization (Benjamin et al. 2004; Okorokov et al. 2004). The geminin dimer binds to Cdt1 with high affinity in the nucleus and prevents it from contributing to pre-RCs during S and G2 phases (Wohlschlegel et al. 2000; Tada et al. 2001). Furthermore, access of the MCM complex to Cdt1 is restricted by the COOH-terminal portion of the coiled-coil domain of geminin (Lee et al. 2004). These observations suggest that geminin antagonizes the function of Cdt1. An excess of Cdt1 overcomes the inhibitory effect of geminin. The expression of geminin in the nucleus is sustained throughout the S and G2 phases, during which time geminin limits Cdt1 activity in order to inhibit re-replication (Mihaylov et al. 2002).
Development must be tightly linked to decisions regarding cell division. Geminin has also been recognized as a factor that contributes to embryonic patterning as well as a regulator of the cell cycle. Microinjection of geminin mRNA into Xenopus blastomeres revealed that geminin possesses neuralizing activity (Kroll et al. 1998). This activity is explained, at least in part, by the observation that geminin directly interacts with Hox and Polycomb proteins (Del Bene et al. 2004; Luo et al. 2004; Seo et al. 2005). Hox proteins induce dissociation of geminin from the Cdt1-geminin complex, rendering Cdt1 available for DNA replication. Such competitive interactions of geminin with Cdt1 and Hox proteins suggest that geminin coordinates development and cell proliferation during embryogenesis.
To evaluate the importance of geminin in mammalian embryogenesis, we have now disrupted the geminin gene by homologous recombination in mice. The resulting geminin-deficient embryos were found to arrest development early during embryogenesis, manifesting an abnormal cell morphology at the eight-cell stage. The geminin-deficient cells entered S phase but had lost the ability to enter M phase. They exhibited DNA damage and underwent apoptosis. Our data suggest that geminin is essential for early embryonic development as a result of its coordination of the progression through S to M phase of the cell cycle.
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Results |
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To elucidate the physiologic functions of geminin, we generated mice deficient in this protein. The geminin gene was disrupted in mouse embryonic stem (ES) cells by replacement of exons 5 and 6, which encode the coiled-coil domain of the protein, with a PGK-neo-poly(A) cassette (Fig. 1A). Given that the coiled-coil domain is required for the dimerization of geminin, which itself is necessary for binding of geminin to Cdt1, a functional protein would not be expected to be expressed from the mutated allele. The targeting construct was introduced into ES cells by electroporation, and 3 of 216 resulting G418- and ganciclovir-resistant colonies were shown to be heterozygous for the geminin locus. Each of these targeted ES clones was used to generate chimeric mice. Chimeras produced from FL29 and FL134 clones were back-crossed to strain C57BL/6 J animals in order to generate mice heterozygous for the mutant geminin allele. We verified the homologous recombination event by Southern blot analysis of the heterozygous mutant mice (Fig. 1B). Adult geminin heterozygous (geminin+/–) mice were overtly normal, and no defect in the proliferation of embryonic fibroblasts derived from these animals was apparent (data not shown). We then intercrossed geminin+/– mice and determined the genotypes of 165 of the resulting offspring by polymerase chain reaction (PCR) analysis of tail DNA at 3 weeks of age. The ratio of heterozygous mice to wild-type mice appeared normal, but no homozygous mutants were detected (Table 1). Given that mortality of neonates was not apparent during the first 3 weeks after birth, this lack of geminin–/– mice indicated that geminin deficiency results in embryonic death.
