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Previous Article | Next Article 
Molecular and Cellular Biology, June 2006, p. 4240-4256, Vol. 26, No. 11
0270-7306/06/$08.00+0 doi:10.1128/MCB.02124-05
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, Pennsylvania 19107,1 Department of Oncology, Lombardi Comprehensive Cancer Center, Georgetown University, 3970 Reservoir Road, NW, Box 571468, Washington, D.C. 20057-1468,2 Department of Developmental and Molecular Biology, Albert Einstein College of Medicine, Yeshiva University, Bronx, New York 104613
Received 2 November 2005/ Returned for modification 13 December 2005/ Accepted 13 March 2006
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ABSTRACT |
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INTRODUCTION |
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Mutational analysis of the human cyclin D1 cDNA has identified several distinct domains involved in binding either pRb, cdk, the p160 coactivator, and histone deacetylases (22, 23, 59). The cdk-binding domain of cyclin D1 is required for the association with cdk4 and sequential phosphorylation of pRb, which in turn, leads to the release of E2F binding proteins. The release of E2F proteins, in turn, leads to the sequential regulation of E2F-responsive genes associated with the induction of DNA synthesis. The association of cyclin D1 with the p160 coactivator SRC1 (AIB1) enhances ligand-independent ER
activity in cultured cells. Recent studies have demonstrated the regulation of several transcription factors through a cdk-independent mechanism, including MyoD, Neuro-D, the androgen receptor, CEBPß, and peroxisome proliferator-activated receptor gamma (PPAR
) (reviewed in reference 73). The abundance of cyclin D1 is rate limiting in progression through the G1 phase of the cell cycle in fibroblasts and mammary epithelial cells. Sustained extracellular signal-regulated kinase (ERK) activation induces cyclin D1 transcription and mRNA and protein abundance, which is required for mid-G1-phase induction of cyclin D1 (2, 56, 75). Tightly coordinated interactions between the Rho GTPases facilitate cell cycle progression through regulating the expression of cyclin D1 and assembly of cyclin D/cdk complexes (12). Rac and Cdc42 induce cyclin D1 independently of ERK involving an NF-
B signaling pathway (12, 31, 79). Rho kinase suppresses Rac/Cdc42-dependent cyclin D1 induction through LIMK (56) independently of cofilin or actin polymerization. The inhibition of Rac/Cdc42 signaling maintains mid-G1-phase ERK-dependent induction of cyclin D1 (56).
The Rho family of small GTPases play an important role in the regulation of cell motility via their effects on the cellular cytoskeleton and adhesion (5, 32). Rac and its effector, PAK, induce membrane ruffles and actin rearrangements including stress fibers that control formation of lamellipodia and new focal contacts at the leading edge that drive cellular motility (54). Rho regulates assembly of stress fibers and associated focal adhesions through its downstream effectors mouse Diaphanous (mDia) and the Rho-activated kinase (ROCK) that phosphorylate cytoskeletal proteins. Major ROCK substrates regulating cellular migration include LIM kinases, which phosphorylate and regulate an actin-depolymerizing protein cofilin, and myosin light chain (MLC) kinase. Although Rho activity negatively influences cell migration by increasing stress fiber-dependent adhesions to substratum, Rho activity is also required for actomyosin contractility needed to drive cell body retraction at the rear of the cell (4). Dynamic activation and inactivation is tightly coordinated, and insufficient levels or excessive Rho GTPase activity will prevent cell migration (52, 57, 58, 71).
A variety of cytokines, chemokines, growth factors, extracellular matrix, and matrix-degrading proteins coordinate their signaling to affect migratory cues through the Rho family GTPases, and these factors are in turn regulated by Rho GTPases. Thrombospondin 1 (TSP-1), for example, is a matrix glyocoprotein that inhibits cellular metastasis and is repressed by oncogenic Ras (64). It is the first protein to be recognized as a naturally occurring inhibitor of angiogenesis (26). TSP-1 overexpression inhibits wound healing and tumorigenesis (55, 63, 64, 65). Conversely, lack of functional TSP-1 increases tissue vascularization. The abundance of TSP-1 is tightly regulated, and it is the alteration from the physiological level that seems to specifically affect migration. Thus, inhibition of TSP-1 from TSP-1-oversecreting cells reverts abnormal migration, but immunoneutralizing antibodies to TSP-1 do not affect migration of normal cells (72).
