Induction of Cell-Cycle Arrest by all-trans Retinoic Acid in Mouse Embryonic Palatal Mesenchymal (MEPM) Cells
http://www.100md.com
《毒物学科学杂志》
School of Stomatology, Peking University, Beijing, 100081, China
School of Public Health, Peking University, Beijing, 100083, China
ABSTRACT
all-trans retinoic acid (atRA), the oxidative metabolite of vitamin A, is essential for normal embryonic development. Also, high levels of atRA are teratogenic in many species and can effectively induce cleft palate in the mouse. Most cleft palate resulted from the failed fusion of secondary palate shelves, and maintenance of the normal cell proliferation is important in this process of shelf growth. To clarify the mechanism by which atRA causes cleft palate, we investigated the effect of atRA on proliferation activity and cell cycle distribution in mouse embryonic palatal mesenchymal (MEPM) cells. atRA inhibited the growth of MEPM cells by inducing apoptosis in a dose-dependent manner. atRA also caused a G1 block in the cell cycle with an increase in the proportion of cells in G0/G1 and a decrease in the proportion of cells in S phase, as determined by flow cytometry. We next investigated the effects of atRA on molecules that regulate the G1 to S phase transition. These studies demonstrated that atRA inhibited expression of cyclins D and E at the protein level. Furthermore, atRA treatment reduced phosphorylated Rb and decreased cdk2 and cdk4 kinase activity. These data suggest that atRA had antiproliferative activity by modulating G1/S cell cycle regulators and by inhibition of Rb phosphorylation in MEPM cells, which might account for the pathogenesis of cleft palate induced by retinoic acid.
Key Words: all-trans retinoic acid; mouse embryonic palatal mesenchymal cells; cell cycle; cyclins; retinoblastoma protein.
INTRODUCTION
In the mouse, secondary palate shelves arise from the developing maxillary process on gestation day (GD) 12.5 by initially growing in a vertical position alongside the tongue and then undergoing rapid elevation to the horizontal position above the tongue, which brings opposing shelves into contact at the midline on GD 14.5 (Eldeib and Reddy, 1988; Ferguson, 1988). At this time, the palate shelves are ready to make contact with each other and thereupon to fuse. During this process of palatogenesis, maintenance of normal proliferation of palatal mesenchymal cells is very important. It is widely believed that one of the primary causes of cleft palate is failure of palatal processes to elevate as a result of growth inhibition. Control of this process at the cellular level is likely to be mediated by specific intracellular signal transduction mechanisms and cell-cycle inhibitory growth factors secreted in response to various teratogenic microenvironments (Dhulipala et al., 2004).
Retinoic acid, especially all-trans retinoic acid (atRA) plays an important role in embryogenesis, by regulating morphogenesis, cell proliferation, differentiation, and extracellular matrix production (Lai et al., 2003; Sasaki et al., 2004; Shibamoto et al., 2004; Wang and Kirsch, 2002). It is also teratogenic at high concentrations. The induction of cleft palate by atRA varies depending on the stage of development exposed. In vivo studies have indicated that, after exposure of embryonic mice to RA on GD 10, abnormally small palatal shelves form. After exposure on GD 12, shelves of normal size form, but fail to fuse, as the medial cells proliferate and the normal growth and differentiation process of palatal mesenchymal cells is inhibited. Because reduced proliferation of palatal mesenchymal cells can contribute to smaller shelf size, the hypothesis that atRA may have adverse effects on this cellular processes was tested in the present study, using primary mouse embryonic palate mesenchymal cell (MEPM) cultures.
Previous studies have indicated that retinoids suppress cell growth by several cellular mechanisms. They inhibit cell proliferation by inducing G1 arrest in many different cell types. The cell cycle in eukaryotes is regulated by cyclin-dependent kinases (cdk). The cyclins, members of the cell cycle regulators, bind to and activate cdk. Activation of cyclin/cdk is required for cell cycle progression and G1/S transition in response to microenvironments. During progression through G1, the amount of D-type cyclin increases, and, in a mitogen-regulated manner, these proteins associate with and activate cdk4 during early-to-mid G1 phase, which phosphorylates Rb on specific residues (including Ser780) (Connell-Crowley et al., 1997; Pan et al., 2001; Zarkowska and Mittnacht, 1997). This releases the activity of E2F toward the cyclin E gene promoter, and consequently, the expression of cyclin E activates cdk2, completing the hyperphosphorylation of Rb (Geng et al., 1996; Kitagawa et al., 1996; Lundberg and Weinberg, 1998; Weinberg, 1995) and allowing the G1/S transition (Donnellan and Chetty, 1999; Harbour and Dean, 2000; Ohtani, 1999; Ohtsubo et al., 1995). Thus, pRb is critical for entry into S phase.
In this study, we evaluated the effect of atRA on cell growth and cell cycle distribution in MEPM cells, and clarified the underlying mechanisms; we investigated the effect of atRA on the above-mentioned molecular regulators, which mainly account for cell proliferation and cell cycle progression. Our data show that atRA has antiproliferative activity by modulating G1/S cell cycle regulators and by inhibition of Rb phosphorylation. These events might account for the pathogenesis of cleft palate induced by retinoic acid.
MATERIALS AND METHODS
Animals. Mature male and female ICR mice were housed at a temperature of 22°C with a 12 h light/dark cycle. Animals were maintained on Purina mouse chow and water ad libitum. Matings were accomplished by placing 1 male and 3 females together at 21:00 and allowing them to mate overnight. The presence of a vaginal plug the following morning was taken as evidence of mating (gestation day 0; GD 0).
Cell culture. Pregnant mice were euthanized on GD 13, a critical stage of murine secondary palate development. Embryos were removed from pregnant dams. The palate shelves were dissected in sterile, cold phosphate-buffered saline (PBS) and were pooled, minced, and converted into single cell suspensions by incubating with 0.25% trypsin/0.05% EDTA in PBS for 10 min at 37°C with constant shaking. Digested samples were briefly triturated and filtered through 70-μm mesh, and cells were seeded on cell-culture dishes and grown to confluence in DMEM medium (Gibco BRL) containing 5% fetal calf serum (FBS, Hyclon Co.) at 37°C in a 95% air/5% CO2 atmosphere, with media replaced every other day.
Trypan blue exclusion assay. Cells were seeded at an initial concentration of 2 x 104 cells/ml cells in 24-well tissue culture plates, and atRA was added to a series of concentrations (0, 0.1, 1, 5, and 10 μM, respectively). The final concentration of vehicle ethanol in all treatments did not exceed 0.1% (v/v) in the medium. Each treatment and time point had four plates. After 24 h, 48 h, and 72 h, cells were collected by trypsinization and counted in duplicate with a hemocytometer. Trypan blue dye exclusion was used to determine a cell growth curve.