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We investigated the time of the presumptive embryonic mortality of geminin–/– mice by analyzing the genotypes of embryos produced from heterozygote intercrosses at various stages of embryogenesis. PCR analysis of postimplantation embryos dissected from the decidua at embryonic day (E) 7.5 to E13.5 (with the morning of vaginal plug detection corresponding to E0.5) showed that none of these embryos were geminin–/– (Table 1). However, geminin–/– preimplantation embryos were detected at E3.5 (Fig. 1C, Table 1). Immunofluorescence analysis confirmed a lack of geminin expression in the homozygous mutant embryos (Fig. 1D,F). Nuclear staining with 4',6-diamidino-2-phenylindole (DAPI) revealed the nuclei of the mutant embryos to be irregular in both size and shape (Fig. 1E,G). Morphologic analysis showed that all 80 geminin–/– embryos examined at E3.5 contained abnormal nuclei, whereas the nuclei of all geminin+/+ or geminin+/– embryos examined appeared normal. These results indicated that loss of geminin function results in gross changes in nuclear morphology, which may be directly responsible for the arrested development of geminin–/– embryos.
The homozygous mutant embryos at E3.5 also manifested other morphologic abnormalities. The trophectoderm epithelium, which normally lines the zona pellucida at this stage, was missing from the mutant embryos, and individual cells constituting the inner cell mass appeared to be dispersed with visible cell boundaries, whereas wild-type cells are densely aggregated and the cell boundaries are usually not visible by phase-contrast microscopy (Fig. 2A,B). In addition, the sizes both of the cells and of their nuclei were heterogeneous in geminin–/– embryos. These morphologic characteristics were suggestive of a defect in cell adhesion in geminin–/– embryos at E3.5. Immunofluorescence analysis with antibodies to E-cadherin and to ß-catenin revealed that, whereas wild-type blastocysts exhibited an organized pattern of E-cadherin and ß-catenin expression along the cell boundaries, the corresponding fluorescence signals were weak or undetectable in the mutant embryos (Fig. 2C–H). The detachment of the cells in E3.5 geminin–/– embryos seemed to be secondary to an abnormality of the cell cycle (see succeeding discussions).
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The presence of aberrant nuclei in geminin–/– embryos suggested that geminin deficiency might affect embryonic cell proliferation. To directly assess the growth capability of geminin–/– embryos, we collected embryos from heterozygote intercrosses at E3.5 and cultured them individually in vitro for 4 days, during which they formed outgrowths. Both geminin+/+ and geminin+/– blastocysts hatched, attached to the culture dish, and produced apparently normal trophoblast giant cells. In contrast, geminin–/– embryos failed to attach to the culture dish or died during in vitro culture (data not shown), suggesting that geminin deficiency results in a defect in cell proliferation at E3.5.
Given that geminin functions in signaling associated with replication licensing, the aberrant nuclei in geminin–/– embryos may have reflected a defect in licensing function. To examine this possibility, we investigated cell cycle progression in the mutant embryos. For detection of cells in S or M phases, E3.5 embryos were cultured briefly in the presence of bromodeoxyuridine (BrdU) and then subjected to immunofluorescence analysis with antibodies specific for BrdU and for histone H3 phosphorylated on Ser10 (pHH3), an M phase-specific marker (Ajiro et al. 1996). Mutant embryos incorporated BrdU, suggesting that geminin–/– cells retained the ability to enter S phase (Fig. 3A–F). In contrast, whereas a substantial proportion of the cells of wild-type embryos were found to enter M phase, cells that stained with the antibodies to pHH3 were not detected in the geminin–/– embryos (Fig. 3G,H). These results thus suggested that the loss of geminin renders cells unable to proceed into M phase, likely as a result of the activation of a checkpoint mechanism that senses aberrant DNA replication or DNA damage (or both).
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We found that geminin–/– embryos at E3.5 were reactive with antibodies to 
H2AX, a variant of histone H2 that is phosphorylated on Ser139 and accumulates in discrete foci at sites of DNA damage (Burma et al. 2001), whereas no signal was detected with these antibodies in wild-type embryos (Fig. 3I–L). These data suggested that the lack of geminin results in accumulation of spontaneous DNA damage. Furthermore, the terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) assay revealed that the mutant embryos underwent apoptosis in response to the DNA damage, whereas no apoptosis was detected in wild-type embryos (Fig. 4).