In the present study, cyclin D1–/– mouse embryo fibroblasts (MEFs) exhibit increased adherence and cellular spreading and decreased cellular motility compared with wild-type cells. Genomewide expression analysis of cyclin D1–/– MEFs transduced with a cyclin D1 expression vector that rescued the defect in adhesion and migration identified a subset of genes governing migration that were regulated by cyclin D1, including TSP-1 and ROCKII. cyclin E knockin to the cyclin D1–/– MEFs rescued the DNA synthesis defect of cyclin D1–/>– MEFs but did not rescue either the migration defect or the abundance of ROCKII. Increased ROCKII activity in cyclin D1–/– cells was corroborated by increased phosphorylation of LIM kinase, cofilin, and MLC. TSP-1 immunoneutralization or ROCK kinase inhibition rescued the migration defect of cyclin D1–/– MEFs. Thus, cyclin D1 inhibits the ROCKII and TSP-1 signaling pathway to promote cellular migration.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cell culture. Cultures of cyclin D1+/+, cyclin D1–/–, and cyclin E knockin to the cyclin D1–/– primary MEFs were prepared as described previously (74). Human kidney 293 and 293T cells were maintained in Dulbecco's modified Eagle's medium (DMEM) containing penicillin and streptomycin (100 mg of each/liter) and supplemented with 10% fetal bovine serum (FBS).
Western blotting. Whole-cell lysates (50 µg) were separated by 10% sodium dodecyl sulfate (SDS-PAGE), and the proteins were transferred to nitrocellulose membrane for Western blotting as previously described (11). The following antibodies were used for Western blotting: rabbit Ab-3 anti-cyclin D1 antibody and mouse Ab-11 antithrombospondin (TSP) from Lab Vision/NeoMarker, Fremont, CA; an antibody against guanine nucleotide dissociation inhibitor (GDI) (35); mouse M2 anti-FLAG antibody, and antivinculin antibody from Sigma, St. Louis, MO; rabbit polyclonal antibody for Thr18/Ser19 phospho-myosin light chain 2, rabbit anti-LIMK1, and rabbit anti-phospho-LIMK (Thr508/Thr505) from Cell Signaling Technology, Beverly, MA; rabbit polyclonal antibody for Ser3 phospho-ADF/cofilin, mouse DCS-6 anti-cyclin D1 antibody, rabbit polyclonal antipaxillin antibody, anti-ROCKII, mouse anti-human cyclin E antibody (HE12), and anti-ß-tubulin antibody from Santa Cruz Biotechnology, Santa Cruz, CA; and rabbit polyclonal anti-Y118 phospho-specific paxillin antibody from Biosource International, Camarillo, CA.
Retroviral production and infection.
The mouse stem cell virus (MSCV)-internal ribosome entry site (IRES)-green fluorescent protein (GFP) retrovirus vector and the ecotropic, packaging vector, pSV-
–E-MLV, which provides ecotropic packaging helper function, and infection methods were as described previously (45). Briefly, the coding region of mouse cyclin D1 cDNA, 3x FLAG-tagged wild-type human cyclin D1, and a series of human cyclin D1 mutant cDNAs (74) were inserted into the MSCV-IRES-GFP vector at the EcoRI site upstream of the IRES driving expression of GFP. MSCV retroviruses were prepared by transient cotransfection with helper virus into 293T cells, using calcium phosphate precipitation. The retroviral supernatants were harvested 48 h after transfection and filtered through a 0.45-µm filter. cyclin D1–/– MEFs were incubated with fresh retroviral supernatants in the presence of 8 µg/ml Polybrene for 24 h, cultured for a further 4 days, and subjected to fluorescence-activated cell sorting (FACS) (FACStar Plus; BD Biosciences, San Jose, CA) to select for cells expressing GFP. GFP+ cells were used for subsequent analysis.
Fluorescent and phase-contrast light microscopy. GFP+ cells were examined in six-well plates. Fluorescent microscopy and phase-contrast imaging were carried out using the x10 objective of an Olympus IX-70 laser-scanning confocal microscope.
Cell cycle analysis, phalloidin staining, and F-actin quantitation. DNA synthesis of MEFs was determined by propidium iodide (PI) staining and FACS analysis as described previously (1). Phalloidin staining was conducted as previously described (45). F-actin quantitation was carried out by FACS analysis (50).