MTT assay. The effect of atRA on cell viability was determined by MTT assay in MEPM cells. Briefly, cells (104 cells per well) were seeded in 96-well microtiter plates (Nunc, Denmark). After exposure to various concentrations of atRA for 72 h, 50 μl MTT solution (Sigma) (2 mg/ml in PBS) was added to each well, and the plates were incubated for additional 4 h at 37°C. MTT solution in medium was aspirated off. To achieve solubilization of the formazan crystal formed in viable cells, 200 μl DMSO was added to each well. The absorbance was read at 540 nm on a Dias automatic microwell plate reader with DMSO as the blank.
Flow cytometry. Cell cycle distribution and subdiploid fraction were determined by staining DNA with PI (propidium iodide) (Sigma). Briefly, 1 x 105 cells were seeded in a 6-well plate and incubated with given concentrations of atRA for 72 h. Cells were then washed in PBS, then fixed in 70% ice ethanol overnight, after which they were resuspended in PBS containing 40 μg/ml PI, 0.1 mg/ml RNase (Sigma), and 0.1% Triton X-100 at 37°C for 30 min. Then the cell cycle distribution and subdiploid population were detected with flow cytometery (FACScan; Becton-Dickinson) and analyzed by using a ModFit software package (Becton–Dickinson).
Western blot analysis. After MEPM cells were seeded in 75 ml flasks and treated with atRA for 72 h, the cells were harvested and lysed in a buffer containing 50 mM Tris-HCl, pH 6.8; 10% glycerol; 2% sodium dodecyl sulfate (SDS); and 5% b-mercaptoethanol. Then 25 μg protein lysates were separated by 12.5% SDS-PAGE (polyacrylamide gel electrophoresis) and transferred onto nitrocellulose. After blocking in a 5% non-fat dry milk solution in washing buffer containing 10 mmol/l Tris (pH 7.5), 150 mmol/l NaCl, and 0.05% Tween-20, membranes were incubated overnight at 4°C with different antibodies: rabbit polyclonal anti-cyclin D (Santa Cruz), rabbit polyclonal anti-cyclin E (Santa Cruz), rabbit polyclonal antibody recognizing phospho-Rb-Ser795 (Cell Signaling Technology), and rabbit polyclonal anti--actin (Santa Cruz). After they were washed three times with Tween 20-PBS, membranes were incubated for 2 h with horseradish peroxidase–coupled secondary antibodies at room temperature. Signals were detected with the ECL kit (Amersham Pharmacia Biotech).
Kinase reaction assay. MEPM cells were treated by atRA and lysed in the same way as for western blot analysis. 100 μg samples of cellular protein were incubated with 1 μg of either anti-cdk2 antibody or anti-cdk4 antibody (Santa Cruz) for 2 h at 4°C. Then, 20 μl of protein A/G agarose (Santa Cruz) was added and incubation was carried out for an additional 1 h. Beads were then pelleted by centrifugation and washed three times with lysis buffer and two times with PBS. To initiate the reaction, 30 μl of kinase buffer (50 mM Hepes (pH 7.5), 10 mM MgCl2, 5 mM MnCl2, 1 mM NaF, 0.1 mM Na3VO4, 1 mM PMSF, 1 mM dithiothreitol, 1 μg/ml leupeptin, 0.2 μg/ml aprotinin) with 2.5 mM EGTA, 50 μM cold ATP, 0.2 mg/ml BSA, 10 μCi of -32P-ATP, and 1 μg of GST-Rb (for cdk4) or 2 μg histone H1 (for cdk2) was then added to the pelleted beads, and the reaction was incubated at 30°C for 30 min with frequent agitation. The reaction was stopped by addition of 8 μl of 6x sample buffer and boiling for 5 min. The products were analyzed by 12% SDS-PAGE electrophoresis and visualized by autoradiography.
Statistical analysis. All data are presented as the mean ± SD (standard deviation). The data were evaluated by a one-way analysis of variance (ANOVA) followed by the least significant difference (LSD) test as a post hoc test or Dunnett's T3 test with the SPSS 10.0 program (SPSS Inc. Chicago, IL). Statistical significance was at P < 0.05.
RESULTS
atRA Inhibited MEPM Cell Growth
The effects of atRA on the growth of MEPM cells were analyzed with the trypan blue dye exclusion test. atRA inhibited the growth of MEPM cells dose dependently, ranging from 0.1 to 10 μM (Fig. 1), and showed significant inhibition at concentrations of 5 and 10 μM after atRA treatment for 24 h, 48 h, and 72 h (P < 0.05).
Effect of atRA on Cell Viability in MEPM Cells
In this set of experiments, we evaluated the effects of atRA on cell viability in MEPM cells. Cells were treated with different concentrations of atRA for 72 h, and cell viability was determined by MTT assay. A significant inhibition in cell viability was observed after treatment with atRA (Fig. 2). Indeed, 5 and 10 μM atRA inhibited cell viable numbers by 42.6% and 54.8%, respectively, compared with control cells that were treated only with vehicle (indicated as 0 μM).
atRA-Induced Cell Cycle Arrest and Apoptosis in MEPM Cells
In light of the fact that proliferation of MEPM cells was inhibited by high concentrations of atRA, we next used flow cytometry to determine where in the cell cycle they accumulate and whether apoptotic events occurred with them in response to atRA. As shown in Figure 3, we found that in control (vehicle-treated) MEPM cells, 53% of the cells were in G0/G1, 29% in S phase, and 18% in G2/M phase. Upon given concentrations of atRA treatment, an increase in the number of cells in G0/G1 was observed in a dose-responsive manner. Indeed, at the concentration of 10 μM, approximately 80% of cells were in G0/G1, whereas the numbers of cells in S and G2/M were reduced to 12% and 8%, respectively (Fig. 3). Thus, atRA treatment leads to G0/G1 arrest in MEPM cells in a dose-dependent manner. At concentrations of 5 μM and 10 μM, atRA treatment induced the appearance of a sub-G1 fraction (17% and 26%, respectively), which is characteristic of apoptosis.
Effect of atRA on Expression of Cell Cycle Regulatory Molecules
We next determined whether atRA affected expression of molecules that are known to regulate entry of cells into the S phase and thereby play key roles in MEPM cell growth, inhibition, and cell cycle progression. These include cyclins D and E, which govern phosphorylation of Rb, the release of E2F, and the control of exit from G1 (Sherr, 1996). As shown in Figure 4, atRA treatment decreased the expression levels of cyclin D1 and E in a dose-responsive manner. When we evaluated the levels of phosphorylated Rb with a specific anti–phospho-Rb antibody (phospho Ser 795), we observed a dose-dependent inhibition of phosphorylation of this critical regulator of G1/S progression.
