Tumor Suppressor p53 and Its Homologue p73{alpha} Affect Cell Migration*,

Anna A. Sablina {ddagger} §, Peter M. Chumakov {ddagger} ¶ || ** and Boris P. Kopnin § || {ddagger}{ddagger}

From the {ddagger}Lerner Research Institute, the Cleveland Clinic Foundation, Cleveland, Ohio 44195, §Institute of Carcinogenesis, Russian Cancer Research Center, Moscow, Russia 115478, and Engelhardt Institute of Molecular Biology, Moscow, Russia 119991

Received for publication, January 17, 2003 , and in revised form, May 13, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
The p53 tumor suppressor plays a central role in the negative control of growth and survival of abnormal cells. Previously we demonstrated that in addition to these functions, p53 expression affects cell morphology and lamellar activity of the cell edge (Alexandrova, A., Ivanov, A., Chumakov, P. M., Kopnin, P. B., and Vasiliev, J. M. (2000) Oncogene 19, 5826–5830). In the present work we studied the effects of p53 and its homologue p73{alpha} on cell migration. We found that loss of p53 function correlated with decreased cell migration that was analyzed by in vitro wound closure test and Boyden chamber assay. The decreased motility of p53-deficient cells was observed in different cell contexts: human foreskin fibroblasts (BJ), human colon and lung carcinoma cell lines (HCT116 and H1299, respectively), as well as mouse normal fibroblasts from lung and spleen, peritoneal macrophages, and keratinocytes. On the other hand, overexpression of the p53 family member p73{alpha} stimulated cell migration. Changes in cell migration correlated directly with transcription activation induced by p53 or p73{alpha}. Noteworthy, p53 modulated cell motility in the absence of stress. The effect of p53 and p73{alpha} on cell migration was mediated through the activity of the phosphatidylinositol 3-kinase/Rac1 pathway. This p53/p73 function was mainly associated with some modulation of intracellular signaling rather than with stimulation of production of secreted motogenic factors. The identified novel activity of the p53 family members might be involved in regulation of embryogenesis, wound healing, or inflammatory response.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
The p53 tumor suppressor is an essential component of an emergency stress-response mechanism that prevents growth and survival of damaged and abnormal cells. Various genotoxic stresses induce functional activity of p53, which consequently stimulates transcription of a set of genes involved in cell cycle checkpoints (p21waf1/cip1, gadd45, 14-3-3{sigma}, etc.) and apoptosis (insulin growth factor-binding protein 3, bax, fas/APO1, killer/DR5, Noxa, PIG3, etc.) (reviewed in Refs. 28). The p53 family members, p63 and p73, have affinity to similar DNA-binding sites and can up-regulate many of the p53-responsive genes. Although nearly half of the genes induced by p53 and p73{alpha} overlap (9), the common targets include the p21waf1/cip1, bax, gadd45, and 14-3-3{sigma} genes whose overexpression leads to growth arrest and/or apoptosis (1013).

In addition to the control of the cell cycle and apoptosis, the p53 family members are involved in other cellular processes. In particular, p53 participates in the DNA repair (4, 14), whereas p73 is involved in the neurogenesis (15, 16). Noteworthy, both p53 and p73 up-regulate a number of genes whose products are implicated in regulation of cell/matrix interactions and cell motility as was recently revealed by cDNA microarray hybridizations (9, 1719). The involvement of p53 in these processes is supported by other reports relating p53 to transcriptional regulation of the genes encoding smooth muscle {alpha}-actin (20), hepatocyte growth factor/scatter factor (21) and its receptor (22), fractalkine (23), HB-EGF1 (24), EGF receptor (25), metalloproteinase-2 (26), etc. Similarly, it was shown that p73 overexpression is associated with increased expression of vascular EGF, fibroblast growth factor 2, and platelet-derived growth factor (27). Although the physiological significance of the p53/p73-mediated up-regulation of the above target genes remains unclear, these data suggest involvement of p53 and p73 in the control of cell migration.

Cell migration is a key aspect of many normal and abnormal biological processes, including embryonic development, protection from infections, wound healing, and metastasis of tumor cells. It is generally accepted that the driving force for the cell movement is provided by the dynamic reorganization of the actin cytoskeleton, directing protrusion at the front of the cell and retraction at the rear. Cooperative action of small GTPases, particularly Rho, Cdc42, and Rac, is required to promote coordinated assembly and disassembly of actin filaments. Rho maintains cell adhesion during the movement and regulates the assembly of contractile actin-myosin filaments to form stress fibers. Cdc42 is required to maintain cell polarity, which includes localization of lamellipodial activity to the leading edge and the reorientation of the Golgi apparatus in the direction of the movement. Rac is necessary for the protrusion of lamellipodia and for the forward movement (reviewed in Refs. 28 and 29).

Previously, we demonstrated that overexpression of exogenous p53 in mouse fibroblasts causes diminution of focal contacts and an increase in lamellar activity of the cell edge that was paradoxically accompanied by inhibition of cell migration into the wound in vitro rather than by its stimulation (1). We explained such an effect by "skidding" of fibroblasts expressing high levels of exogenous p53 due to their inability to form effective focal contacts because of p53-induced repression of fibronectin production (1). The goal of this work was to study the effect of physiological levels of p53 on the Rho GTPase activity and cell migration in various cell contexts. Here we present the data demonstrating the ability of p53 as well as its homologue p73{alpha} to affect migration ability of different cell types through activation of the PI3-kinase/Rac1 pathway.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
Cell Cultures—p53-positive colon carcinoma HCT116 cell line, its derivative HCT116 p53–/– with the p53 gene knockout (30), and p53-deficient H1299 lung adenocarcinoma cell line were used. Sublines of the HCT116, HCT116 p53–/–, and H1299 cells expressing dominant-negative C-terminal fragment of rat p53 cDNA (GSE22 (31)), GFP, p73{alpha}, p73{alpha}His293, or p53His175 were generated by retrovirus-mediated gene transfer with pBabe/puro-GSE22, pBabe/puro-GFP, pPS/hygro-p73{alpha}, pPS/hygro-p73{alpha}His293, or pPS/hygro-p53His175. The HCT116, HCT116 p53–/–, and H1299 sublines with introduced empty pBabe/puro or pPS/hygro vectors were used as a control. Normal human foreskin fibroblasts BJ (ATCC CRL-2522) with inhibited p53 expression were obtained by infection with lentivirus vector pLV-si-p53-GFP expressing hairpin transcript corresponding to 19 bp of human p53 cDNA under the control of H1 RNA promoter and GFP under the control of the cytomegalovirus promoter. The H1299tet/wt-p53 and 10(1)tet/wt-p53 cell lines with tetracycline-repressible expression of the exogenous p53 have been described previously (32). Normal lung and spleen fibroblasts, peritoneal macrophages, and tail keratinocytes were isolated from 8-week-old wild type (p53+/+) or homozygous p53 gene knockout (p53–/–) C57BL mice obtained from The Jackson Laboratory.