Immunofluorescence. cyclin D1+/+, cyclin D1–/–, and cyclin D1–/– MEFs infected with MSCV-cyclin D1-IRES-GFP or GFP vector control grown in four-well chamber slides were fixed with cold methanol for 20 min at 4°C. The slides were air dried at room temperature. The primary antibodies used were mouse monoclonal antipaxillin (clone 5H11; Upstate) (1/100) and rabbit polyclonal anti-phospho-paxillin (pY118) (Biosource) (1/100). The secondary antibodies used were Alexa Fluor 488-conjugated F(ab')2 fragment of goat anti-mouse immunoglobulin G (IgG) (Molecular Probes, Inc.) (1/250) and rhodamine red X-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch Lab.) (1/50). The samples were visualized on an Olympus IX70 laser-scanning confocal microscope with a x60 PlanApo oil immersion objective and using Olympus Fluoview FV-300 software. The images were processed with MetaMorph (Molecular Devices).
SEM. For scanning electron microscopy (SEM), cells were plated on fibronectin-coated glass coverslips and grown to approximately 80% confluence. The cells were fixed as described previously (45) to prevent agonal membrane artifacts. Dehydrated cells were critical point dried using liquid carbon dioxide in a Samdri 790 critical point drier (Tousimis Research, Rockville, MD), sputter-coated with gold-palladium in a Vacuum Desk-1 sputter coater (Denton, Cherry Hill, NJ), mounted, and viewed in a AMRAY 1820D scanning electron microscope by using an accelerated voltage of 20 kV for electron microscopy.
IRM. Interference reflection microscopy (IRM) was performed as described as previously (45). Briefly, IRM was performed using a x60 objective of an Olympus IX-70 laser-scanning confocal microscope in reflectance mode with polarization filters at either 488 or 568 nm. Direct adherence or apposition of the cell to the substrate is imaged as black (destructive interference), and greater distance is viewed as white (direct reflection or constructive interference). Image analysis was performed on the images based on the assumption that more adherent cells would have more dark pixels per unit area of spread than less-adherent cells. NIH ImageJ software was used to manually trace the boundaries of individual cells, and a grayscale histogram was derived for each cell. The threshold was set based on the shape of the histogram (15). Values below the threshold corresponded to regions that appeared black. A ratio was derived for each cell (percentage close apposition) that consisted of the sum of pixels below the threshold divided by the total number of pixels within the boundaries of the cells.
Adhesion assay. Adhesion assays were conducted as previously described (45). Briefly, cells (2 x 104) were seeded in serum-free medium in a 96-well plate coated with 10 µg/ml fibronectin (Sigma, St. Louis, MO). After 30 min, plates were washed twice with phosphate-buffered saline (PBS), fixed, and stained with crystal violet. Adherent cells were represented as A550.
Spreading assay. Spreading assays were conducted as described previously (47). Briefly, cells were plated on 60-mm plastic tissue culture dishes in DMEM containing 10% FBS for the indicated time points. Dark cells were considered to be spread, and bright cells were considered to be unspread. Pictures of three independent fields were taken under the x10 objective. Experiments were done in triplicate and repeated at least three times.
Migration assay. Transwell migration assays were performed as described before (38). Briefly, GFP-positive cells were seeded on 8-µm-pore-size Transwell filter insert (Costar) coated with 10 µg/ml fibronectin (Sigma, St. Louis, MO). After 16 h of incubation at 37°C and 5% CO2, cells adherent to the upper surface of the filter were removed using a cotton applicator. Cells were fixed with 3.7% formaldehyde and stained with crystal violet, and the numbers of cells on the bottom were counted. Data are from three experiments done in triplicate (mean ± standard error).
Wound healing. Cells were grown to confluence on 12-well plates, and the monolayers were wounded with a P10 micropipette tip (45). DMEM with 10% FBS and HEPES was changed immediately after scoring. The wound-healing videos were taken at 20-min intervals (30 min in p16INK4a peptide experiments) using a Nikon Eclipse TE-300 inverted microscope system. The cell movement velocity was determined by tracing single cells at different time points using MetaMorph software (Molecular Devices Corporation, Downington, PA).
p16 peptides.