Alterations in Cell-Cycle Enzyme Activity of cdk2 and cdk4
To further understand the mechanism by which atRA inhibit Rb phosphorylation, we examined their effect on the activity of cdks. cdk4 is active during early-to-mid G1 phase and Cdk2 is active in late G1 phase; both were required for progression through the G1/S transition through phosphorylating a critical serine residue of Rb (Ser 795) (Koff et al., 1992; Rosenblatt et al., 1992; Swanton, 2004; van den Heuvel and Harlow 1993). The present data show that within the range of concentrations from 0.1 μM to 10 μM, atRA inhibited reaction activity of both cdk2 and cdk4 (Fig. 5).
DISCUSSION
The physiologic doses (usually 0.01 μM) of RA play a important role in embryogenesis. The unsuitable maternal use of certain retinoid-containing medications and excessive supplements at specific critical periods of facial morphogenesis could lead to aberrant development and subsequent cleft palate. The greatest success in the RA-induced cleft palate in mice was demonstrated by a single dose of exogenous atRA on GD 11 at a level of 100 mg/kg. Given the absence of maternal toxicity, it is evident that all pharmacological doses (1 μM) of RA are at the toxic thresholds. To gain insight into possible mechanisms of atRA-induced cleft palate, we mainly studied the effects of pharmacologic (1 μM) doses of atRA on growth of MEPM cells.
In atRA-treated MEPM cultures, we found a dose-dependent survival suppression that blocked cell cycle progression at the G1/S transition, ultimately leading to G1/G0 arrest and eliciting apoptosis. Data here are in agreement with a previous in vivo study, which showed that the inhibition of growth and excess apoptosis of MEPM cells contributed to the formation of cleft palate and other orofacial congenital malformations (Suwa et al., 2001). It is well known that atRA treatment induced decrease of cell viability and increase of apoptotic phenomena in diverse tumor cell types, including cancers of the breast, lung, liver, and blood (Guzey et al., 1998; Hsu et al., 2001; Jimenez-Lara et al., 2004; Mangiarotti et al., 1998). Some teratogenic effects of retinoids may be due to their ability to inhibit cell growth and induce apoptosis. For example, RA treatment of midgestation mouse embryos induces excess limb bud apoptosis, resulting in malformed limbs (Jiang and Kochhar, 1992). Cleft palate results from failure of fusion between the palatal shelves. One possible cause is retarded mesenchymal proliferation and differentiation in those shelves, which will lead to retardation in the development of the palatal bones (Furukawa et al., 2004; Stark, 1954).
Proliferating mammalian cells pass through several cell cycle checkpoints, mainly G1-to-S and G2-to-M transitions. Three important targets have emerged as likely candidates for effectors of this outcome including the expression of cyclin D1 and E and the phosphorylated status of Rb. Data here show that atRA induced a dose-dependent decrease in cyclins D and E. During early G1 phase, cyclins D bind to and activates cdk 4, cyclin D–cdk4 complexes phosphorylate their primary target protein, the tumor suppressor retinoblastoma (Rb) protein. In late G1, cyclins E interact with and activate cdk2. Cyclin E–Cdk2 complexes complete this phosphorylation (Duman-Scheel et al., 2002; Yamada et al., 2004). Hyperphosphorylated Rb no longer binds to its target, E2F, thereby allowing E2F to activate the transcription of genes required for S phase in several consecutive steps and thereby ultimately relieving Rb-mediated repression from S phase, as Rb becomes dephosphorylated just prior to M/G1 transition (Bai et al., 2003; DiPietrantonio et al., 1998; Johnson and Schneider-Broussard, 1998; Sherr, 2000). Given the importance of Rb, it is reasonable that inhibition of phosphorylation of Rb is thought to be one of the important mechanisms by which retinoids inhibit proliferation in MEPM cells.
The formation of secondary palate involves several ordered steps, starting with shelf growth and elevation, and followed by contact, adhesion, and disappearance of the medial edge epithelium. Disruption at any stage of palate development could potentially cause cleft palate. It has been known for some time that maternal treatment of atRA produces embryonic palatal shelves deficient in mesenchymal tissue (Horie and Yasuda, 2001; Ikemi et al., 2001) and exhibiting a decreased rate of 3H-thymidine uptake, indicative of inhibition of cell proliferation (Watanabe et al., 1990). Retinoic acid has also been shown to inhibit proliferation of, and extracellular matrix synthesis in mouse fetal palate mesenchymal cells. These data suggest that perturbation of the process of cell proliferation might account for the formation of small palatal shelves and the failure of palatal fusion in vivo (Watanabe et al., 1988).
Together, these results are consistent with the hypothesis that retinoid-induced cleft palate might depend in part on the effect of RA on growth inhibition and modification of cell cycle distribution of palate mesenchymal cells in the developing palatal shelves. We proposed that decreased cdk activity, followed by reduced phosphorylation of Rb, may account for the failure of palate fusion exposed to RA as the elements of the secondary palate are just beginning to make contract.
ACKNOWLEDGMENTS
This work was supported by National Natural Science Foundation grant nos. 30371224 and 30371559, People's Republic of China, and the Major Basic Research Development Program of the People's Republic of China (2001CB510305).
NOTES
REFERENCES
Bai, S., Goodrich, D., Thron, C. D., Tecarro, E., and Obeyesekere, M. (2003). Theoretical and experimental evidence for hysteresis in cell proliferation. Cell Cycle 2, 46–52.
Connell-Crowley, L., Harper, J. W., and Goodrich, D. W. (1997). Cyclin D1/Cdk4 regulates retinoblastoma protein-mediated cell cycle arrest by site specific phosphorylation. Mol. Biol. Cell. 8, 287–301.
Dhulipala, V. C., Hanumegowda, U. M., Balasubramanian, G., and Reddy, C. S. (2004). Relevance of the palatal protein kinase A pathway to the pathogenesis of cleft palate by secalonic acid D in mice. Toxicol. Appl. Pharmacol. 194, 270–279.
DiPietrantonio, A. M., Hsieh, T. C., Olson, S. C., and Wu, J. M. (1998). Regulation of G1/S transition and induction of apoptosis in HL-60 leukemia cells by fenretinide (4HPR). Int. J. Cancer 78, 53–61.
Donnellan, R., and Chetty, R. (1999). Cyclin E in human cancers. FASEB J. 13, 773–780.
Duman-Scheel, M., Weng, L., Xin, S., and Du, W. (2002). Hedgehog regulates cell growth and proliferation by inducing cyclin D and cyclin E. Nature 417, 299–304.
Eldeib, M. M. R., and Reddy, C. S. (1988). Secalonic acid D–induced changes in palatal cyclic AMP and cyclic GMP in developing mice. Teratology 37, 343–352.
Ferguson, M. W. (1988). Palate development. Development 103(Suppl), 41–60.
Furukawa, S., Usuda, K., Abe, M., and Ogawa, I. (2004). Histopathological findings of cleft palate in rat embryos induced by triamcinolone acetonide. J. Vet. Med. Sci. 66, 7–402.
Geng, Y., Eaton, E. N., Picon, M., Roberts, J. M., Lundberg, A. S., Gifford, A., Sardet, C., and Weinberg, R. A. (1996). Regulation of cyclin E transcription by E2Fs and retinoblastoma protein. Oncogene 12, 1173–1180.