All the cells except keratinocytes were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum (Fetaclone). The keratinocytes were cultured in keratinocyte/serum-free medium (Invitrogen) containing bovine pituitary extract (30 µg/ml, Invitrogen), recombinant EGF (0.1 ng/ml, Invitrogen), and 0.02 mM CaCl2. The H1299tet/wt-p53 and 10(1)tet/wt-p53 cells were grown in the presence of tetracycline (2 µg/ml).

Boyden Chamber Assay—Boyden chamber cell migration assay was performed using transwell chambers with 8-µm pore size membranes (BD Biosciences). The chambers were inserted into 24-well culture plates containing Dulbecco's modified Eagle's medium with either 0.5% bovine serum albumin, 10% FBS, 100 ng/ml EGF (Invitrogen), or conditioned media. To prepare the conditioned media, equal quantities of the cells were plated for 48 h before harvesting the culture media. After collection of the conditioned media, the cells were harvested and counted again. To equalize the conditioned media they were diluted in proportion to the cell quantities.

The cells (5 x 104) were loaded into the upper volume of the Boyden chambers. Non-migrated cells were removed with a cotton swab, and the cells were fixed with methanol for 15 min and stained with crystal violet. The migration activity was quantified by blind counting of the migrated cells on the lower surface of the membrane of at least 10 fields per chamber using a x20 objective.

Western Blot Analysis of Protein Expression—Whole cell extracts were lysed in ice-cold RIPA buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1% sodium deoxycholate, 1% Nonidet P-40, 0.1% SDS, 100 mM phenylmethylsulfonyl fluoride, 1 mM pepstatin A, and 1 mM E64). Protein concentration in the extracts was determined with a protein assay system (Bio-Rad). 100 µg of protein was separated on a 10% SDS-polyacrylamide gel and transferred to polyvinylidene difluoride membrane (Amersham Biosciences). The membranes were probed with antibodies specific to p53 (monoclonal antibody 421, a gift of Dr. A. J. Levine, or DO-1, Santa Cruz Biotechnology), p73{alpha} (Ab-1, Oncogene Research Products), PI3-kinase subunits p85{alpha} (polyclonal, Sigma) or p110 (polyclonal, Santa Cruz Biotechnology). The membranes were treated with secondary sheep anti-mouse horseradish peroxidase antibodies or anti-rabbit horseradish peroxidase (Jackson Immuno-Research). The filters were developed by ECL chemiluminescence reagents (PerkinElmer Life Sciences) according to the manufacturer's protocol.

Detection of Transcriptional Activity of p53 and p73{alpha}For luciferase assay, cells were transiently transfected using LipofectaAMINE reagent (Invitrogen) with 1 µg of the pWWp-GL2 expressing luciferase under the control of the wild type p21waf1/cip1 gene promoter (33). To control the efficiency of the transfection, the cells were co-transfected with 0.5 µg of pCH100 (Amersham Biosciences) expressing {beta}-galactosidase. The luciferase activity was measured using Dual-Light luminescent reporter gene assay system (Tropix) 48 h after the transfection.

{beta}-Galactosidase staining of HCT116-Waf1ConA cells expressing {beta}-galactosidase reporter gene under the p53-responsive promoter was performed as described previously (34).

Measuring Activity of the Rho GTPases—The cells were incubated in serum-free media for 24 h. Thirty minutes or 2 h after stimulation with 10% FBS or EGF (100 ng/ml), the cells were lysed in the MLB buffer (Upstate Biotechnology, Inc.). The amounts of GTP-bound Rac1 or GTP-bound Rho were determined using the Rac1 activation assay kit and the Rho activation assay kit (Upstate Biotechnology, Inc.), respectively, according to the manufacturer's protocol.

Measuring Activity of PI3-Kinase—The serum-starved cells were stimulated with 10% FBS and lysed at 4 °C in the lysis buffer (150 mM NaCl, 1% Triton X-100, 4 mM Na3VO4, 200 mM NaF, 10 mM EDTA, 2 mM phenylmethylsulfonyl fluoride, and 10% glycerol, pH 7.4). The lysates were incubated overnight at 4 °C with the anti-p85{alpha} antibodies (Sigma) and protein A-agarose. The beads were pelleted at 14,000 x g, washed three times with Buffer A (Tris-buffered saline, pH 7.5, 0.1% Nonidet P-40, and 100 µM Na3VO4), three times with the Buffer B (100 mM Tris, pH 7.5, 500 mM LiCl2, and 100 µM Na3VO4), and twice with the Buffer C (10 mM Tris, pH 7.5, 100 mM NaCl, 1 mM EDTA, and 100 µM Na3VO4). The pellets were resuspended in the Buffer D (50 mM Hepes, pH 7.5, 1 mM EDTA, and 100 µM NaH2PO3), and the activity of PI3-kinase was assessed by phosphorylation of a mixture of phosphatidylinositols in the presence of 40 µM ATP (1 µCi of [{gamma}-32P]ATP). The reaction was carried out for 15 min and stopped with 150 µl of ice-cold 1 N HCl. The lipids were extracted by vortexing with 450 µl of chloroform/methanol (1:1) mixture. After centrifugation the organic phase was washed twice with 200 µl of 1 N HCl, and 40-µl aliquots were separated on potassium oxalate-pretreated TLC plates (Whatman) with 35 ml of 2 N acetic acid and 65 ml of 1-propanol as a mobile phase. The TLC plates were dried and exposed to Fuji film.

The Wound Healing Experiments—Eight-week-old wild type and homozygous p53 gene knockout C57BL mice were anesthetized and shaved, and full-thickness skin wounds were inflicted by excising ~0.5 x 0.5-cm patches of the skin. The wounds were then allowed to dry and to form a scab. The wound closure was measured daily. The area of wounds was measured by covering each wound of an anesthetized animal with a transparent plastic film and recording the wound contour. The wound areas were quantified and expressed as a percentage of the initial wound size (100%). The mean values (n = 10 animals) were plotted for each time point.