Peptides corresponding to amino acids 84 to 103 of human p16INK4a protein with a C-terminal sequence of 16 amino acids encoding the Antennapedia homeodomain (Penetratin) (20, 21, 27) were synthesized (Bio-synthesis, Inc. Lewisville, TX). Peptide 20 (DAAREGFLATLVVLHRAGARRQIKIWFQNRRMKWKK) with the D92A substitution has a lower 50% inhibitory concentration (IC50) to inhibit cdk4-cyclin D1 phosphorylation of a glutathione S-transferase (GST)-Rb protein in vitro and to arrest cell cycle progression in G1 than the corresponding peptide containing the wild-type sequence (20, 21). Peptide 21 (DAAREGFLDTLAALHRAGARRQIKIWFQNRRMKWKK) carrying VV95 and 96AA mutations, has an increased IC50 in vitro and has lost 
60% of the cell cycle inhibitory capacity (20, 21). These peptides were added to the cell culture medium at a concentration of 20 µM.
Time-lapse video. For time-lapse observation of cell movement, cells on 12-well plates were maintained in DMEM with 10% fetal calf serum (FCS) and HEPES. Cells were placed in a temperature and CO2 control incubator to maintain the temperature at 37°C and CO2 at 5%. The cell movement videos were taken at 5-min intervals by using a Nikon Eclipse TE-300 inverted microscope system. The cell movement velocity was determined by tracing the single cells at different time points using MetaMorph software. To observe the effect of ROCK inhibition, cells were treated with 10 µM Y27632 (Calbiochem) for 30 min before time-lapse recording. For assays with neutralizing antibody, the A4.1 anti-TSP-1 monoclonal antibody (Neomarkers) was added at 10 µg/ml as previously described (72).
Microarray analysis. Total RNA was isolated from retrovirus vector-infected cyclin D1–/– MEFs (infected with either MSCV-cyclin D1-IRES-GFP or MSCV-IRES-GFP control vector) using Trizol and used to probe Affymetrix MU74v2 arrays (Affymetrix, Santa Clara, CA). RNA quality was determined by gel electrophoresis. Probe synthesis and hybridization were performed according to the manufacturer's manual (see the eukaryotic target preparation section at http://www.affymetrix.com/support). Three arrays were used for each condition. Analysis of the arrays was performed using the statistical package R statistics package (14) and the limma library (62) of the Bioconductor software package. Arrays were normalized using robust multiarray analysis (RMA), and the genes were ranked using the log odds ratios for differential expression (9, 53). Briefly, a linear model was constructed using a factorial design and differentially expressed genes were obtained from cyclin D1–/– plus cyclin D1 or a cyclin D1–/– vector control. Finally, the top-ranked genes that are differentially expressed in a cyclin D1-dependent manner were determined based on their log odds ratio. These genes were then clustered using hierarchical clustering with "complete" agglomeration, and each cluster was further analyzed based upon the known function of the genes contained in the cluster. All six arrays were analyzed and show good separation based upon their cyclin D1 expression levels (8, 62).
Luciferase assays. A luciferase reporter plasmid containing bp –2800 to +48 of the murine thrombospondin 1 (mTSP-1) promoter was kindly provided by Paul Bornstein (University of Washington, Seattle) (7). HEK 293 cells were used in mTSP-1 reporter assays. The cells were transiently transfected with increasing amounts of wild-type or KE mutant cyclin D1 (in a cytomegalovirus 10 [CMV10] vector) in combination with the reporter using Superfect reagent (QIAGEN, Valencia, CA), according to the manufacturer's instructions. Forty-eight hours posttransfection, luciferase assays were performed at room temperature using an Autolumat LB 953 (EG&G Berthold) as previously described (74). A CMV-ß-galactosidase plasmid was cotransfected as a control for transfection efficiency. Luciferase activities were measured and normalized by the empty vector CMV10 control.
Semiquantitative RT-PCR. Total cellular RNA was prepared from cyclin D1+/+ and cyclin D1–/– P-3 MEFs. Semiquantitative reverse transcription-PCR (RT-PCR) was performed as previously described (72) using the following primers: forward primer 5' GGGCTAGAGAAACCCCCCAC 3' and reverse primer 5' CCAAAGGGAGAAAGTCC 3' to amplify murine TSP-1 and forward primer 5' TGTTACCAACTGGGACGACA 3' and reverse primer 5' AAGGAAGGCTGGAAAAGAGC 3' to amplify murine ß-actin.