Guzey, M., Demirpence, E., Criss, W., and DeLuca, H. F. (1998). Effects of retinoic acid (all-trans and 9-cis) on tumor progression in small cell lung carcinoma. Biochem. Biophys. Res. Commun. 242, 369–375.
Harbour, J. W., and Dean D. C. (2000). The Rb/E2F pathway: expanding roles and emerging paradigms. Genes Dev. 14, 2393–409.
Horie, S., and Yasuda, M. (2001). Alterations in palatal ruga patterns in Jcl:ICR mouse fetuses from dams treated with all-trans-retinoic acid. Hiroshima J. Med. Sci. 50, 17–25.
Hsu, S. L., Cheng, C. C., Shi, Y. R., and Chiang, C. W. (2001). Proteolysis of integrin alpha5 and beta1 subunits involved in retinoic acid-induced apoptosis in human hepatoma Hep3B cells. Cancer Lett. 167, 193–204.
Ikemi, N., Otani, Y., Ikegami, T., and Yasuda, M. (2001). Palatal ruga anomaly induced by all-trans-retinoic acid in the Crj:SD rat: Possible warning sign of teratogenicity. Reprod. Toxicol. 15, 87–93.
Jiang, H., and Kochhar, D. M. (1992). Induction of tissue transglutaminase and apoptosis by retinoic acid in the limb bud. Teratolog. 46, 333–340.
Jimenez-Lara, A. M., Clarke, N., Altucci, L., and Gronemeyer, H. (2004). Retinoic-acid-induced apoptosis in leukemia cells. Trends Mol. Med. 10, 508–515
Johnson, D. G., and Schneider-Broussard, R. (1998). Role of E2F in cell cycle control and cancer. Frontiers Biosci. 27, 447–8.
Kitagawa, M., Higashi, H., Jung, H. K, Suzuki-Takahashi, I., Ikeda, M., Tamai, K., Kato, J., Segawa, K., Yoshida, E., Nishimura, S., and Taya, Y. (1996). The consensus motif for phosphorylation by cyclin D1-Cdk4 is different from that for phosphorylation by cyclin A/E-Cdk2. EMBO J. 15, 7060–7069.
Koff, A. A., Giordano, D., Desai, K., Yamashita, J. W., Harper, S., Elledge, T., Nishimoto, D. O., Morgan, B. R., Franza, B. R., and Roberts, J. M. (1992). Formation and activation of a cyclin E-cdk2 complex during G1 phase of the human cell cycle. Science 257, 1689–1694.
Lai, L., Bohnsack, B. L., Niederreither, K., and Hirschi, K. K. (2003). Retinoic acid regulates endothelial cell proliferation during vasculogenesis. Development 130, 6465–6474.
Lundberg, A. S., and Weinberg, R. A. (1998). Functional inactivation of the retinoblastoma protein requires sequential medication by at least two distinct cyclin-cdk complexes. Mol. Cell Biol. 18, 753–761.
Mangiarotti, R., Danova, M., Alberici, R., and Pellicciari, C. (1998). All-trans retinoic acid (ATRA)-induced apoptosis is preceded by G1 arrest in human MCF-7 breast cancer cells. Br. J. Cancer 77, 186–191.
Ohtani, K. (1999). Implication of transcription factor E2F in regulation of DNA replication. Frontiers Biosci. 4, D793–804.
Ohtsubo, M., Theodoras, A. M., Schumacher, J., Roberts, J. M., and Pagano, M. (1995). Human cyclin E, a nuclear protein essential for the G1-to-S phase transition. Mol. Cell Biol. 15, 2612–2624.
Pan, W., Cox, S., Hoess, R. H., and Grafstrom, R. H. (2001). A cyclin D1/cyclin dependent kinase 4 binding site within the C domain of the retinoblastoma protein. Cancer Res. 61, 2885–2891.
Rosenblatt, J., Gu, Y., and Morgan, D. O. (1992). Human cyclin dependent kinase 2 is activated during the S and G2 phases of the cell cycle and associates with cyclin A. Proc. Natl. Acad. Sci. U.S.A. 89, 2824–2828.
Sasaki, Y., Iwai, N., Tsuda, T., and Kimura, O. (2004). Sonic hedgehog and Bone morphogenetic protein 4 expressions in the hindgut region of murine embryo with anorectal malformations. J. Pediatr. Surg. 39, 170–173.
Shibamoto, S., Winer, J., Williams, M., and Polakis, P. (2004). A blockade in Wnt signaling is activated following the differentiation of F9 teratocarcinoma cells. Exp. Cell Res. 292, 11–20.
Sherr, C. J. (1996). Cancer cell cycles. Science 274, 1672–1677
Sherr, C. J. (2000). The Pezcoller Lecture: Cancer cell cycles revisited. Cancer Res. 60, 3689–3695.
Stark, R. B. (1954). The pathogenesis of harelip and cleft palate. Plast. Reconstruct. Surg. 13, 20–32.
Suwa, F., Jin, Y., Lu, H., Li, X., Tipoe, G. L., Lau, T. Y., Tamada, Y., Kuroki, K., and Fang, Y. R. (2001). Alteration of apoptosis in cleft palate formation and ectomesenchymal stem cells influenced by retinoic acid. Okajimas Folia. Anat. Jpn. 78, 179–186.
Swanton, C. (2004). Cell-cycle targeted therapies. Lancet Oncol. 5, 27–36.
van den Heuvel, S., E., and Harlow, E. (1993). Distinct roles for cyclin-dependent kinases in cell cycle control. Science 262, 2050–2054.
Wang, W., and Kirsch, T. (2002). Retinoic acid stimulates annexin-mediated growth plate chondrocyte mineralization. J. Cell Biol. 157, 1061–1069.
Watanabe, T., Goulding, E. H., and Pratt, R. M. (1988). Alterations in craniofacial growth induced by isotretinoin (13-cis-retinoic acid) in mouse whole embryo and primary mesenchymal cell culture. J. Craniofac. Genet. Dev. Biol. 8, 21–33.
Watanabe, T., Willis, W. D., and Pratt, R. M. (1990). Effect of retinoids on proliferation of human embryonic palatal mesenchymal cells in culture. J. Nutr. Sci. Vitaminol. (Tokyo) 36, 311–325.
Weinberg, R. A. (1995). The retinoblastoma protein and cell cycle control. Cell 81, 323–330.
Yamada, S., Sumrejkanchanakij, P., Amagasa, T., and Ikeda, M. A. (2004). Loss of cyclin E requirement in cell growth of an oral squamous cell carcinoma cell line implies deregulation of its downstream pathway. Int. J. Cancer 111, 17–22.