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To determine when the morphological abnormalities of geminin–/– embryos first become apparent, we collected embryos at the two-cell stage (E1.5) and microscopically examined the progress of their development in culture. The geminin–/– embryos were indistinguishable from wild-type embryos until the four-cell stage (Fig. 5A–F), after which their development was retarded. The mutant embryos thus remained at the four-cell stage 1.5 days after initiation of culture, whereas wild-type embryos had progressed to the eight-cell stage by this time (Fig. 5G–I). Irregularly sized cells were occasionally observed in the mutant embryos at this stage (Fig. 5I). Despite the retardation of development and morphologic abnormalities, geminin–/– embryos appeared to undergo compaction (Fig. 5J–L). However, the trophectoderm epithelium lining the zona pellucida was not formed and the cell mass became fragmented in geminin–/– embryos (Fig. 5M–O). No cavity was formed in the mutant embryos. These observations indicated that the morphologic abnormalities and developmental retardation of geminin–/– embryos become evident at the transition from the four- to eight-cell stages and markedly worsen at the eight-cell stage (Fig. 6). Immunofluorescence analysis revealed that maternal geminin was present until the four-cell stage in geminin–/– embryos, with the subsequent disappearance of the maternal protein coinciding with the onset of developmental retardation at the transition from the four- to eight-cell stages (Fig. 7). These data suggested that the normal progression of development of geminin–/– embryos until the four-cell stage may be attributable to the presence of maternal geminin.
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Given that E3.5 mutant embryos exhibited defects in cell cycle progression and cell-cell adhesion, we attempted to determine which of these defects occurs first. Immunofluorescence staining of E2.0 embryos at the four- to eight-cell stages with antibodies to pHH3 revealed that the number of cells in mitosis was decreased in geminin–/– embryos (Fig. 8A–D), possibly accounting for the delay in the cleavage of mutant blastomeres. Cell adherence complexes and tight junctions are formed at the late eight-cell stage in wild-type embryos (Fig. 8E,G). Mutant embryos that remained at the four-cell stage manifested normal distributions of E-cadherin (Fig. 8F,H) and ß-catenin (data not shown) at the cell periphery. These data suggested that the expression and distribution of adhesion molecules such as E-cadherin are maintained after deterioration of cell cycle progression. The impairment of the cell cycle therefore appears to be the primary cause of developmental failure in geminin–/– embryos.
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Discussion |
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Mammalian Chk1 is activated in cells treated with inhibitors of DNA replication such as hydroxyurea and aphidicolin. Chk1 is also activated by DNA double-strand breaks generated by exposure of cells to ionizing radiation (Feijoo et al. 2001). Mouse embryos deficient in Chk1 die between E3.5 and E7.5, and ES cells lacking Chk1 undergo apoptosis. Both Chk1-deficient embryos and ES cells are defective in the G2-M checkpoint evoked by inhibition of DNA replication by aphidicolin or hydroxyurea. Moreover, such cells lacking Chk1 are unable to arrest the cell cycle in response to ionizing or ultraviolet irradiation. Chk1 thus appears to function as a primary effector kinase that enforces checkpoint control in response to aberrant replication or DNA damage in mammalian cells (Takai et al. 2000).
We have now shown that cell cycle progression is delayed in geminin-deficient embryos, and that this defect is associated with heterogeneity in cell and nuclear size. Although DNA synthesis occurs in the nuclei of these abnormal cells, the cells do not enter M phase, likely resulting in over-replication of genomic DNA and in activation of the Chk1-dependent checkpoint. Although the precise mechanism remains unclear, abnormal replication results in double-strand breakage in genomic DNA, as revealed by the presence of phosphorylated histone H2AX. We also detected phosphorylated histone H2AX in geminin–/– blastomeres. Over-replication and subsequent DNA double-strand breakage thus likely trigger the checkpoint response and prevent the mutant blastomeres from entering mitosis.