Rho pulldown assay. The amount of activated Rho present in cyclin D1–/– MEFs infected with wild-type cyclin D1 and vector was determined by using the Rho activation assay kit (Upstate Cell Signaling Solutions, Lake Placid, NY), according to the manufacturer's instruction. The amounts of bound (activated) and total Rho (RhoA, -B, and -C) were measured by immunoblotting.
ROCKII kinase assay.
For ROCKII kinase activity assays, cell lysates (500 µg protein in NP40 lysis buffer: 1% NP40, 150 mM NaCl, 50 mM Tris, pH 7.4) were incubated with 10 µl of polyclonal anti-ROCKII (H-85; Santa Cruz Biotechnology) and 50 µl protein A agarose (Roche) overnight at 4°C. The immunocomplexes were washed and subjected to kinase reaction in 20 µl of kinase buffer (50 mM Tris, pH 7.4, 10 mM MgCl2, 3 mM NaCl, 1 mM dithiothreitol, 1 mM EDTA) in the presence of 10 µCi of [-32P]ATP and 20 µg of myelin basic protein (MBP) (Sigma). Reaction mixtures were incubated for 20 min at 37°C, and reactions were terminated by the addition of 4 µl of 6x SDS sample buffer. Samples were resolved by 15% SDS-polyacrylamide gel electrophoresis, and the gel was dried and subjected to autoradiography.
Online supplemental material. An online supplement includes the wound-healing videos (http://www.kimmelcancercenter.org/pestell/papers/zl/video/) of cyclin D1+/+ (see Fig. 5Avideo1 at the URL mentioned above), cyclin D1–/– MEFs (Fig. 5Avideo2), cyclin D1–/– MEFs infected with wild type (Fig. 5Bvideo1) or mutant GH (Fig. 5Bvideo2), LLAA (Fig. 5Bvideo3), KE (Fig. 5Bvideo5) cyclin D1 or vector control (Fig. 5Bvideo4), and cyclin D1+/+ MEFs treated with p16INK4a peptide 20 (Fig. 5Dvideo1) or p16INK4a peptide 21 (Fig. 5Dvideo3) or without peptide treatment control (Fig. 5Dvideo2). Microarray data of cyclin D1–/– plus cyclin D1 and vector control were also included in the online supplemental material (http://www.kimmelcancercenter.org/pestell/papers/zl/D1vsGFP.xls). Supplements 1 and 2 may also be found online (http://www.kimmelcancercenter.org/pestell/papers/zl/Supplement1.pdf and http://www.kimmelcancercenter.org/pestell/papers/zl/Supplement2.pdf).
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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0.02). Mutation of the cdk4-binding site in cyclin D1 abrogated this rescue of DNA synthesis. All cyclin D1–/– MEFs transduced with the MSCV-cyclin D1-IRES-GFP vector reverted to the fibroblastoid morphology, yet only 50% of the cells were in the DNA synthetic phase.
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0.008). In contrast, expression of the cdk-binding defective (KE) mutant, failed to reduce the closely apposed area detected by IRM (P = 0.12 and P = 0.14, respectively) (see supplement 1B at http://www.kimmelcancercenter.org/pestell/papers/zl/Supplement1.pdf).