Expression of p53 was analyzed in 5-day skin wounds by immunohistochemistry of paraffin-embedded wounded skin sections using standard protocol. The FL-293 antibodies (Santa Cruz Biotechnology) were used for the immunostaining of p53.


    RESULTS AND DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
Loss of p53 Activity Decreases Cell Migration—To study the influence of physiological levels of p53 on cell motility, we analyzed how inhibition of endogenous p53 expression affects cell behavior in wound closure test and Boyden chamber assay. The effect of p53 elimination on the ability of cells to migrate into the wound in vitro was determined in two cell contexts: human normal foreskin fibroblasts BJ and colon carcinoma HCT116 cells. Both inhibition of p53 expression by siRNA-p53 in BJ cells and the p53 gene knockout in HCT116 cells resulted in significantly lower ability to migrate into the mechanical scratch (in vitro wound) made on the surface of growing cell culture (Fig. 1, and Supplemental Material Figs. 1 and 2). This result was in disagreement with our previous observation that mouse fibroblasts 10(1) expressing high levels of exogenous p53 showed decreased cell migration into the wound in vitro despite increased lamellar activity. This paradoxical effect was probably a result of repression of fibronectin production induced by overexpressed p53 (1). In BJ cell cultures we found no difference between p53-positive and p53-negative cells in fibronectin production (see Supplemental Material, Fig. 3). Probably the differential effects of endogenous and exogenous p53 on fibronectin production in human BJ and mouse 10(1) fibroblasts, respectively, are responsible for the difference in their behavior in wound closure assay.



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FIG. 1.
Effect of the p53 expression on wound closure in vitro. A, the p53 expression in BJ and HCT116 cell cultures. Expression of both siRNA-p53 in BJ fibroblasts and the p53 gene knockout in HCT116 cells inhibited p53 expression as revealed by Western blotting using DO-1 monoclonal antibodies (anti-p53). B, influence of p53 expression on migration of BJ fibroblasts into the wound in vitro. Wounds were done on confluent BJ and BJ/siRNAp53 cell cultures by the pipette tip (arrowheads show the size of the initial wound). After incubation for 24 h the cells were fixed and stained by Oregon-Green phalloidin. To estimate cell migration the culture was stained by Hoechst 33258 24 h after wounding. The number of nuclei of cells migrated into the wound was counted. The quantified result of the experiment is represented on the histogram. The difference in migration levels between p53-positive and p53-negative cells was statistically significant (p = 0.02 by Student's t test). C, effect of p53 expression on wound closure in HCT116 cell cultures. The p53-positive HCT116 and the GFP-labeled HCT116 p53–/– cells were plated in equal proportions. A wound was made on confluent cell culture by a pipette tip (arrowheads show the size of initial wound). On the left part the mixture HCT116 and HCT116 p53–/–/GFP cells is shown by phase contrast microscopy, while on the right part one can see only the GFP-labeled HCT116 p53–/– cells. The wound was filled mainly by the unlabeled p53-positive HCT116 cells. To exclude the inhibitory effect of GFP expression on cell migration, we performed a similar experiment using GFP-labeled HCT116 and unlabeled HCT116 p53–/–. In this case GFP-labeled p53-positive HCT116 cells mainly migrated into the wound (see Supplemental Material, Fig. 1). (The experiments were repeated twice with similar results.)

 


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FIG. 2.
The p53 gene knockout in HCT116 cells is accompanied by decreased cell migration. Boyden chamber assay was performed using uncoated or Matrigel-coated 8-µm pore-size membranes. HCT116 and HCT116 p53–/– cells were allowed to migrate toward 10% FBS for 12 h. The cells migrated through Matrigel-coated membrane and stained with crystal violet are shown on the top of the figure. The migration was quantified by blind counting of the migrated cells on the lower surface of the membrane. At least 10 fields per chamber were counted using a x20 objective. The quantified result of the experiment is represented on the histogram. The difference in migration levels between p53-positive and p53-negative cells was statistically significant (p < 0.03 by Student's t test).

 


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FIG. 3.
Inactivation of p53 by the GSE22 decreases migration of the HCT116 cells. A, the expression level of p53 and its C-terminal fragment (GSE22) in the HCT116 cells as revealed by Western blotting using 421 monoclonal antibodies (anti-p53). The HCT116 cell line carrying pBabe/puro vector was used as a control (The data of one of three typical experiments are shown.) B, the activity of p53 reporter gene in control HCT116, HCT116 p53–/– cells, and their derivatives carrying dominant-negative GSE22. The cells were transiently transfected with the pWWp-GL2 construct expressing luciferase gene under the p53-responsive promoter. (Each point represents average data obtained from six samples). C, the ability of the control HCT116, HCT116 p53–/– cells, and their derivatives carrying dominant-negative GSE22 to migrate toward 10% FBS for 12 h. The difference in migration level between HCT116 and HCT116/GSE22 cells was statistically significant (p = 0.003). No significant difference in migration level was found by Student's t test between HCT116 p53–/– and HCT116 p53–/–/GSE22 cells (p = 0.4407).

 

The decreased cell motility of various p53-deficient cells was further confirmed by Boyden chamber assay. Indeed, the HCT116 p53–/– cells showed lower ability to migrate through both uncoated and Matrigel-coated 8-µm pore-size membranes (Fig. 2). The increased number of p53-positive HCT116 cells on the lower side of the membrane as compared with their p53-negative counterparts could not be explained by differences in cell attachment or cell proliferation/survival on the upper side of the filter (see Supplemental Material, Fig. 4). The difference in migration level between HCT116 and HCT116 p53–/– cells remained unchanged at different time points of incubation (Supplemental Material, Fig. 5). So the p53 gene knockout in HCT16 cells caused an inhibition rather than a delay of cell migration.



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FIG. 4.
The effect of p53 expression on cell migration in different cell contexts. A, BJ and BJ/siRNA-p53 cells were allowed to migrate toward 10% FBS for 6 h. (The experiment was repeated three times.) B, withdrawal of tetracycline from the culture media induced p53 expression in H1299tet/wt-p53 cells as revealed by Western blotting using DO-1 antibodies (anti-p53, shown on the bottom). Migration activity of H1299tet/wt-p53 cells in the presence or absence of tetracycline (2 µg/ml) was analyzed. (The experiment was repeated three times). C, cells of several types were obtained from wild type and homozygous p53 gene knockout mice. Spleen and lung fibroblasts and peritoneal macrophages were allowed to migrate toward 10% FBS for 6 h; keratinocytes were allowed to move toward EGF (100 ng/ml) for 18 h. (The experiment was repeated three times.)

