Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M601959200 on March 23, 2006

J. Biol. Chem., Vol. 281, Issue 21, 14596-14603, May 26, 2006
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
281/21/14596    most recent
M601959200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Hizli, A. A.
Right arrow Articles by Black, J. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Hizli, A. A.
Right arrow Articles by Black, J. D.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Protein Kinase C {alpha} Signaling Inhibits Cyclin D1 Translation in Intestinal Epithelial Cells*Formula

A. Asli Hizli1, Adrian R. Black1, Marybeth A. Pysz2, and Jennifer D. Black3

From the Department of Pharmacology and Therapeutics, Roswell Park Cancer Institute, Buffalo, New York 14263

Received for publication, March 1, 2006


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cyclin D1 is a key regulator of cell proliferation, acting as a mitogen sensor and linking extracellular signaling to the cell cycle machinery. Strict control of cyclin D1 levels is critical for maintenance of tissue homeostasis. We have reported previously that protein kinase C {alpha} (PKC{alpha}), a negative regulator of cell growth in the intestinal epithelium, promotes rapid down-regulation of cyclin D1 (Frey, M. R., Clark, J. A., Leontieva, O., Uronis, J. M., Black, A. R., and Black, J. D. (2000) J. Cell Biol. 151, 763–778). The current study explores the mechanisms underlying PKC{alpha}-induced loss of cyclin D1 protein in non-transformed intestinal epithelial cells. Our findings exclude several mechanisms previously implicated in down-regulation of cyclin D1 during cell cycle exit/differentiation, including alterations in cyclin D1 mRNA expression and protein turnover. Instead, we identify PKC{alpha} as a novel repressor of cyclin D1 translation, acting at the level of cap-dependent initiation. Inhibition of cyclin D1 translation initiation is mediated by PKC{alpha}-induced hypophosphorylation/activation of the translational suppressor 4E-BP1, association of 4E-BP1 with the mRNA cap-binding protein eIF4E, and sequestration of cyclin D1 mRNA in 4E-BP1-associated complexes. Together, these post-transcriptional effects ensure rapid disappearance of the potent mitogenic molecule cyclin D1 during PKC{alpha}-induced cell cycle withdrawal in the intestinal epithelium.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Tight control of cell proliferation is essential for maintenance of normal homeostasis in self-renewing tissues such as the intestinal epithelium. Previous studies have identified protein kinase C (PKC)4 signaling as an important negative regulator of cell growth/cell cycle progression in intestinal epithelial cells (1–3). Several members of the PKC family are predominantly expressed/activated in non-proliferating and terminally differentiated intestinal cells (4–6), pointing to a major function in regulation of post-mitotic events in this tissue. Consistent with this role, we have demonstrated that PKC{alpha} signaling triggers a program of cell cycle withdrawal in non-transformed intestinal crypt cells, paralleling the membrane translocation/activation of this enzyme precisely at the point of growth arrest within intestinal crypts in situ (3, 4, 7, 8). Although the extracellular signals that trigger growth arrest in the intestine in situ remain poorly characterized, a possible physiological activator of PKC{alpha} in this system is transforming growth factor beta. This potent growth inhibitory factor is known to promote G0/G1 arrest in intestinal epithelial cells (9–11) and to activate PKC{alpha} signaling in other systems (12).

One of the earliest events following PKC{alpha} activation in intestinal epithelial cells is down-regulation of cyclin D1 (3, 7, 8), indicating that this molecule is an important target of PKC{alpha} control. Cyclin D1 is a key regulator of cell proliferation, acting as a mitogen sensor and linking extracellular signaling to the cell cycle machinery (13). Progression through early G1 involves the activity of holoenzymes consisting of cyclin D (D1, D2, or D3 depending on the cell type) in association with cdk4 or cdk6. Cyclin D-dependent kinases promote cell cycle progression by initiating phosphorylation/inactivation of the retinoblastoma growth suppressor protein in mid G1, a process that is completed later in G1 by cyclin E/cdk2. Cyclin D/cdk complexes also have an important non-catalytic function that involves sequestration of cdk-inhibitory proteins of the Cip/Kip family, thus relieving repression of cyclin E- and cyclin A-cdk2 activity.

Precise regulation of cyclin D1 accumulation is of critical importance. Increased expression of the molecule shortens the G1 interval in many cell types and can reduce/overcome dependence on physiological growth stimuli (14). Decreased levels of the protein, on the other hand, lengthen G1 phase and reduce proliferation (14, 15). Thus, cyclin D1 expression is subject to strict control at multiple levels, including transcription, message stability and nucleocytoplasmic transport, protein synthesis, and protein turnover (14, 16–21). Notably, aberrant overexpression of cyclin D1 is a key component of tumor development in various tissues, including the intestine (14, 22, 23), and both transcriptional and post-transcriptional mechanisms have been implicated in deregulation of cyclin D1 expression in tumors (14).