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30% at 30 min after plating (P = 0.0007 and P = 0.0004, respectively) (see supplement 2 at http://www.kimmelcancercenter.org/pestell/papers/zl/Supplement2.pdf). Both the pRb-binding-defective mutant (GH mutant) and the SRC1-binding-defective mutant (LLAA mutant) decreased the percentage of spread cells at 30 min (P = 0.0032) (see supplement 2 at the URL above). However, the cdk-binding-defective mutant (KE mutant) failed to reduce the percentage of spread cells (see supplement 2 at the URL above). Cyclin D1-deficient MEFs were more adherent than wild-type cells on fibronectin (P = 0.03) (Fig. 4D). Exogenous mouse and human cyclin D1 wild-type, GH mutant, or LLAA cyclin D1 mutants decreased adherence on fibronectin (P = 8.7 x 10–10, P = 1 x 10–6, P = 9.1 x 10–6, P = 4.4 x 10–6) (see supplement 2 at the URL above), while the cdk-binding-defective mutant (KE mutant) failed to reduce adhesion (see supplement 2 at the URL above). p16INK4a inhibits and cyclin D1 enhances the velocity of cellular migration. cyclin D1–/– MEFs migrated significantly less than wild-type cells on fibronectin-coated Transwell plates (P = 0.0008) (Fig. 4E). Retroviral introduction of the murine or human cyclin D1 cDNA into cyclin D1–/– MEFs increased the cell migration rate significantly (P = 0.0006 and P = 0.001, respectively) (see supplement 2C at the URL above). The pRb-binding-defective cyclin D1 mutant (GH mutant) and the SRC1-binding-defective mutant (LLAA mutant) both increased cell migration (P = 0.016 and P = 0.0005, respectively) (see supplement 2C at the URL above), which was not enhanced by the cdk-binding-defective mutant (KE mutant).
By overnight time-lapse videomicroscopy, the single-cell motility of cyclin D1–/– MEFs was significantly slower than the motility of wild-type cells (P = 0.0006) (Fig. 5A; see Fig. 5Avideo1 and -2 at http://www.kimmelcancercenter.org/pestell/papers/zl/video/). Reintroduction of the cyclin D1 wild type or the cyclin D1 mutants GH or LLAA increased the velocity of cell movement compared with that in vector control cells (P = 0.0048, P = 0.0009, and P = 0.0002, respectively) (Fig. 5B; see Fig. 5Bvideo1 to -4 at http://www.kimmelcancercenter.org/pestell/papers/zl/video/), whereas there was no significant effect of the cdk-binding-defective mutant (KE mutant) (P = 0.125) (Fig. 5B; see Fig. 5Bvideo5 at http://www.kimmelcancercenter.org/pestell/papers/zl/video/). Thus, cyclin D1 enhances the velocity of cell motility.
A synthetic peptide that spans the two -helixes of the third ankyrin repeat of p16INK4a, interacts with the cdk, inhibiting cyclin D1-kinase activity. The derivative of this peptide carrying a D92A substitution (peptide 20) (Fig. 5D) has a lower IC50 compared with the wild-type sequence (20). The p16INK4a peptide 20 linked to the Antennapedia carrier sequence inhibited wound closure compared with the no-peptide-treatment control (P = 0.0016) (Fig. 5D; see Fig. 5Dvideo1 and -2 at http://www.kimmelcancercenter.org/pestell/papers/zl/video/). The p16INK4a peptide 21, which carries alanine substitutions of two valine residues corresponding to positions 95 and 96 of the p16INK4a that dramatically increases its IC50 in vitro (20), had reduced ability to inhibit MEF migration (P = 0.98) (Fig. 5E; see Fig. 5Dvideo3 at http://www.kimmelcancercenter.org/pestell/papers/zl/video/). This finding is consistent with a model in which cdk binding regulates MEF cellular migration by cyclin D1.
Cyclin D1 regulates a molecular genetic cluster governing cellular adhesion and migration. We conducted a comprehensive genomewide interrogation to identify gene targets regulated by cyclin D1 that may play a role in cell adhesion and migration. cyclin D1–/– MEFs were infected with either wild-type cyclin D1 or the empty GFP vector as control. Analyses were conducted in triplicate, and the subsets of genes differentially regulated by cyclin D1 were identified (Fig. 6 and Table 1; see also http://www.kimmelcancercenter.org/pestell/papers/zl/D1vsGFP.xls). Differential expression of genes previously identified as regulating cellular migration and adhesion included those coding for TSP-1 and ROCKII. Given the central role of ROCKII and TSP-1 in regulating cellular migration, we considered the possibility that cyclin D1 may regulate the abundance and activity of these two key proteins.