We detected apoptosis in geminin-deficient embryos but not in wild-type ones. Depletion of geminin by RNAi in normal human cells did not trigger apoptosis despite the checkpoint activation induced by re-replication. However, prevention of Chk1 activation through inhibition of the upstream kinases ATM and ATR by caffeine in cells depleted of geminin by RNAi released the cells from checkpoint-induced arrest at G2 phase, resulting in entry into mitosis followed by apoptosis. These findings suggest that Chk1 is activated and inhibits both the G2-M transition and apoptosis in the geminin-depleted cells (Melixetian et al. 2004). In contrast, during mouse development, although geminin deficiency results in delayed cell cycle progression, it also elicits apoptosis. This apparent discrepancy might be explained if geminin performs other unidentified functions during development.
Although geminin-deficient embryos appear to undergo compaction, cell–cell adhesion is impaired soon thereafter, likely as a result of apoptosis. Compaction is a crucial event of early embryonic development and normally occurs within 2–3 h of embryos having progressed to the eight-cell stage (Fleming & Johnson 1988). During compaction, blastomeres that are loosely associated with each other become densely aggregated and the cell boundaries become indistinct on examination by phase-contrast microscopy. Compaction is accompanied by regionalization of cell adherence complexes at sites of cell–cell contact, resulting in the generation of spatial heterogeneity and polarity in the embryos. Indeed, cell adherence complex proteins have been shown to be essential for compaction, with antibodies to E-cadherin preventing this process (Ekblom et al. 1986). The localization of E-cadherin and ß-catenin did not appear to differ between geminin-deficient and wild-type mouse embryos at E2.0, even though progression of the cell cycle was already delayed in the mutant embryos. These observations suggest that cell–cell adhesion and the expression of adhesion molecules such as E-cadherin are independent of developmental stage or the cell cycle, being dependent instead on time after fertilization. The cell cycle defect of geminin–/– embryos thus appears to be central to the failure of development. It remains possible, however, that the premature compaction of geminin–/– blastomeres at the four- to eight-cell stage may also contribute to their developmental abnormality.
The imbalance between geminin and Cdt1 may also contribute to genomic instability and tumorigenesis. Subcutaneous injection of primary cells that over-express Cdt1 induces tumor formation in mice (Arentson et al. 2002). In contrast, over-expression of a stable mutant of geminin in the HCT116 colorectal carcinoma cell line suppresses tumorigenesis in vivo (Yoshida et al. 2004). However, a search for mutations or deletions of the geminin gene in breast carcinoma tissue samples did not support the possibility that geminin functions as a tumor suppressor (Yoshida et al. 2004). On the other hand, geminin was identified as an effective independent marker for tumor prognosis (Yu et al. 2004). Further evaluation of the anti-tumor activity of geminin will require the establishment of animal models in which geminin is conditionally ablated.