 


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FIG. 5.
Overexpression of p73{alpha} increases cell migration. A, Western blotting revealed only a background level of endogenous p73{alpha} in control HCT116 and H1299 cells carrying pPS/hygro vector. Retroviral transfer of p73{alpha} increased its level in the both cell lines. (The result of one of three typical experiments is shown.) B, the activity of p53 reporter gene in control and p73{alpha} expressing HCT116 and H1299 cells. The cells were transiently transfected with the pWWp-GL2 expressing luciferase gene under the p53-responsive promoter. The HCT116 and H1299 cells bearing pPS/hygro vector were used as a control. (Each point represents the average of six samples.) C, migration activity toward 10% FBS of control and p73{alpha} expressing HCT116 and H1299 cells assayed for 12 h. (The experiment was repeated three times.)

 

To exclude that the effect observed in HCT116 cells was due to variations between different clones (the construction of the HCT116 p53-knockout cell line involved selection of individual cell clones), we used retroviral gene transfer to produce mass cultures of HCT116 cells expressing C-terminal dominant-negative fragment of p53 (GSE22 (31)). Introduction of GSE22 into the HCT116 cells decreased p53 transcriptional activity 5 times as measured by the luciferase reporter assay without significant changes in the levels of expressed p53 protein, whereas all HCT116 p53–/– derivatives showed background level of luciferase activity and no expression of the p53 protein (Fig. 3, A and B). Boyden chamber assay revealed that GSE22 expression dramatically decreased migration of the HCT116 cells without affecting motility of the HCT116 p53–/– cells (Fig. 3C). Hence, there is direct correlation between the level of p53 activity and the migration ability of the HCT116 cells.

The influence of p53 expression on cell motility was also observed in different cell systems. Inhibition of p53 expression in human BJ fibroblasts by siRNA-p53 resulted in decreased cell motility (Fig. 4A). On the other hand the tetracycline-regulated expression of the exogenous p53 in p53-deficient human lung carcinoma H1299 cell line considerably stimulated cell migration (Fig. 4B). Similar correlation between cell migration activity and p53 state was observed in various cells types derived from wild type and homozygous p53 gene knockout mice, in particular for lung and spleen fibroblasts, peritoneal macrophages, and keratinocytes. The most pronounced difference was observed for the peritoneal macrophages (Fig. 4B). Thus, the effect of p53 on cell motility was characteristic for both tumor and normal cells, although the extent of this effect depends on particular cell context.

Overexpression of the p53 Homologue p73{alpha} Stimulates Cell Migration—The p53 family member p73{alpha} displays a high degree of structural and functional similarity with p53 (10, 11, 13, 16) suggesting that p73{alpha} can also affect cell migration. Transcriptionally active p73{alpha} was overexpressed in the p53-positive HCT116 and in the p53-deficient H1299 cell lines (Fig. 5A). Overexpression of the p73{alpha} in the H1299 cells considerably enhanced both the p53-dependent luciferase reporter expression and the migration activity (Fig. 5, B and C). In the p53-positive HCT116 cells the effect of p73{alpha} was detectable but less significant as compared with the p53-deficient H1299 cells (Fig. 5, B and C). Hence, as with p53, the increase in p73{alpha} transcriptional activity was accompanied by stimulation of cell migration.

Transcriptional Activity Is Required for the p53- or p73{alpha}-dependent Stimulation of Cell Migration—To study whether p53 or p73{alpha} affects cell motility through their ability to activate transcription of the target genes, we introduced transcriptionally inactive mutants p53His175 (35, 36) or p73{alpha}His293 (10) into H1299 cells (Fig. 6A). Expression of either p53 or p73{alpha} mutant had no effect on cell motility (Fig. 6B), suggesting that transcriptional up-regulation of certain p53/p73{alpha} target genes is involved in the modulation of cell migration.



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FIG. 6.
Transcriptional activity is required for the p53/p73{alpha}-mediated cell migration. A, expression of transcriptionally inactive mutants p53His175 or p73{alpha}His293 in H1299 cells as revealed by Western blotting with the DO-1 (anti-p53) or Ab-1 (anti-p73) antibodies. The H1299 cells carrying pPS/hygro vector were used as a control. (The result of one of three typical experiments is shown.) B, migration of the control, p53His175, and p73{alpha}His293 expressing H1299 cells through 8-µm pore-size membranes toward 10% FBS. (The experiment was repeated three times.)

 

Because transcriptional activity of p53 and its family members can be modulated at post-translational level by various stresses, we tested whether p53 can affect motility under unstressed conditions. We investigated the following: (i) whether conditions of Boyden chamber assay or wounding in vitro induce transcriptional activation of p53 and (ii) whether the fraction of the migrated cells is enriched by the cells with transcriptionally activated p53. We used HCT116 cells carrying {beta}-galactosidase reporter gene under the control of the p53-responsive promoter. We found that below 2% of the cells were positively stained for {beta}-galactosidase 12 h after plating into Boyden chambers, whereas over half the cells became {beta}-galactosidase-positive after treatment with the DNA-damaging drug EMS (see Supplemental Material, Fig. 6). Moreover, we observed the same proportion of the {beta}-galactosidase-positive cells on both sides of the porous membrane (data not shown). No considerable increase of activity of p53 or its homologues was observed at the wound edge 24 h after wounding (see Supplemental Material Fig. 6). This result suggests that conditions of Boyden chamber assay or wound closure in vitro do not induce stress-related activation of p53. So even the "latent" form of p53 is capable of affecting cell migration. This idea is in agreement with the data that the unstressed latent form of p53 is bound to p53-binding sites within the p53-responsive genes, including the p21waf1/cip1 gene (37). Moreover, p53 expression under unstressed conditions induces significant changes in the gene expression profiles revealed by cDNA microarray hybridization (9, 18, 19, 38). The genes responsible for the p53-dependent modulation of cell motility might belong to the group of genes that p53 regulates under normal physiological conditions.