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6-fold (Fig. 7B and C). To determine whether the cyclin E could rescue the migration defect of cyclin D1 deficiency, we examined MEFs derived from mice in which the cyclin D1 gene coding sequence was replaced with the human cyclin E gene cDNA (cyclin E knockin to the cyclin D1–/– mice) (24). The cell movement velocity of the cyclin E knockin MEFs was reduced to approximately 20% of that of wild-type littermate controls (Fig. 7D and E). Cyclin E protein was well expressed in the cyclin E knockin MEFs (Fig. 7F); however, as with the cyclin D1–/– MEFs, ROCKII abundance remained increased. The cyclin E knockin was capable of rescuing the defect in DNA synthesis of the cyclin D1–/– cells (Fig. 7G), as previously described (24). The reduction in ROCK II kinase activity in the cyclin D1+/+ cells compared with cyclin D1–/– cells suggested cyclin D1 inhibited Rho activity. To determine whether cyclin D1 was sufficient to inhibit Rho GTPase activity, cyclin D1–/– cells were transduced with a cyclin D1-expressing retrovirus. Consistent with the relative reduction in ROCKII kinase in the cyclin D1-expressing cells, Rho-GTP activity and total Rho abundance were decreased upon cyclin D1 transduction of cyclin D1–/– cells compared to vector control-transduced cells (Fig. 7H).
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3-fold compared with normal mouse IgM control (Fig. 9E and F). Addition of TSP-1 antibody to normal wild-type MEFs did not affect cellular migration, consistent with previous studies (72). The concurrent addition of the TSP-1 antibody to the Y27632-treated cyclin D1–/– cells provided no significant additional increase in cellular velocity compared with Y27632 alone (data not shown).
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DISCUSSION |
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The current studies are consistent with recent findings linking factors that regulate cellular spreading and cell cycle progression. AP-1 proteins are known to promote both DNA synthesis and cellular motility (68, 71). Under several circumstances, cellular adhesion and migration promote G1-phase progression. Increased Rho activity, as observed in the cyclin D1–/– cells in the current studies, has been shown to natively regulate motility by increasing stress fiber-dependent adhesion (16). Although cellular adhesion is frequently required for cell cycle progression to occur, high levels of cell adhesion with large focal contacts results in failed cellular migration. Rho may also promote migration, being required for actomyosin contraction (4). Mutational analysis of the Rho/Rac chimera demonstrates distinct residues within the Rac effector domain regulate cellular morphology versus cyclin D1 expression and DNA synthesis (31, 80). Further evidence for a dissociation between the regulation of cellular morphology through Rho activity and DNA synthesis through pRb includes findings that unlike Rac and Cdc42, activated RhoA, despite its ability to induce morphological changes, does not inactivate pRb in NIH 3T3 cells (25). Together, these studies suggest the regulation of cell cycle progression and adhesion may be coupled or uncoupled, depending upon the cell type and signaling pathway involved.
Cyclin D1 promoted cell migration by inhibiting ROCKII expression and activity. ROCK kinase inhibitor reversed the defect in cellular motility in cyclin D1–/– cells, generating the more polarized morphology of wild-type cells. These findings are consistent with studies that ROCKII small interfering RNA induces an elongated morphology and that ROCK inhibitor treatment of cells induces a more polarized morphology (37) and increased cellular migration (6). Cyclin D1-deficient cells demonstrated evidence of increased ROCK activity using the in vitro substrate of myelin basic protein (MBP) and with increased phosphorylation of MLC2 (Thr18/Ser19), LIMK1/2 (Thr508/505), and cofilin (Ser3). The relationship between the activity of ROCK and cellular migration is complex and cell type specific. Thus, the ROCK inhibitor Y-27632 may either promote or inhibit cell migration (29, 46, 69, 81). The present findings are consistent with several recent findings. B-raf–/– MEFs demonstrated reduced ROCKII, reduced phosphocofilin, and increased migration (50), further confirming the correlation of increased ROCKII and reduced migration as seen in our studies. The possibility that the spread phenotype in cyclin D1–/– MEFs was a function of early senescence was excluded by the finding that pH-sensitive ß-galactosidase staining (18) was similar between the cyclin D1–/– and cyclin D1+/+ MEFs used in the studies (data not shown).