In this study, we have explored the mechanisms underlying PKC{alpha}-induced down-regulation of cyclin D1 in IEC-18 non-transformed intestinal crypt cells. Our analysis of cross-talk between PKC signaling and cyclin D1 control unveiled a novel function for PKC{alpha} as a negative regulator of cyclin D1 translation initiation. PKC{alpha} modulates the activity of key translational regulators, including eukaryotic translation initiation factor eIF4E and eIF4E-binding protein 1 (4E-BP1), to repress cyclin D1 protein synthesis during intestinal epithelial cell cycle withdrawal. Importantly, the engagement of translational rather than transcriptional mechanisms ensures a rapid effect (24), with disappearance of cyclin D1 protein preceding other hallmark events of cell cycle withdrawal, including induction of Cip/Kip cyclin-dependent kinase inhibitors and activation of the growth suppressor function of pocket proteins (3, 7, 8).


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—Anti-cyclin D1 (sc-753, sc-450), -PKC{alpha} (sc-8393), -PKC{delta} (sc-213), and -PKC{epsilon} (sc-214) antibodies were from Santa Cruz Biotechnology. Antibodies to eIF4E, phospho-Ser-9-GSK-3beta, 4E-BP1, and eIF2{alpha} were from Cell Signaling Technology. Anti-phospho-Ser-51-eIF2{alpha} antibody was from Stressgen, and anti-FLAG M2 Affinity Gel was from Sigma. Phorbol 12-myristate 13-acetate (PMA), phorbol 12,13-dibutyrate (PdBu), 1,2-dioctanoyl-sn-glycerol (DiC8), and lithium were from Sigma, and bryostatin 1 (Bryo) was from Biomol or LC Laboratories. Bisindolylmaleimide I and Gö6976 were from Alexis and Calbiochem, respectively.

Cell Culture and Drug Treatments—IEC-18 non-transformed rat intestinal epithelial cells were maintained as described (3). PKC{alpha}, -{delta}, and -{epsilon} were activated in subconfluent cells by treatment with PMA (100 nM), PDBu (1 µM), Bryo (100 nM), or DiC8 (20 µg/ml). Control cells were treated with the appropriate vehicle (ethanol or Me2SO). Membrane translocation/activation of PKC isozymes was analyzed by subcellular fractionation and anti-PKC isozyme immunoblotting as previously described (7, 8). For depletion of PKC{alpha}, -{delta}, and -{epsilon}, cells were treated with 1 µM PDBu for 16–24 h (7, 8). Selective down-regulation of PKC{delta} and -{epsilon} was accomplished by pulse treatment with 10 nM PMA for 15 min, followed by two washes in PBS and return to fresh medium for 24 h (7, 8). Inhibition of PKC{alpha}, -{delta}, and -{epsilon} activity, or of PKC{alpha} activity alone, was achieved using 5 µM bisindolylmaleimide I or 1 µM Gö6976, respectively (8).

Analysis of Protein Expression—Cell lysis and immunoblot analysis were performed as described previously (4, 8). Blots were routinely stained with 0.1% Fast Green (Sigma) to confirm equal loading and even transfer. Primary antibody dilutions were: 1:1000 for cyclin D1, p-GSK-3beta, p-eIF2{alpha}, PKC{alpha}, and PKC{epsilon}; 1:2000 for eIF4E, 4E-BP1, total eIF2{alpha}, and PKC{delta}.

Northern Blot Analysis—Total cellular and cytoplasmic RNA were isolated using the RNeasy system (Qiagen) as recommended by the manufacturer. Nuclear RNA was similarly isolated from the nuclear pellet generated during the cytoplasmic RNA isolation procedure. Northern blot analysis of RNA samples (10 µg) was performed using randomly primed, 32P-labeled probe corresponding to mouse cyclin D1 cDNA, and specific hybridization was detected by phosphorimaging as we have described (25).

Immunoprecipitation—Immunoprecipitation using 2–4 µg of sc-753 anti-cyclin D1 antibody and Protein A/G-plus-agarose slurry (Santa Cruz Biotechnology) was performed essentially as we have described (26). In some experiments, cells were lysed in 20 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, and protease/phosphatase inhibitor cocktails (Sigma), and extracts were incubated with 2–3 µg of sc-450 anti-cyclin D1 antibody. For immunoprecipitation of FLAG-tagged cyclin D1, transfected cells were lysed in 20 mM Tris, pH 7.6, 120 mM NaCl, 0.5% Nonidet P-40, and protease/phosphatase inhibitor cocktails, and lysates were incubated with 50 µl of anti-FLAG M2 Affinity Gel.