Zarkowska, T., and Mittnacht, S. (1997). Differential phosphorylation of the retinoblastoma protein by G1/S cyclin-dependent kinases. J. Biol. Chem. 272, 12738–12746.(Zengli Yu, Jiuxiang Lin, )
School of Public Health, Peking University, Beijing, 100083, China
ABSTRACT
all-trans retinoic acid (atRA), the oxidative metabolite of vitamin A, is essential for normal embryonic development. Also, high levels of atRA are teratogenic in many species and can effectively induce cleft palate in the mouse. Most cleft palate resulted from the failed fusion of secondary palate shelves, and maintenance of the normal cell proliferation is important in this process of shelf growth. To clarify the mechanism by which atRA causes cleft palate, we investigated the effect of atRA on proliferation activity and cell cycle distribution in mouse embryonic palatal mesenchymal (MEPM) cells. atRA inhibited the growth of MEPM cells by inducing apoptosis in a dose-dependent manner. atRA also caused a G1 block in the cell cycle with an increase in the proportion of cells in G0/G1 and a decrease in the proportion of cells in S phase, as determined by flow cytometry. We next investigated the effects of atRA on molecules that regulate the G1 to S phase transition. These studies demonstrated that atRA inhibited expression of cyclins D and E at the protein level. Furthermore, atRA treatment reduced phosphorylated Rb and decreased cdk2 and cdk4 kinase activity. These data suggest that atRA had antiproliferative activity by modulating G1/S cell cycle regulators and by inhibition of Rb phosphorylation in MEPM cells, which might account for the pathogenesis of cleft palate induced by retinoic acid.
Key Words: all-trans retinoic acid; mouse embryonic palatal mesenchymal cells; cell cycle; cyclins; retinoblastoma protein.
INTRODUCTION
In the mouse, secondary palate shelves arise from the developing maxillary process on gestation day (GD) 12.5 by initially growing in a vertical position alongside the tongue and then undergoing rapid elevation to the horizontal position above the tongue, which brings opposing shelves into contact at the midline on GD 14.5 (Eldeib and Reddy, 1988; Ferguson, 1988). At this time, the palate shelves are ready to make contact with each other and thereupon to fuse. During this process of palatogenesis, maintenance of normal proliferation of palatal mesenchymal cells is very important. It is widely believed that one of the primary causes of cleft palate is failure of palatal processes to elevate as a result of growth inhibition. Control of this process at the cellular level is likely to be mediated by specific intracellular signal transduction mechanisms and cell-cycle inhibitory growth factors secreted in response to various teratogenic microenvironments (Dhulipala et al., 2004).
Retinoic acid, especially all-trans retinoic acid (atRA) plays an important role in embryogenesis, by regulating morphogenesis, cell proliferation, differentiation, and extracellular matrix production (Lai et al., 2003; Sasaki et al., 2004; Shibamoto et al., 2004; Wang and Kirsch, 2002). It is also teratogenic at high concentrations. The induction of cleft palate by atRA varies depending on the stage of development exposed. In vivo studies have indicated that, after exposure of embryonic mice to RA on GD 10, abnormally small palatal shelves form. After exposure on GD 12, shelves of normal size form, but fail to fuse, as the medial cells proliferate and the normal growth and differentiation process of palatal mesenchymal cells is inhibited. Because reduced proliferation of palatal mesenchymal cells can contribute to smaller shelf size, the hypothesis that atRA may have adverse effects on this cellular processes was tested in the present study, using primary mouse embryonic palate mesenchymal cell (MEPM) cultures.
Previous studies have indicated that retinoids suppress cell growth by several cellular mechanisms. They inhibit cell proliferation by inducing G1 arrest in many different cell types. The cell cycle in eukaryotes is regulated by cyclin-dependent kinases (cdk). The cyclins, members of the cell cycle regulators, bind to and activate cdk. Activation of cyclin/cdk is required for cell cycle progression and G1/S transition in response to microenvironments. During progression through G1, the amount of D-type cyclin increases, and, in a mitogen-regulated manner, these proteins associate with and activate cdk4 during early-to-mid G1 phase, which phosphorylates Rb on specific residues (including Ser780) (Connell-Crowley et al., 1997; Pan et al., 2001; Zarkowska and Mittnacht, 1997). This releases the activity of E2F toward the cyclin E gene promoter, and consequently, the expression of cyclin E activates cdk2, completing the hyperphosphorylation of Rb (Geng et al., 1996; Kitagawa et al., 1996; Lundberg and Weinberg, 1998; Weinberg, 1995) and allowing the G1/S transition (Donnellan and Chetty, 1999; Harbour and Dean, 2000; Ohtani, 1999; Ohtsubo et al., 1995). Thus, pRb is critical for entry into S phase.
In this study, we evaluated the effect of atRA on cell growth and cell cycle distribution in MEPM cells, and clarified the underlying mechanisms; we investigated the effect of atRA on the above-mentioned molecular regulators, which mainly account for cell proliferation and cell cycle progression. Our data show that atRA has antiproliferative activity by modulating G1/S cell cycle regulators and by inhibition of Rb phosphorylation. These events might account for the pathogenesis of cleft palate induced by retinoic acid.
MATERIALS AND METHODS
Animals. Mature male and female ICR mice were housed at a temperature of 22°C with a 12 h light/dark cycle. Animals were maintained on Purina mouse chow and water ad libitum. Matings were accomplished by placing 1 male and 3 females together at 21:00 and allowing them to mate overnight. The presence of a vaginal plug the following morning was taken as evidence of mating (gestation day 0; GD 0).
Cell culture. Pregnant mice were euthanized on GD 13, a critical stage of murine secondary palate development. Embryos were removed from pregnant dams. The palate shelves were dissected in sterile, cold phosphate-buffered saline (PBS) and were pooled, minced, and converted into single cell suspensions by incubating with 0.25% trypsin/0.05% EDTA in PBS for 10 min at 37°C with constant shaking. Digested samples were briefly triturated and filtered through 70-μm mesh, and cells were seeded on cell-culture dishes and grown to confluence in DMEM medium (Gibco BRL) containing 5% fetal calf serum (FBS, Hyclon Co.) at 37°C in a 95% air/5% CO2 atmosphere, with media replaced every other day.
Trypan blue exclusion assay. Cells were seeded at an initial concentration of 2 x 104 cells/ml cells in 24-well tissue culture plates, and atRA was added to a series of concentrations (0, 0.1, 1, 5, and 10 μM, respectively). The final concentration of vehicle ethanol in all treatments did not exceed 0.1% (v/v) in the medium. Each treatment and time point had four plates. After 24 h, 48 h, and 72 h, cells were collected by trypsinization and counted in duplicate with a hemocytometer. Trypan blue dye exclusion was used to determine a cell growth curve.