Several lines of evidence suggest the reduced migration of cyclin D1 deficiency is not a direct consequence of reduced DNA synthesis. First, all cyclin D1–/– cells exhibited the migratory defect, whereas only 7% of the cells were in the DNA synthetic phase. Second, all cyclin D1–/– cells transduced by cyclin D1 demonstrated the change in cellular morphology, alterations in focal contact distribution and cellular migration, yet only a fraction of the cells were undergoing DNA synthesis. Third, the reintroduction of cyclin E through a knockin of the human cyclin E cDNA rescued the DNA synthetic defect of cyclin D1–/– cells but did not rescue either the migration defect nor the ROCKII abnormality. Collectively these studies suggest the effect of cyclin D1 on cell cycle progression and cellular migration may be dissociable functions. The effect of Rho, and its signaling components, including mDia, on cellular morphology can be uncoupled from effects on DNA synthesis. ROCK inhibition stimulates passage through G1 phase in the absence of cell spreading (37), and ROCK activity is not required for cell-shape-dependent G1 progression in microvascular endothelial cells. mDia, which promotes actin polymerization and regulates the alignment of stress fibers by targeting microtubules to focal adhesions, fails to promote G1-phase progression (37). Thus, interaction between ROCK and its regulation of cellular adhesion and DNA synthesis is dissociable and cell type dependent.
Cyclin D1 deficiency increased adhesion in both macrophages and MEFs, changes correlating with the induction of circumferential cortical F-actin stress fibers in MEFs and in macrophages (45). As with cyclin D1, genetic deletion of several other proteins known to regulate migration including RhoB, cyclin B, and p27kip1, do not appear to affect embryonic development (6, 10, 38, 40, 44, 49). The stable adhesive structures in cyclin D1-deficient cells are thought to contribute to the increased adhesion and reduced migration of these cells. It has been proposed that an increase in either Rac (40) or Rho (6) activity and/or their effectors can contribute to reduced cellular migration. Cyclin D1 transcription is induced by the Rho GTPases (12, 31, 79). Cyclin D1 induction in response to growth factors as a delayed early response at 6 h requires Rho-dependent sustained ERK activation and results in the induction of DNA synthesis. Rho also inhibits an alternative Rac/Cdc42-dependent induction of cyclin D1, thus preventing its premature induction (77). The inhibition of Rac-dependent expression of cyclin D1 involves LIM kinase through an effect that is independent of cofilin phosphorylation and actin polymerization. Herein, cyclin D1 also functions as an upstream inhibitor of Rho/ROCK/LIMK. As the effect of nuclear LIM kinase on cyclin D1 abundance regulates the duration of G1 phase (56), together these studies suggest cyclin D1 may function as a fine-tuning feedback-regulating LIM kinase.
Cyclin D1 physically interacts with pRb, p160 (AIB1), cdks, and the cell cycle inhibitor proteins p21CIP1 and p27KIP1. p21CIP1 promotes cell motility in Ras-transformed cells, through forming a complex with ROCK and thereby blocking Rho kinase action (36). p27KIP1 regulates actin dynamics, promoting cell migration (40, 43) independently of cyclin-cdk binding, suggesting that the mechanisms by which p27KIP1 and cyclin D1 regulate motility are distinct (40). p27KIP1 can function upstream of RhoA, inhibiting its activation (6). p21CIP1 and p27KIP1 can either reduce cdk activity or promote assembly and nuclear transport of D-type cyclins (59). Thus, new possible interactions between cyclin D1 and p27KIP1 in regulating cellular migration may require further analysis. The association of cyclin D1 overexpression with poor prognosis and tumor metastasis (19, 30) raised the intriguing possibility that cyclin D1 may play a distinct role in promoting cellular migration and invasion. The present studies demonstrate cyclin D1 promotes cellular migration through TSP-1 and ROCKII. The identification of compounds that selectively block the ATP pocket of cdk to selectively inhibit cellular kinase activity has proven challenging. The current findings that the cyclin D1 protein, through K112, promotes cellular migration may provide an important new avenue for therapy of metastatic disease in which cyclin D1 is overexpressed.
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ACKNOWLEDGMENTS |
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Support for these studies was from NIH R01CA70896, R01CA75503, R01CA86072, R01CA93596, R01CA107382, and Kimmel Cancer Center Support grant P30 CA56036 to R.G.P.; CA26504, P01 100324-02, and P30 CA13330-31 to E.R.S.; and the Breast Cancer Alliance through an Exceptional Project grant to A.A.Q.
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REFERENCES |
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20 cells (**, P < 0.01) assessed at 5-min intervals for 2 h. (D and E) Single-cell analysis of cell movement velocity was taken at 5-min intervals from cyclin D1+/+ or cyclin E knockin to cyclin D1–/– MEFs. Cellular movement velocity is shown as the mean ± standard error for n 