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Experimental procedures |
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The mouse geminin gene (geminin) was isolated by screening a mouse 129/Sv Lambda FIX II Genomic Library (Stratagene) with mouse geminin cDNA as a probe. Three overlapping genomic clones containing exons 1–7 of geminin were isolated and subcloned into the pBluescript II SK+ vector (Stratagene). The targeting vector pGem.KO was designed to replace a 1.2 kb XbaI-EcoRV genomic fragment containing exons 5 and 6 of geminin with a PGK-neo-poly(A) cassette in the same transcriptional orientation as that of geminin. The targeting vector contained 5.7 kb (XbaI-XbaI) and 2.8 kb (EcoRV-EcoRV) regions of homology located 5' and 3', respectively, relative to the neomycin-resistance gene (neo). A PGK-tk-poly(A) cassette was ligated at the 5' end of the targeting construct. The maintenance, transfection, and selection of ES cells were performed as previously described (Nakayama et al. 1996). The recombination event was confirmed by Southern blot analysis with a 0.5-kb EcoRV-EcoRV fragment of genomic DNA that flanked the 3' homology region as the probe (Fig. 1A). The expected sizes of hybridizing fragments after digestion of genomic DNA with EcoRI were 9.2 and 7.8 kb for the wild-type and mutant geminin alleles, respectively. Mutant ES cells were micro-injected into C57BL/6 mouse blastocysts, and the resulting male chimeras were mated with C57BL/6 females. Germ-line transmission of the mutant allele was confirmed by Southern blot analysis. Heterozygous offspring were intercrossed to produce homozygous mutant animals. For genotyping of embryos, DNA was extracted from whole embryos at E2.5 to E3.5. The extracted DNA was then analyzed by nested PCR with the primers F1 (5'-CTCTGGTGTTCGCTCTTCCTATTG-3'), F2 (5'-CAAACGCTACTCCTGGGACAAG-3'), R1 (5'-TTCGCAGGGGCTTTGATGTATTTC-3'), R2 (5'-GGTGGATGTGGAATGTGTGCGAGG-3'), R3 (5'-AAATGGCTAAGTGGGCAAAGC-3'), and R4 (5'-CGAGGCCAGAGGCCACTTGTGTAG-3'). The wild-type allele of geminin was amplified with primers F2 and R3 followed by F1 and R1; the mutant allele was amplified with F1 and R4 followed by F1 and R2. The present study conformed to the guidelines of Tohoku University for the care and use of laboratory animals.
Immunofluorescence and TUNEL staining
Embryos were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 10 min at room temperature and were then permeabilized for 12 min at room temperature with PBS containing 0.2% Triton X-100. The embryos were stained with rabbit polyclonal antibodies to geminin (kindly provided by H. Nishitani) or to E-cadherin (DECMA-1, Sigma) or with mouse monoclonal antibodies to ß-catenin (610153, BD Biosciences Pharmingen), to histone H3 phosphorylated on Ser10 (06–570, Upstate Biotechnology), or to histone H2AX phosphorylated on Ser139 (07–164, Upstate Biotechnology). Immune complexes were detected with Alexa 488– or Alexa 546–conjugated secondary antibodies to rabbit or mouse immunoglobulin G (Molecular Probes). Nuclei were stained with DAPI. Blasotomeres in the S phase were labeled by incubation of embryos for 3 h in medium containing 10 µM BrdU (Sigma) and were then detected by immunostaining with rat monoclonal antibodies to BrdU (BU1/75 ICR1, Abcam) (Aoki & Schultz 1999). For the TUNEL assay, fixed embryos were permeabilized for 6 min at room temperature with PBS containing 0.5% Triton X-100, pre-incubated in the TUNEL Label mixture (Roche) for 10 min at room temperature, and then incubated for 1 h at 37 °C with TUNEL Label mixture containing 10% (v/v) TUNEL enzyme (Roche) (Kim et al. 2002). Specimens were mounted on slides with ProLong Gold anti-fade reagent (Molecular Probes) and were observed with a laser-scanning confocal microscope (Zeiss).
Culture of preimplantation embryos
Heterozygous male and female mutant mice were intercrossed, and the morning of the day on which a vaginal plug was detected was designated E0.5. Embryos at different stages of development (E0.5 to E3.5) were collected by flushing the oviduct or uterus with HEPES-buffered medium (M2, Sigma). After washing with HEPES-free M16 medium (Sigma), embryos were cultured for the indicated times under a humidified atmosphere of 5% CO2 at 37 °C in tissue culture plates containing M16 medium before staining or examination by phase-contrast microscopy.
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Acknowledgements |
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Footnotes |
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* Correspondence: E-mail: nakayak2{at}mail.tains.tohoku.ac.jp
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References |
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Aoki, E. & Schultz, R.M. (1999) DNA replication in the 1-cell mouse embryo: stimulatory effect of histone acetylation. Zygote 7, 165–172.[CrossRef][Medline]
Arentson, E., Faloon, P., Seo, J., Moon, E., Studts, J.M., Fremont, D.H. & Choi, K. (2002) Oncogenic potential of the DNA replication licensing protein CDT1. Oncogene 21, 1150–1158.[CrossRef][Medline]
Benjamin, J.M., Torke, S.J., Demeler, B. & McGarry, T.J. (2004) Geminin has dimerization, Cdt1-binding, and destruction domains that are required for biological activity. J. Biol. Chem. 279, 45957–45968.