To test whether stress-induced activation of p53 can additionally stimulate cell motility, we analyzed migration of the cells treated with the DNA-damaging agent EMS or with the hypoxia-mimicking agent deferoxamine mesylate. Either of the treatments increased the expression level and transcriptional activity of p53 (data not shown). Boyden chamber assay carried in the presence of the drugs showed proportional increase in migration of both p53-positive and p53-negative cells (Fig. 7) suggesting significant p53-independent component in stress-induced activation of cell motility. Thus, the contribution of activated p53 into stimulation of cell migration under stress conditions can hardly be assessed.



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FIG. 7.
Effect of DNA damage and hypoxia on migration activity of p53-positive and p53-negative cells. The HCT116 and HCT116 p53–/– cells pretreated with ethyl methanesulfonate (EMS, Sigma, 600 µg/ml, 2 h) or deferoxamine mesylate (DFO, Sigma, 300 ng/ml, 12 h) were allowed to migrate toward 10% FBS for 12 h in the presence of corresponding drugs. (The experiment was repeated twice.)

 

The Rac1 and PI3-Kinase Activity Correlates with the Expression of p53 and p73{alpha}The GTPase proteins of the Rho family are known as key regulators of cell motility (reviewed in Refs. 28 and 29). To study possible mechanisms of the p53/p73{alpha}-mediated cell migration, we tested the effect of p53 or p73{alpha} expression on activity of the Rho family GTPases. We assessed the Rac1 activity by its ability to bind the effector Pak1 protein. In serum-starved HCT116 and HCT116 p53–/– cells, the active form of Rac1 was detectable at low level, and unlike Guo et al. (39) we failed to establish clear differences between p53-positive and p53-negative cells (see Supplemental Material, Fig. 7). After serum stimulation for 30 min or 2 h, the amount of the active GTP-bound Rac1 was considerably higher in p53-positive HCT116 cells as compared with their p53-negative counterparts (Fig. 8A and Supplemental Material, Fig. 7). Inactivation of p53 by GSE22 also decreased the Rac1 activity in HCT116 cells without affecting the HCT116 p53–/– cells (Fig. 8A). On the other hand, expression of p53 or p73{alpha} in the p53-deficient H1299 cells considerably increased the amount of active Rac1 (Fig. 8, A and B). Hence, the Rac1 activity correlates directly with the level of p53 or p73{alpha} expression. These data agree with our previous observations (1) of the p53-mediated increase in lamellar activity at the cell edge, because Rac1 is the essential inductor of membrane ruffling (reviewed in Refs. 28 and 29). Unlike changes in the Rac1 activity, the expression of p53 did not affect the activities of Rho (Fig. 8C) or Cdc42 (40) GTPases. The changes in activity of Rac1 can constitute a part of the main mechanism by which p53 and p73{alpha} modulate cell migration.



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FIG. 8.
The effect of p53 and p73{alpha} expression on activity of Rac1 and Rho GTPases. A, the p53 affects Rac1 activity. Serum-starved derivatives of HCT116 cells and the H1299tet/wt-p53 cells were stimulated with 10% FBS for 30 min. The amount of GTP-bound Rac1 in the lysates was determined by its binding to the effector Pak1 protein. Total Rac1 protein expression was assayed in 1/20 of the lysate. (The data of one of three typical experiments are shown.) B, the effect of p73{alpha} overexpression on Rac1 activity. The amount of the GTP-bound Rac1 in serum-stimulated HCT116 and H1299 cells bearing pPS/hygro vector or expressing p73{alpha} was determined as described above. (The data of one of three typical experiments are shown.) C, the effect of p53 on Rho activity. Serum-starved HCT116 cells with different p53 states were stimulated with 10% FBS for 30 min. The amount of GTP-bound Rho in the lysates was determined by its ability to bind the Rhotekinin domain. Total Rho protein was assayed in 1/20 of the lysate. (The data of one of three experiments are shown.)

 

One of the major regulators of Rac1 activity are guanine nucleotide exchange factors (GEFs). Over 30 different GEFs regulating the Rho family GTPases have been identified to date. Some of the GEFs, including representatives of the best characterized Vav family members, are activated by the second messengers generated by the enzymatic activity of PI3-kinase (reviewed in Refs. 28 and 41). Trying to learn more about possible pathways leading to the p53-mediated up-regulation of Rac1, we tested whether p53 expression could modulate PI3-kinase activity. We found that p53 expression did not alter the expression of p85 or p110 subunits of PI3-kinase (Fig. 9A). However, study of the PI3-kinase functional activity in p53-positive and p53-deficient serum-stimulated HCT116 cells showed that PI3-kinase precipitated from the p53-deficient cells had significantly lower phosphorylation activity than the enzyme from the parental HCT116 cells (Fig. 9B). Hence, we conclude that p53 may stimulate cell motility through the PI3-kinase/Rac1 pathway.



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FIG. 9.
Effect of the p53 expression on the PI3-kinase activity. A, the expression level of p85 and p110 subunits of PI3-kinase in HCT116 and HCT116 p53–/– cells as revealed by Western blotting using polyclonal antibodies (anti-p85 or anti-p110). (The data of one of three typical experiments are shown.) B, serum-starved HCT116 and HCT116 p53–/– cells were stimulated by 10% FBS for the indicated periods. PI3-kinase was precipitated from the obtained lysates by specific antibodies to the p85 subunit of the PI3-kinase. PI3-kinase activity was assessed by measuring phosphorylation of phosphatidylinositol mixtures in the presence of [{gamma}-32P]ATP. (The typical result of a thin layer chromatography is shown.)

 

The p53/p73{alpha}-induced Stimulation of Cell Migration Depends on Modulation of Intracellular Signaling Pathways Rather Than on Enhanced Production of Secreted Motogenic/Chemoattractant Factors—There are two possible mechanisms by which p53 could up-regulate the PI3-kinase/Rac1 pathway. First, the p53 expression can increase production of secreted motogenic/chemoattractant factors. Second, p53 could act downstream by modulating some steps of intracellular signaling involved in activation of PI3-kinase/Rac1 in response to motogenic factors. The former supposition is based on the data that p53 can up-regulate transcription of HB-EGF leading to activation the PI3-kinase/Akt pathway (24). Some other genes encoding motogenic factors proved to be direct transcriptional targets of p53 (17, 18, 21, 23, 24). Therefore, p53 may enhance the expression of motogenic cytokines such as HB-EGF, hepatocyte growth factor/SF, IGF1, etc. and consequently induces the activity of their receptors whose phosphorylation mediates activation of the PI3-kinase function.