MTT assay. The effect of atRA on cell viability was determined by MTT assay in MEPM cells. Briefly, cells (104 cells per well) were seeded in 96-well microtiter plates (Nunc, Denmark). After exposure to various concentrations of atRA for 72 h, 50 μl MTT solution (Sigma) (2 mg/ml in PBS) was added to each well, and the plates were incubated for additional 4 h at 37°C. MTT solution in medium was aspirated off. To achieve solubilization of the formazan crystal formed in viable cells, 200 μl DMSO was added to each well. The absorbance was read at 540 nm on a Dias automatic microwell plate reader with DMSO as the blank.
Flow cytometry. Cell cycle distribution and subdiploid fraction were determined by staining DNA with PI (propidium iodide) (Sigma). Briefly, 1 x 105 cells were seeded in a 6-well plate and incubated with given concentrations of atRA for 72 h. Cells were then washed in PBS, then fixed in 70% ice ethanol overnight, after which they were resuspended in PBS containing 40 μg/ml PI, 0.1 mg/ml RNase (Sigma), and 0.1% Triton X-100 at 37°C for 30 min. Then the cell cycle distribution and subdiploid population were detected with flow cytometery (FACScan; Becton-Dickinson) and analyzed by using a ModFit software package (Becton–Dickinson).
Western blot analysis. After MEPM cells were seeded in 75 ml flasks and treated with atRA for 72 h, the cells were harvested and lysed in a buffer containing 50 mM Tris-HCl, pH 6.8; 10% glycerol; 2% sodium dodecyl sulfate (SDS); and 5% b-mercaptoethanol. Then 25 μg protein lysates were separated by 12.5% SDS-PAGE (polyacrylamide gel electrophoresis) and transferred onto nitrocellulose. After blocking in a 5% non-fat dry milk solution in washing buffer containing 10 mmol/l Tris (pH 7.5), 150 mmol/l NaCl, and 0.05% Tween-20, membranes were incubated overnight at 4°C with different antibodies: rabbit polyclonal anti-cyclin D (Santa Cruz), rabbit polyclonal anti-cyclin E (Santa Cruz), rabbit polyclonal antibody recognizing phospho-Rb-Ser795 (Cell Signaling Technology), and rabbit polyclonal anti--actin (Santa Cruz). After they were washed three times with Tween 20-PBS, membranes were incubated for 2 h with horseradish peroxidase–coupled secondary antibodies at room temperature. Signals were detected with the ECL kit (Amersham Pharmacia Biotech).
Kinase reaction assay. MEPM cells were treated by atRA and lysed in the same way as for western blot analysis. 100 μg samples of cellular protein were incubated with 1 μg of either anti-cdk2 antibody or anti-cdk4 antibody (Santa Cruz) for 2 h at 4°C. Then, 20 μl of protein A/G agarose (Santa Cruz) was added and incubation was carried out for an additional 1 h. Beads were then pelleted by centrifugation and washed three times with lysis buffer and two times with PBS. To initiate the reaction, 30 μl of kinase buffer (50 mM Hepes (pH 7.5), 10 mM MgCl2, 5 mM MnCl2, 1 mM NaF, 0.1 mM Na3VO4, 1 mM PMSF, 1 mM dithiothreitol, 1 μg/ml leupeptin, 0.2 μg/ml aprotinin) with 2.5 mM EGTA, 50 μM cold ATP, 0.2 mg/ml BSA, 10 μCi of -32P-ATP, and 1 μg of GST-Rb (for cdk4) or 2 μg histone H1 (for cdk2) was then added to the pelleted beads, and the reaction was incubated at 30°C for 30 min with frequent agitation. The reaction was stopped by addition of 8 μl of 6x sample buffer and boiling for 5 min. The products were analyzed by 12% SDS-PAGE electrophoresis and visualized by autoradiography.
Statistical analysis. All data are presented as the mean ± SD (standard deviation). The data were evaluated by a one-way analysis of variance (ANOVA) followed by the least significant difference (LSD) test as a post hoc test or Dunnett's T3 test with the SPSS 10.0 program (SPSS Inc. Chicago, IL). Statistical significance was at P < 0.05.
RESULTS
atRA Inhibited MEPM Cell Growth
The effects of atRA on the growth of MEPM cells were analyzed with the trypan blue dye exclusion test. atRA inhibited the growth of MEPM cells dose dependently, ranging from 0.1 to 10 μM (Fig. 1), and showed significant inhibition at concentrations of 5 and 10 μM after atRA treatment for 24 h, 48 h, and 72 h (P < 0.05).
Effect of atRA on Cell Viability in MEPM Cells
In this set of experiments, we evaluated the effects of atRA on cell viability in MEPM cells. Cells were treated with different concentrations of atRA for 72 h, and cell viability was determined by MTT assay. A significant inhibition in cell viability was observed after treatment with atRA (Fig. 2). Indeed, 5 and 10 μM atRA inhibited cell viable numbers by 42.6% and 54.8%, respectively, compared with control cells that were treated only with vehicle (indicated as 0 μM).
atRA-Induced Cell Cycle Arrest and Apoptosis in MEPM Cells
In light of the fact that proliferation of MEPM cells was inhibited by high concentrations of atRA, we next used flow cytometry to determine where in the cell cycle they accumulate and whether apoptotic events occurred with them in response to atRA. As shown in Figure 3, we found that in control (vehicle-treated) MEPM cells, 53% of the cells were in G0/G1, 29% in S phase, and 18% in G2/M phase. Upon given concentrations of atRA treatment, an increase in the number of cells in G0/G1 was observed in a dose-responsive manner. Indeed, at the concentration of 10 μM, approximately 80% of cells were in G0/G1, whereas the numbers of cells in S and G2/M were reduced to 12% and 8%, respectively (Fig. 3). Thus, atRA treatment leads to G0/G1 arrest in MEPM cells in a dose-dependent manner. At concentrations of 5 μM and 10 μM, atRA treatment induced the appearance of a sub-G1 fraction (17% and 26%, respectively), which is characteristic of apoptosis.
Effect of atRA on Expression of Cell Cycle Regulatory Molecules
We next determined whether atRA affected expression of molecules that are known to regulate entry of cells into the S phase and thereby play key roles in MEPM cell growth, inhibition, and cell cycle progression. These include cyclins D and E, which govern phosphorylation of Rb, the release of E2F, and the control of exit from G1 (Sherr, 1996). As shown in Figure 4, atRA treatment decreased the expression levels of cyclin D1 and E in a dose-responsive manner. When we evaluated the levels of phosphorylated Rb with a specific anti–phospho-Rb antibody (phospho Ser 795), we observed a dose-dependent inhibition of phosphorylation of this critical regulator of G1/S progression.
Alterations in Cell-Cycle Enzyme Activity of cdk2 and cdk4
To further understand the mechanism by which atRA inhibit Rb phosphorylation, we examined their effect on the activity of cdks. cdk4 is active during early-to-mid G1 phase and Cdk2 is active in late G1 phase; both were required for progression through the G1/S transition through phosphorylating a critical serine residue of Rb (Ser 795) (Koff et al., 1992; Rosenblatt et al., 1992; Swanton, 2004; van den Heuvel and Harlow 1993). The present data show that within the range of concentrations from 0.1 μM to 10 μM, atRA inhibited reaction activity of both cdk2 and cdk4 (Fig. 5).