Blow, J.J. & Laskey, R.A. (1988) A role for the nuclear envelope in controlling DNA replication within the cell cycle. Nature 332, 546–548.[CrossRef][Medline]
Burma, S., Chen, B.P., Murphy, M., Kurimasa, A. & Chen, D.J. (2001) ATM phosphorylates histone H2AX in response to DNA double-strand breaks. J. Biol. Chem. 276, 42462–42467.
Del Bene, F., Tessmar-Raible, K. & Wittbrodt, J. (2004) Direct interaction of geminin and Six3 in eye development. Nature 427, 745–749.[CrossRef][Medline]
Delmolino, L.M., Saha, P. & Dutta, A. (2001) Multiple mechanisms regulate subcellular localization of human CDC6. J. Biol. Chem. 276, 26947–26954.
Diffley, J.F., Nishitani, H. & Lygerou, Z. (2004) Regulation of early events in chromosome replication. Control of DNA replication licensing in a cell cycle. Curr. Biol. 14, R778–R786.[CrossRef][Medline]
Ekblom, P., Vestweber, D. & Kemler, R. (1986) Cell–matrix interactions and cell adhesion during development. Annu. Rev. Cell Biol. 2, 27–47.[Medline]
Feijoo, C., Hall-Jackson, C., Wu, R., Jenkins, D., Leitch, J., Gilbert, D.M. & Smythe, C. (2001) Activation of mammalian Chk1 during DNA replication arrest: a role for Chk1 in the intra-S phase checkpoint monitoring replication origin firing. J. Cell Biol. 154, 913–923.
Fleming, T.P. & Johnson, M.H. (1988) From egg to epithelium. Annu. Rev. Cell Biol. 4, 459–485.[CrossRef][Medline]
Kelly, T.J. & Brown, G.W. (2000) Regulation of chromosome replication. Annu. Rev. Biochem. 69, 829–880.[CrossRef][Medline]
Kim, J.M., Ogura, A., Nagata, M. & Aoki, F. (2002) Analysis of the mechanism for chromatin remodeling in embryos reconstructed by somatic nuclear transfer. Biol. Reprod. 67, 760–766.
Kroll, K.L., Salic, A.N., Evans, L.M. & Kirschner, M.W. (1998) Geminin, a neuralizing molecule that demarcates the future neural plate at the onset of gastrulation. Development 125, 3247–3258.[Abstract]
Laskey, R.A., Fairman, M.P. & Blow, J.J. (1989) S phase of the cell cycle. Science 246, 609–614.
Lee, C., Hong, B., Choi, J.M., Kim, Y., Watanabe, S., Ishimi, Y., Enomoto, T., Tada, S., Kim, Y. & Cho, Y. (2004) Structural basis for inhibition of the replication licensing factor Cdt1 by geminin. Nature 430, 913–917.[CrossRef][Medline]
Lei, M. & Tye, B.K. (2001) Initiating DNA synthesis: from recruiting to activating the MCM complex. J. Cell Sci. 114, 1447–1454.[Abstract]
Luo, L., Yang, X., Takihara, Y., Knoetgen, H. & Kessel, M. (2004) The cell-cycle regulator geminin inhibits Hox function through direct and polycomb-mediated interactions. Nature 427, 749–753.[CrossRef][Medline]
McGarry, T.J. (2002) Geminin deficiency causes a Chk1-dependent G2 arrest in Xenopus. Mol. Biol. Cell 13, 3662–3671.