To answer whether p53 or p73{alpha} affect cell migration through increased production of secreted motogenic/chemoattractant factors, we studied the changes in cell migration toward conditioned media obtained from the cells with different p53 state. The conditioned medium obtained from the cells expressing p53 or p73{alpha} showed only a slightly increased motogenic effect as compared with the medium conditioned by the cells with inhibited p53 function (Fig. 10A). In agreement with these data, we observed no changes in the migration activity of HCT116 cells toward 10(1)tet/wt-p53 cells after stimulation of p53 expression by withdrawal of tetracycline (Fig. 10B). This result allowed us to propose that p53/p73{alpha}-mediated stimulation of cell migration is mainly related to some changes in intracellular signaling rather than to enhanced production of secreted motogenic factors.



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FIG. 10.
Effect of the conditioned media from cells with different p53 and/or p73{alpha} state on cell migration. A, the migration of the HCT116 cells toward the conditioned media obtained from the cells with different p53/p73 state. The conditioned medium was prepared as described under "Experimental Procedures." The cells migrated through 8-µm pore-size membranes toward the conditioned media for 12 h. (The experiment was repeated twice.) B, the migration of HCT116 cells toward 10(1) cells with tetracycline-repressible expression of p53. 10(1)tet/wt-p53 cells cultivated in the presence or absence of tetracycline (2 µg/ml) for 3 days on the bottom of 24-well plate were used as the attracting cells. The tested HCT116 cells were plated on 8-µm membranes and allowed to migrate toward the 10(1)tet/wt-p53 cells in the presence or absence of tetracycline for 12 h. (The experiment was repeated twice.) The activation of p53 expression after tetracycline withdrawal was confirmed by immunofluorescent staining (not shown).

 

Next, we tested whether expression of p53 could modify cell response to motogenic factors, in particular to EGF. Stimulation with EGF of the p53-deficient HCT116/GSE22 cells resulted in significantly lower migration activity and proportion of the GTP-bound Rac1 as compared with the control p53-positive HCT116 cells (Fig. 11A). A similar result was obtained after addition of the serum that apparently contains a mixture of motogenic factors (Fig. 11A). Thus, p53-negative cells might have some defects in transduction signals from growth/motogenic factor receptors to PI3-kinase. This is consistent with the observation that the conditioned media additionally stimulates migration of p53-positive HCT116 cells, whereas it has almost no effect on the isogenic cells with the disrupted p53 gene (Fig. 10A).



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FIG. 11.
The p53 expression affects the response to motogenic factors. A, the migration activity and the amount of the GTP-bound Rac1 were determined in the HCT116 and HCT116 p53–/– cells stimulated by 10% FBS or EGF (100 ng/ml) (The experiment was repeated twice.) B, the influence of AG1478 (30 mM, Biomol) or wortmannin (50 nM, Biomol) on migration of the p53-positive and p53-deficient HCT116 cells and peritoneal macrophages isolated from p53+/+ and p53–/– C57BL mice. (The experiment was repeated twice.)

 

It should be noted that the EGF-dependent pathway does not seem to be the only and, probably, the major mechanism through which p53 affects cell motility. In fact, treatment with a specific inhibitor of EGF receptor (AG1478) only partially inhibited the migration and did not eliminate the difference in migration level of p53-positive and p53-negative HCT116 cells. Meanwhile, wortmannin, a specific inhibitor of PI3-kinase, almost completely inhibited migration of all tested cells (Fig. 11B). Moreover, treatment of peritoneal macrophages that may not express the EGF receptor (42, 43) with the AG1478 inhibitor did not affect migration of either p53-positive or p53-deficient cells (Fig. 11B). This allows us to suggest that different p53 targets are probably responsible for stimulation of cell migration in distinct cell contexts.

Analysis of gene expression profiles has revealed a number of p53- or p73{alpha}-regulated genes involved in positive control of cell migration, such as Eps8, Ha-Ras-1, ACK, GNA13, inositol-1,3,4-triphosphate 5/6-kinase, smooth muscle actin, etc. (9, 1719, 24, 38). Notably, the set of p53/p73{alpha}-up-regulated genes is cell type-specific. So it is likely that several different p53/p73 target genes are involved in positive control of cell migration. On the other hand, p53 and p73{alpha} can cause some cell type-specific changes in gene expression resulting in inhibition of directed cell migration such as repression of fibronectin in fibroblasts (1, 45), repression of metalloproteinase-2 in melanocytes (46), or yet unidentified modulation of downstream targets of Cdc42 in mouse embryo fibroblasts (40). Additional studies are required for identification and verification of particular p53/p73-dependent genes that are involved in regulation of cell migration in each cell context.

The Role of the p53/p73-regulated Cell Migration in Vivo— The question arises what is the physiological significance of the p53/p73-mediated modulation of cell migration. We assumed that such regulation might stimulate remodeling of tissues and, in particular, could facilitate wound healing. This idea is based on two observations. First, the p53-deficient cells showed decreased migration into the in vitro wound scratched on the surface of the plate with growing culture (Fig. 1). Second, the obtained data (data not shown) and the published data (47, 48) demonstrated that the re-epithelialization of skin wounds was accompanied by enhanced levels of p53 protein in the basal layers of the epidermis. However, measuring the kinetics of skin wound closing showed only marginal difference with a tendency of delayed wound healing in the p53 knockout mice (Fig. 12). The absence of a clear effect of p53 gene knockout could be explained by compensatory mechanisms. For example, the lack of p53 could lead to elevated transcriptional activity of the p53 family members, because p53 proved to up-regulate expression of the dominant-negative form of p73 (49, 50). Thus, the decrease in cell migration due to p53 deficiency could be compensated by stimulation of motility through increased functional activity of its family members, particularly p73. Hence, involvement of p53 in tissue remodeling and wound healing remains in question.



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FIG. 12.
Effect of p53 on wound healing in vivo. Kinetics of the skin wound closure in p53+/+ and p53–/– C57BL mice. Wound areas are given as percent of the initial wound size (100%). The mean values (n = 10) were plotted for each time point.