DISCUSSION
The physiologic doses (usually 0.01 μM) of RA play a important role in embryogenesis. The unsuitable maternal use of certain retinoid-containing medications and excessive supplements at specific critical periods of facial morphogenesis could lead to aberrant development and subsequent cleft palate. The greatest success in the RA-induced cleft palate in mice was demonstrated by a single dose of exogenous atRA on GD 11 at a level of 100 mg/kg. Given the absence of maternal toxicity, it is evident that all pharmacological doses (1 μM) of RA are at the toxic thresholds. To gain insight into possible mechanisms of atRA-induced cleft palate, we mainly studied the effects of pharmacologic (1 μM) doses of atRA on growth of MEPM cells.
In atRA-treated MEPM cultures, we found a dose-dependent survival suppression that blocked cell cycle progression at the G1/S transition, ultimately leading to G1/G0 arrest and eliciting apoptosis. Data here are in agreement with a previous in vivo study, which showed that the inhibition of growth and excess apoptosis of MEPM cells contributed to the formation of cleft palate and other orofacial congenital malformations (Suwa et al., 2001). It is well known that atRA treatment induced decrease of cell viability and increase of apoptotic phenomena in diverse tumor cell types, including cancers of the breast, lung, liver, and blood (Guzey et al., 1998; Hsu et al., 2001; Jimenez-Lara et al., 2004; Mangiarotti et al., 1998). Some teratogenic effects of retinoids may be due to their ability to inhibit cell growth and induce apoptosis. For example, RA treatment of midgestation mouse embryos induces excess limb bud apoptosis, resulting in malformed limbs (Jiang and Kochhar, 1992). Cleft palate results from failure of fusion between the palatal shelves. One possible cause is retarded mesenchymal proliferation and differentiation in those shelves, which will lead to retardation in the development of the palatal bones (Furukawa et al., 2004; Stark, 1954).
Proliferating mammalian cells pass through several cell cycle checkpoints, mainly G1-to-S and G2-to-M transitions. Three important targets have emerged as likely candidates for effectors of this outcome including the expression of cyclin D1 and E and the phosphorylated status of Rb. Data here show that atRA induced a dose-dependent decrease in cyclins D and E. During early G1 phase, cyclins D bind to and activates cdk 4, cyclin D–cdk4 complexes phosphorylate their primary target protein, the tumor suppressor retinoblastoma (Rb) protein. In late G1, cyclins E interact with and activate cdk2. Cyclin E–Cdk2 complexes complete this phosphorylation (Duman-Scheel et al., 2002; Yamada et al., 2004). Hyperphosphorylated Rb no longer binds to its target, E2F, thereby allowing E2F to activate the transcription of genes required for S phase in several consecutive steps and thereby ultimately relieving Rb-mediated repression from S phase, as Rb becomes dephosphorylated just prior to M/G1 transition (Bai et al., 2003; DiPietrantonio et al., 1998; Johnson and Schneider-Broussard, 1998; Sherr, 2000). Given the importance of Rb, it is reasonable that inhibition of phosphorylation of Rb is thought to be one of the important mechanisms by which retinoids inhibit proliferation in MEPM cells.
The formation of secondary palate involves several ordered steps, starting with shelf growth and elevation, and followed by contact, adhesion, and disappearance of the medial edge epithelium. Disruption at any stage of palate development could potentially cause cleft palate. It has been known for some time that maternal treatment of atRA produces embryonic palatal shelves deficient in mesenchymal tissue (Horie and Yasuda, 2001; Ikemi et al., 2001) and exhibiting a decreased rate of 3H-thymidine uptake, indicative of inhibition of cell proliferation (Watanabe et al., 1990). Retinoic acid has also been shown to inhibit proliferation of, and extracellular matrix synthesis in mouse fetal palate mesenchymal cells. These data suggest that perturbation of the process of cell proliferation might account for the formation of small palatal shelves and the failure of palatal fusion in vivo (Watanabe et al., 1988).
Together, these results are consistent with the hypothesis that retinoid-induced cleft palate might depend in part on the effect of RA on growth inhibition and modification of cell cycle distribution of palate mesenchymal cells in the developing palatal shelves. We proposed that decreased cdk activity, followed by reduced phosphorylation of Rb, may account for the failure of palate fusion exposed to RA as the elements of the secondary palate are just beginning to make contract.
ACKNOWLEDGMENTS
This work was supported by National Natural Science Foundation grant nos. 30371224 and 30371559, People's Republic of China, and the Major Basic Research Development Program of the People's Republic of China (2001CB510305).
NOTES
REFERENCES
Bai, S., Goodrich, D., Thron, C. D., Tecarro, E., and Obeyesekere, M. (2003). Theoretical and experimental evidence for hysteresis in cell proliferation. Cell Cycle 2, 46–52.
Connell-Crowley, L., Harper, J. W., and Goodrich, D. W. (1997). Cyclin D1/Cdk4 regulates retinoblastoma protein-mediated cell cycle arrest by site specific phosphorylation. Mol. Biol. Cell. 8, 287–301.
Dhulipala, V. C., Hanumegowda, U. M., Balasubramanian, G., and Reddy, C. S. (2004). Relevance of the palatal protein kinase A pathway to the pathogenesis of cleft palate by secalonic acid D in mice. Toxicol. Appl. Pharmacol. 194, 270–279.
DiPietrantonio, A. M., Hsieh, T. C., Olson, S. C., and Wu, J. M. (1998). Regulation of G1/S transition and induction of apoptosis in HL-60 leukemia cells by fenretinide (4HPR). Int. J. Cancer 78, 53–61.
Donnellan, R., and Chetty, R. (1999). Cyclin E in human cancers. FASEB J. 13, 773–780.
Duman-Scheel, M., Weng, L., Xin, S., and Du, W. (2002). Hedgehog regulates cell growth and proliferation by inducing cyclin D and cyclin E. Nature 417, 299–304.
Eldeib, M. M. R., and Reddy, C. S. (1988). Secalonic acid D–induced changes in palatal cyclic AMP and cyclic GMP in developing mice. Teratology 37, 343–352.
Ferguson, M. W. (1988). Palate development. Development 103(Suppl), 41–60.
Furukawa, S., Usuda, K., Abe, M., and Ogawa, I. (2004). Histopathological findings of cleft palate in rat embryos induced by triamcinolone acetonide. J. Vet. Med. Sci. 66, 7–402.
Geng, Y., Eaton, E. N., Picon, M., Roberts, J. M., Lundberg, A. S., Gifford, A., Sardet, C., and Weinberg, R. A. (1996). Regulation of cyclin E transcription by E2Fs and retinoblastoma protein. Oncogene 12, 1173–1180.