McGarry, T.J. & Kirschner, M.W. (1998) Geminin, an inhibitor of DNA replication, is degraded during mitosis. Cell 93, 1043–1053.[CrossRef][Medline]
Melixetian, M., Ballabeni, A., Masiero, L., Gasparini, P., Zamponi, R., Bartek, J., Lukas, J. & Helin, K. (2004) Loss of Geminin induces rereplication in the presence of functional p53. J. Cell Biol. 165, 473–482.
Mihaylov, I.S., Kondo, T., Jones, L., Ryzhikov, S., Tanaka, J., Zheng, J., Higa, L.A., Minamino, N., Cooley, L. & Zhang, H. (2002) Control of DNA replication and chromosome ploidy by geminin and cyclin A. Mol. Cell. Biol. 22, 1868–1880.
Nakayama, K., Ishida, N., Shirane, M., Inomata, A., Inoue, T., Shishido, N., Horii, I., Loh, D.Y. & Nakayama, K.I. (1996) Mice lacking p27Kip1 display increased body size, multiple organ hyperplasia, retinal dysplasia, and pituitary tumors. Cell 85, 707–720.[CrossRef][Medline]
Nishitani, H. & Lygerou, Z. (2002) Control of DNA replication licensing in a cell cycle. Genes Cells 7, 523–534.[Abstract]
Okorokov, A.L., Orlova, E.V., Kingsbury, S.R., Bagneris, C., Gohlke, U., Williams, G.H. & Stoeber, K. (2004) Molecular structure of human geminin. Nat. Struct. Mol. Biol. 11, 1021–1022.[CrossRef][Medline]
Saxena, S., Dutta, A., Diffley, J.F., Nishitani, H. & Lygerou, Z. (2005) Geminin-Cdt1 balance is critical for genetic stability; regulation of early events in chromosome replication control of DNA replication licensing in a cell cycle. Mutat. Res. 569, 111–121.[Medline]
Seo, S., Herr, A., Lim, J.W., Richardson, G.A., Richardson, H. & Kroll, K.L. (2005) Geminin regulates neuronal differentiation by antagonizing Brg1 activity. Genes Dev. 19, 1723–1734.
Tada, S., Li, A., Maiorano, D., Mechali, M. & Blow, J.J. (2001) Repression of origin assembly in metaphase depends on inhibition of RLF-B/Cdt1 by geminin. Nat. Cell Biol. 3, 107–113.[CrossRef][Medline]
Takai, H., Tominaga, K., Motoyama, N., Minamishima, Y.A., Nagahama, H., Tsukiyama, T., Ikeda, K., Nakayama, K., Nakanishi, M. & Nakayama, K.I. (2000) Aberrant cell cycle checkpoint function and early embryonic death in Chk1–/– mice. Genes Dev. 14, 1439–1447.
Takisawa, H., Mimura, S. & Kubota, Y. (2000) Eukaryotic DNA replication: from pre-replication complex to initiation complex. Curr. Opin. Cell Biol. 12, 690–696.[CrossRef][Medline]
Wohlschlegel, J.A., Dwyer, B.T., Dhar, S.K., Cvetic, C., Walter, J.C. & Dutta, A. (2000) Inhibition of eukaryotic DNA replication by geminin binding to Cdt1. Science 290, 2309–2312.
Yoshida, K., Oyaizu, N., Dutta, A. & Inoue, I. (2004) The destruction box of human Geminin is critical for proliferation and tumor growth in human colon cancer cells. Oncogene 23, 58–70.[CrossRef][Medline]
Yu, K., Lee, C.H., Tan, P.H., Hong, G.S., Wee, S.B., Wong, C.Y. & Tan, P. (2004) A molecular signature of the Nottingham prognostic index in breast cancer. Cancer Res. 64, 2962–2968.
Zhu, W., Chen, Y. & Dutta, A. (2004) Rereplication by depletion of Geminin is seen regardless of p53 status and activates a G2/M checkpoint. Mol. Cell. Biol. 24, 7140–7150.
Received: 27 June 2006
Accepted: 10 August 2006
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