 

Involvement of the p53/p73-mediated regulation of cell motility in other physiological processes can be proposed. It can take place during inflammatory response directing certain type of cells to the sites of infection, or during embryogenesis to establish tissue patterns and to drive certain morphogenetic pathways. The role of p53 and p73 in inflammation response is still poorly studied. We found that the p53-positive peritoneal macrophages showed significantly higher motility (Fig. 4B) and thus could have increased phagocyte activity as well. Notably, it was shown that p73 knockout mice have severe defects in inflammation response (15). In addition, the p53 family members, particularly p73, are implicated in the control of cell migration during neurogenesis. Deficiency in p73 leads to a severe distortion of hippocampal formation suggesting that p73 function is important for proper differentiation and migration of neurons (15, 16). The p53 can be also implicated in neurogenesis, as a certain proportion of p53-deficient mice show defects in neuronal tube closure (44, 51). Moreover, p53 regulates a set of genes involved in neural crest cell migration and neural tube closure (19). We can speculate that the ability of p53 to modulate cell migration is a feature of its evolutional ancestors that, similar to p63 and p73, played an important role in ontogenesis. This feature could be attributed to a constitutive function of p53 in the absence of stress, which explains why stress activation is not important for p53-mediated increase in cell motility. Hence, our finding demonstrates a novel activity of p53 and p73{alpha} that might be related to obscure functions of the p53 family under normal physiological conditions.


    FOOTNOTES
 
* This work was supported by startup funds from the Lerner Research Institute (to P. M. C.), the International Research Scholars Program of the Howard Hughes Medical Institute (to B. P. K.), and the Russian Foundation for Basic Research (to A. A. S., P. M. C., and B. P. K.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

The on-line version of this article (available at http://www.jbc.org) contains Figs. 1-7. Back

|| Both senior authors contributed equally to this work. Back

** To whom correspondence may be addressed. Tel.: 216-444-9540; E-mail: chumakp{at}ccf.org.

{ddagger}{ddagger} To whom correspondence may be addressed. Tel.: 7-095-324-1739; E-mail: bkopnin{at}yahoo.com.

1 The abbreviations used are: HB-EGF, heparin-binding EGF-like growth factor; EGF, epidermal growth factor; PI3, phosphatidylinositol 3; GFP, green fluorescent protein; GSE22, genetic suppressor element 22; FBS, fetal bovine serum; EMS, ethyl methanesulfonate; GEF, guanine-nucleotide-exchange factor. Back


    ACKNOWLEDGMENTS
 
We are grateful to Dr. A. Alexandrova and Dr. J. Vasiliev for stimulating discussions.