Guzey, M., Demirpence, E., Criss, W., and DeLuca, H. F. (1998). Effects of retinoic acid (all-trans and 9-cis) on tumor progression in small cell lung carcinoma. Biochem. Biophys. Res. Commun. 242, 369–375.
Harbour, J. W., and Dean D. C. (2000). The Rb/E2F pathway: expanding roles and emerging paradigms. Genes Dev. 14, 2393–409.
Horie, S., and Yasuda, M. (2001). Alterations in palatal ruga patterns in Jcl:ICR mouse fetuses from dams treated with all-trans-retinoic acid. Hiroshima J. Med. Sci. 50, 17–25.
Hsu, S. L., Cheng, C. C., Shi, Y. R., and Chiang, C. W. (2001). Proteolysis of integrin alpha5 and beta1 subunits involved in retinoic acid-induced apoptosis in human hepatoma Hep3B cells. Cancer Lett. 167, 193–204.
Ikemi, N., Otani, Y., Ikegami, T., and Yasuda, M. (2001). Palatal ruga anomaly induced by all-trans-retinoic acid in the Crj:SD rat: Possible warning sign of teratogenicity. Reprod. Toxicol. 15, 87–93.
Jiang, H., and Kochhar, D. M. (1992). Induction of tissue transglutaminase and apoptosis by retinoic acid in the limb bud. Teratolog. 46, 333–340.
Jimenez-Lara, A. M., Clarke, N., Altucci, L., and Gronemeyer, H. (2004). Retinoic-acid-induced apoptosis in leukemia cells. Trends Mol. Med. 10, 508–515
Johnson, D. G., and Schneider-Broussard, R. (1998). Role of E2F in cell cycle control and cancer. Frontiers Biosci. 27, 447–8.
Kitagawa, M., Higashi, H., Jung, H. K, Suzuki-Takahashi, I., Ikeda, M., Tamai, K., Kato, J., Segawa, K., Yoshida, E., Nishimura, S., and Taya, Y. (1996). The consensus motif for phosphorylation by cyclin D1-Cdk4 is different from that for phosphorylation by cyclin A/E-Cdk2. EMBO J. 15, 7060–7069.
Koff, A. A., Giordano, D., Desai, K., Yamashita, J. W., Harper, S., Elledge, T., Nishimoto, D. O., Morgan, B. R., Franza, B. R., and Roberts, J. M. (1992). Formation and activation of a cyclin E-cdk2 complex during G1 phase of the human cell cycle. Science 257, 1689–1694.
Lai, L., Bohnsack, B. L., Niederreither, K., and Hirschi, K. K. (2003). Retinoic acid regulates endothelial cell proliferation during vasculogenesis. Development 130, 6465–6474.
Lundberg, A. S., and Weinberg, R. A. (1998). Functional inactivation of the retinoblastoma protein requires sequential medication by at least two distinct cyclin-cdk complexes. Mol. Cell Biol. 18, 753–761.
Mangiarotti, R., Danova, M., Alberici, R., and Pellicciari, C. (1998). All-trans retinoic acid (ATRA)-induced apoptosis is preceded by G1 arrest in human MCF-7 breast cancer cells. Br. J. Cancer 77, 186–191.
Ohtani, K. (1999). Implication of transcription factor E2F in regulation of DNA replication. Frontiers Biosci. 4, D793–804.
Ohtsubo, M., Theodoras, A. M., Schumacher, J., Roberts, J. M., and Pagano, M. (1995). Human cyclin E, a nuclear protein essential for the G1-to-S phase transition. Mol. Cell Biol. 15, 2612–2624.
Pan, W., Cox, S., Hoess, R. H., and Grafstrom, R. H. (2001). A cyclin D1/cyclin dependent kinase 4 binding site within the C domain of the retinoblastoma protein. Cancer Res. 61, 2885–2891.
Rosenblatt, J., Gu, Y., and Morgan, D. O. (1992). Human cyclin dependent kinase 2 is activated during the S and G2 phases of the cell cycle and associates with cyclin A. Proc. Natl. Acad. Sci. U.S.A. 89, 2824–2828.
Sasaki, Y., Iwai, N., Tsuda, T., and Kimura, O. (2004). Sonic hedgehog and Bone morphogenetic protein 4 expressions in the hindgut region of murine embryo with anorectal malformations. J. Pediatr. Surg. 39, 170–173.
Shibamoto, S., Winer, J., Williams, M., and Polakis, P. (2004). A blockade in Wnt signaling is activated following the differentiation of F9 teratocarcinoma cells. Exp. Cell Res. 292, 11–20.
Sherr, C. J. (1996). Cancer cell cycles. Science 274, 1672–1677
Sherr, C. J. (2000). The Pezcoller Lecture: Cancer cell cycles revisited. Cancer Res. 60, 3689–3695.
Stark, R. B. (1954). The pathogenesis of harelip and cleft palate. Plast. Reconstruct. Surg. 13, 20–32.
Suwa, F., Jin, Y., Lu, H., Li, X., Tipoe, G. L., Lau, T. Y., Tamada, Y., Kuroki, K., and Fang, Y. R. (2001). Alteration of apoptosis in cleft palate formation and ectomesenchymal stem cells influenced by retinoic acid. Okajimas Folia. Anat. Jpn. 78, 179–186.
Swanton, C. (2004). Cell-cycle targeted therapies. Lancet Oncol. 5, 27–36.
van den Heuvel, S., E., and Harlow, E. (1993). Distinct roles for cyclin-dependent kinases in cell cycle control. Science 262, 2050–2054.
Wang, W., and Kirsch, T. (2002). Retinoic acid stimulates annexin-mediated growth plate chondrocyte mineralization. J. Cell Biol. 157, 1061–1069.
Watanabe, T., Goulding, E. H., and Pratt, R. M. (1988). Alterations in craniofacial growth induced by isotretinoin (13-cis-retinoic acid) in mouse whole embryo and primary mesenchymal cell culture. J. Craniofac. Genet. Dev. Biol. 8, 21–33.
Watanabe, T., Willis, W. D., and Pratt, R. M. (1990). Effect of retinoids on proliferation of human embryonic palatal mesenchymal cells in culture. J. Nutr. Sci. Vitaminol. (Tokyo) 36, 311–325.
Weinberg, R. A. (1995). The retinoblastoma protein and cell cycle control. Cell 81, 323–330.
Yamada, S., Sumrejkanchanakij, P., Amagasa, T., and Ikeda, M. A. (2004). Loss of cyclin E requirement in cell growth of an oral squamous cell carcinoma cell line implies deregulation of its downstream pathway. Int. J. Cancer 111, 17–22.
Zarkowska, T., and Mittnacht, S. (1997). Differential phosphorylation of the retinoblastoma protein by G1/S cyclin-dependent kinases. J. Biol. Chem. 272, 12738–12746.(Zengli Yu, Jiuxiang Lin, )