Analysis of Protein Stability—Cells were incubated in methionine-free Dulbecco's modified Eagle's medium containing 4 mM glutamine, 5 µg/ml of insulin, and 5% dialyzed fetal bovine serum for 30 min and then labeled with 94 µCi of [35S]Met/Cys Expre35S35S protein-labeling mixture (PerkinElmer Life Sciences) for 1 h, prior to the addition of PMA or vehicle (ethanol) for 1 h. [35S]Met/Cys was chased for various times in label-free complete medium containing PMA or vehicle, and cells were harvested for cyclin D1 immunoprecipitation using sc-753 anti-cyclin D1 antibody. Immunoprecipitates were resolved on 20% SDS-PAGE gels, transferred to nitrocellulose membrane, and 35S-labeled protein was detected and quantified using the Storm/ImageQuant PhosporImaging system (Amersham Biosciences). For analysis of cyclin D1 decay following protein synthesis inhibition, cells were treated with 50 µg/ml of cycloheximide after 1 h of PMA/vehicle treatment and harvested for Western blot analysis at various times thereafter.

Biosynthetic Labeling—IEC-18 cells were incubated in methionine-free medium as described above and treated with PMA or ethanol for 30 min. Cells were then incubated in [35S]Met/Cys and PMA/vehicle for the indicated times as described (19). Cyclin D1 was immunoprecipitated using sc-450 antibody, and immunoprecipitates were analyzed as above.

Plasmids and Transfection—FLAG-tagged wild-type and T286A mutant cyclin D1 constructs were gifts from Dr. C. Sherr (St. Jude Children's Research Hospital, Memphis, TN). Destruction box mutants (R29Q and L32A) were generated from the wild-type cyclin D1 construct using the QuikChange site-directed mutagenesis kit (Stratagene) and appropriate primers (sense strands: GGGTGCTGCAGGCCATGCTCAAG; GCGAGCCATGGCCAAGACGGAG). Cells in 60-mm plates were transfected with 5 µg of each construct (0.5 µg for the T286A mutant) using Lipofectamine 2000 (Invitrogen).

In Vitro Cap Affinity Assay—Cap affinity chromatography was performed essentially as described (27). PMA- or vehicle-treated IEC-18 cells were harvested in lysis buffer (50 mM Tris, pH 7.4, 100 mM NaCl, 1 mM EDTA, and 1% Triton X-100, supplemented with protease/phosphatase inhibitor cocktails), and particulate material was removed by centrifugation. Cell lysates (350–400 µg of protein) were incubated with 50 µl of 7-methyl-GTP-Sepharose 4B slurry (Amersham Biosciences) for 16 h at 4 °C. Beads were washed three times with lysis buffer, and cap-bound protein was eluted with Laemmli sample buffer and subjected to immunoblot analysis.

Immunoprecipitation-RT-PCR Assay—PMA- or vehicle-treated cells were harvested in detergent-free lysis buffer (50 mM Tris-HCl, pH 7.5, 100 mM NaCl, 0.1 mM EDTA, 1 mM MgCl2, and 10% glycerol, containing protease/phosphatase inhibitors and 56 units/ml of RNase inhibitor (Sigma)) (28). Following removal of particulate material by centrifugation, lysates were supplemented with Tween 20 (0.5%), RNase inhibitor (53 units/ml), and dithiothreitol (0.1 mM), and 4E-BP1-containing complexes were immunoprecipitated using rabbit anti-4E-BP1 antibody (1.8 µg) and Protein A Dynabeads (Dynal). Beads were then washed twice (5 min) with lysis buffer containing 0.5% Tween 20 and once with lysis buffer without Tween 20 and resuspended in 45 µl of lysis buffer. 25 µl were taken for RNA isolation as described (28), and 20 µl were boiled in 2x Laemmli sample buffer for immunoblot analysis. Levels of cyclin D1 mRNA were assessed by RT-PCR in the presence of 50 µCi of [{alpha}32P]CTP using the Titanium one-step RT-PCR kit (Clonetech) and specific primers (GTCTTCCCGCTGGCCATGAACTAC; AAGAAAGTGCGTTGTGCGGTAGCA). PCR cycle number was empirically determined to be in the linear range of amplification. PCR products were resolved on 5% PAGE gels and visualized by phosphorimaging. Quantitative RT-PCR analysis of cyclin D1 mRNA levels in immunoprecipitates was performed using the Full Velocity SYBR Green QRT-PCR Master Mix (Stratagene) and a 7300 real-time PCR system (Applied Biosciences).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
PKC/PKC{alpha} Signaling Promotes Down-regulation of Cyclin D1 in Intestinal Epithelial Cells
We have previously shown that activation of PKC signaling in IEC-18 non-transformed intestinal crypt cells results in cell cycle withdrawal (3, 8). Maintenance of cell cycle arrest requires sustained activation of PKC{alpha}, and the effect is reversed coincident with down-regulation of the enzyme. As shown in Fig. 1, PKC-induced cell cycle blockade is associated with marked changes in the levels of cyclin D1. Activation of PKC{alpha}, -{delta}, and/or -{epsilon} (the only phorbol ester-responsive PKC isozymes expressed in IEC-18 cells) (3, 8) by PMA results in rapid down-regulation of cyclin D1 protein, detectable by 30 min and maximal by ~2 h (Fig. 1A). Down-regulation of cyclin D1 is also seen in response to PKC activation by the phorbol ester PdBu, the macrocyclic lactone Bryo, and the diacylglycerol analog DiC8 (Fig. 1B), excluding potential nonspecific effects of PMA treatment (29). PKC agonist-induced down-regulation of cyclin D1 is blocked by the general PKC inhibitor bisindolylmaleimide I and by PKC{alpha}, -{delta}, and -{epsilon} depletion with 1 µM PdBu (Fig. 1C), confirming that the effect is PKC dependent. Notably, the PKC{alpha}-selective inhibitor Gö6976 also prevents the effect of PKC agonists, pointing to a key role for PKC{alpha} in regulating cyclin D1 levels in intestinal epithelial cells (Fig. 1D).