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 

  1. Alexandrova, A., Ivanov, A., Chumakov, P. M., Kopnin, P. B., and Vasiliev, J. M. (2000) Oncogene 19, 5826–5830[CrossRef][Medline] [Order article via Infotrieve]
  2. el-Deiry, W. S. (1998) Semin. Cancer Biol. 8, 345–357[CrossRef][Medline] [Order article via Infotrieve]
  3. Amundson, S. A., Myers, T. G., and Fornace, A. J. J. (1998) Oncogene 17, 3287–3299[CrossRef][Medline] [Order article via Infotrieve]
  4. Ko, L. J., and Prives, C. (1996) Genes Dev. 10, 1054–1072[CrossRef][Medline] [Order article via Infotrieve]
  5. Prives, C., and Hall, P. A. (1999) J. Pathol. 187, 112–126[CrossRef][Medline] [Order article via Infotrieve]
  6. Sionov, R. V., and Haupt, Y. (1999) Oncogene 18, 6145–6157[CrossRef][Medline] [Order article via Infotrieve]
  7. Vogelstein, B., Lane, D., and Levine, A. J. (2000) Nature 408, 307–310[CrossRef][Medline] [Order article via Infotrieve]
  8. Vousden, K. (2000) Cell 103, 691–694[Medline] [Order article via Infotrieve]
  9. Fontemaggi, G., Kela, I., Amariglio, N., Rechavi, G., Krishnamurthy, J., Strano, S., Sacchi, A., Givol, D., and Blandino, G. (2002) J. Biol. Chem. 277, 43359–43368[Abstract/Free Full Text]
  10. Kaghad, M., Bonnet, H., Yang, A., Creancier, L., Biscan, J. C., Valent, A., Minty, A., Chalon, P., Lelias, J. M., Dumont, X., Ferrara, P., McKeon, F., and Caput, D. (1997) Cell 90, 809–819[Medline] [Order article via Infotrieve]
  11. Jost, C. A., Marin, M. C., and Kaelin, W. G., Jr. (1997) Nature 389, 191–194[CrossRef][Medline] [Order article via Infotrieve]
  12. Yang, A., Kaghad, M., Wang, Y., Gillett, E., Fleming, M. D., Dotsch, V., Andrews, N. C., Caput, D., and McKeon, F. (1998) Mol. Cell 2, 305–316[Medline] [Order article via Infotrieve]
  13. Zhu, J., Jiang, J., Zhou, W., and Chen, X. (1998) Cancer Res. 58, 5061–5065[Abstract]
  14. Albrechtsen, N., Dornreiter, I., Grosse, F., Kim, E., Wiesmuller, L., and Deppert, W. (1999) Oncogene 18, 7706–7717[CrossRef][Medline] [Order article via Infotrieve]
  15. Yang, A., Walker, N., Bronson, R., Kaghad, M., Oosterwegel, M., Bonnin, J., Vagner, C., Bonnet, H., Dikkes, P., Sharpe, A., McKeon, F., and Caput, D. (2000) Nature 404, 99–103[CrossRef][Medline] [Order article via Infotrieve]
  16. Yang, A., and McKeon, F. (2000) Nat. Rev. Mol. Cell Biol. 1, 199–207[CrossRef][Medline] [Order article via Infotrieve]
  17. Zhao, R., Gish, K., Murphy, M., Yin, Y., Notterman, D., Hoffman, W. H., Tom, E., Mack, D. H., and Levine, A. J. (2000) Genes Dev. 14, 981–993[Abstract/Free Full Text]
  18. Yu, J., Zhang, L., Hwang, P. M., Rago, C., Kinzler, K. W., and Vogelstein, B. (1999) Proc. Natl. Sci. Acad. U. S. A. 96, 14517–14522[Abstract/Free Full Text]
  19. Yoon, H., Liyanarachchi, S., Wright, F. A., Davuluri, R., Lockman, J. C., De La Chapelle, A., and Pellegata, N. S. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 15632–15637[Abstract/Free Full Text]
  20. Comer, K. A., Dennis, P. A., Armstrong, L., Catino, J. J., Kastan, M. B., and Kumar, C. C. (1998) Oncogene 16, 1299–12308[CrossRef][Medline] [Order article via Infotrieve]
  21. Metcalfe, A. M. J., Dixon, R. M., and Radda, G. C. (1997) Nucleic Acids Res. 25, 983–986[Abstract/Free Full Text]
  22. Seol, D. W., Chen, Q., Smith, M. L., and Zarnegar, R. (1999) J. Biol. Chem. 274, 3565–3572[Abstract/Free Full Text]
  23. Shiraishi, K., Fukuda, S., Mori, T., Matsuda, K., Yamaguchi, T., Tanikawa, C., Ogawa, M., Nakamura, Y., and Arakawa, H. (2000) Cancer Res. 60, 3722–3726[Abstract/Free Full Text]
  24. Fang, L., Li, G., Liu, G., Lee, S. W., and Aaronson, S. A. (2001) EMBO J. 20, 1931–1939[Abstract/Free Full Text]
  25. Ludes-Meyers, J. H., Subler, M. A., Shivakumar, C. V., Munoz, R. M., Jiang, P., Bigger, J. E., Brown, D. R., Deb, S. P., and Deb, S. (1996) Mol. Cell. Biol. 16, 6009–6019[Abstract]
  26. Bian, J., and Sun, Y. (1997) Mol. Cell. Biol. 17, 6330–6338[Abstract]
  27. Vikhanskaya, F., Bani, M. R., Borsotti, P., Ghilardi, C., Ceruti, R., Ghisleni, G., Marabese, M., Giavazzi, R., Broggini, M., and Taraboletti, G. (2001) Oncogene 20, 7293–7300[CrossRef][Medline] [Order article via Infotrieve]
  28. Bishop, A. L., and Hall, A. (2000) Biochem. J. 348, 241–255[CrossRef][Medline] [Order article via Infotrieve]
  29. Ridley, A. J. (2001) J. Cell Sci. 114, 2713–2722[Medline] [Order article via Infotrieve]
  30. Bunz, F., Dutriaux, A., Lengauer, C., Waldman, T., Zhou, S., Brown, J. P., Sedivy, J. M., Kinzler, K. W., and Vogelstein, B. (1998) Science 282, 1497–1500[Abstract/Free Full Text]
  31. Ossovskaya, V. S., Mazo, I. A., Chernov, M. V., Chernova, O. B., Strezoska, Z., Kondratov, R., Stark, G. R., Chumakov, P. M., and Gudkov, A. V. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 10309–10314[Abstract/Free Full Text]
  32. Pugacheva, E. N., Ivanov, A. V., Kravchenko, J. E., Kopnin, B. P., Levine, A. J., and Chumakov, P. M. (2002) Oncogene 21, 4595–4600[CrossRef][Medline] [Order article via Infotrieve]
  33. Buckbinder, L., Talbott, R., Seizinger, B. R., and Kley, N. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 10640–10644[Abstract/Free Full Text]
  34. Sablina, A. A., Ilyinskaya, G. V., Rubtsova, S. N., Agapova, L. S., Chumakov, P. M., and Kopnin, B. P. (1998) J. Cell Sci. 111, 977–984[Abstract/Free Full Text]
  35. Kern, S. E., Pietenpol, J. A., Thiagalingam, S., Seymour, A., Kinzler, K. W., and Vogelstein, B. (1992) Science 256, 827–830[Medline] [Order article via Infotrieve]
  36. Farmer, G., Bargonetti, J., Zhu, H., Friedman, P., Prywes, R., and Prives, C. (1992) Nature 358, 83–86[CrossRef][Medline] [Order article via Infotrieve]
  37. Kaeser, M. D., and Iggo, R. D. (2001) Proc. Natl. Acad. Sci. U. S. A. 99, 95–100[Medline] [Order article via Infotrieve]
  38. Kannan, K., Amariglio, N., Rechavi, G., Jakob-Hirsch, J., Kela, I., Kaminski, N., Getz, G., Domany, E., and Givol, D. (2001) Oncogene 20, 2225–2234[CrossRef][Medline] [Order article via Infotrieve]
  39. Guo, F., Gao, Y., Wang, L., and Zheng, Y. (2003) J. Biol. Chem. 278, 14414–14419[Abstract/Free Full Text]
  40. Gadea, G., Lapasset, L., Gauthier-Rouviere, C., and Roux, P. (2002) EMBO J. 21, 2373–2382[Abstract/Free Full Text]
  41. Cantrell, D. A. (2001) J. Cell Sci. 114, 1439–1445[Abstract/Free Full Text]
  42. Mograbi, B., Rochet, N., Imbert, V., Bourget, I., Bocciardi, R., Emiliozzi, C., and Rossi, B. (1997) Eur. Cytokine Netw. 8, 73–81[Medline] [Order article via Infotrieve]
  43. Hardie, W., Bejarano, P. A., Miller, M. A., Yankaskas, J. R., Ritter, J. H., Whitsett, J. A., and Korfhagen, T. R. (1999) Pediatr. Dev. Pathol. 2, 415–423[CrossRef][Medline] [Order article via Infotrieve]
  44. Sah, V. P., Attardi, L. D., Mulligan, G. J., Williams, B. O., Bronson, R. T., and Jacks, T. (1995) Nat. Genet. 10, 175–180[Medline] [Order article via Infotrieve]
  45. Iotsova, V., and Stehelin, D. (1996) Cell Growth Differ. 7, 629–634[Abstract]
  46. Toschi, E., Rota, R., Antonini, A., Melillo, G., and Capogrossi, M. C. (2000) J. Invest. Dermatol. 114, 1188–1194[Abstract/Free Full Text]
  47. Hausmann, R., Nerlich, A., and Betz, P. (1998) Int. J. Leg. Med. 111, 169–172[Medline] [Order article via Infotrieve]
  48. Kane, C. D., and Greenhalgh, D. G. (2000) Wound Repair Regen. 8, 45–58[Medline] [Order article via Infotrieve]
  49. Kartasheva, N. N., Contente, A., Lenz-Stoppler, C., Roth, J., and Dobbelstein, M. (2002) Oncogene 21, 4715–4727[CrossRef][Medline] [Order article via Infotrieve]
  50. Chen, X., Zheng, Y., Zhu, J., Jiang, J., and Wang, J. (2001) Oncogene 20, 769–774[CrossRef][Medline] [Order article via Infotrieve]
  51. Armstrong, J. F., Kaufman, M. H., Harrison, D. J., and Clarke, A. R. (1995) Curr. Biol. 5, 931–936[Medline] [Order article via Infotrieve]