Figure 1
View larger version (26K):
[in this window]
[in a new window]
  		FIGURE 1.
Activation of PKC/PKC{alpha} in IEC-18 cells promotes down-regulation of cyclin D1. A, IEC-18 cells were treated with 100 nM PMA for the indicated times, and cyclin D1 expression was determined by immunoblot analysis. The loading control is a nonspecific band consistently seen on cyclin D1 immunoblots that is not affected by PKC agonist treatment. C, control cells. IB, immunoblot. B, effects of 100 nM PMA (P), 1 µM PdBu (PB), 100 nM Bryo (B), or 20 µg/ml DiC8 (D) on cyclin D1 protein levels in IEC-18 cells. C, control cells. C and D, PMA-induced down-regulation of cyclin D1 is PKC/PKC{alpha} dependent. C, cells were pretreated (30 min) with the general PKC inhibitor bisindolylmaleimide I (BIM)(5µM) or were depleted of PKC{alpha}, -{delta}, and -{epsilon} by prolonged incubation with PdBU as described (7) (PKC-depleted) prior to addition of PMA for 2 h. D, cells were pretreated (30 min) with the PKC{alpha}-selective inhibitor Gö6976 (1 µM) and treated with PMA as in panel C. The increased levels of cyclin D1 seen in PKC-depleted and PKC inhibitor-treated IEC-18 cells reflects relief of a repressive effect of unstimulated/basal PKC/PKC{alpha} signaling on cyclin D1 accumulation in cycling cells. Data are representative of at least three independent experiments.

 
PKC/PKC{alpha}-induced Down-regulation of Cyclin D1 Does Not Involve Alterations in mRNA Levels/Subcellular Distribution or Protein Stability
Effects of PKC Signaling on Cyclin D1 mRNA—To investigate the molecular mechanisms underlying PKC-induced loss of cyclin D1, the effects of PMA on cyclin D1 mRNA were examined by Northern blotting. As shown in Fig. 2, PMA did not affect total cyclin D1 mRNA levels at 1 h of treatment, a time when cyclin D1 protein was significantly down-regulated (Fig. 1A), although a slight decrease in mRNA levels was observed at later times (e.g. 2 h, data not shown). Analysis of nuclear and cytosolic mRNA further excluded the possibility that PKC activation affects cyclin D1 mRNA nucleocytoplasmic distribution (Fig. 2). Thus, PKC-induced reduction in cyclin D1 protein does not involve effects on the levels or distribution of cyclin D1 mRNA, at least during the initial stages of PMA treatment.

PKC/PKC{alpha} Does Not Alter Cyclin D1 Protein Turnover—Cell cycle exit (e.g. under conditions of serum starvation or cell differentiation) has been linked to accelerated turnover of cyclin D1 protein (20, 30). Therefore, [35S]methionine pulse-chase experiments were performed to determine whether PKC activation alters cyclin D1 stability in IEC-18 cells. As expected, PMA treatment resulted in lower recovery of 35S in cyclin D1 immunoprecipitates, reflecting the reduced cyclin D1 protein levels at the time points examined (1–2.5 h of PMA treatment) (Fig. 3A). However, PKC activation had no effect on the rate of decay of cyclin D1 protein (t1/2 ~ 30 min), a result that was confirmed by examining cyclin D1 disappearance following inhibition of protein synthesis with cycloheximide (data not shown).


Figure 2
View larger version (45K):
[in this window]
[in a new window]
  		FIGURE 2.
PKC/PKC{alpha}-induced down-regulation of cyclin D1 does not involve changes in cyclin D1 mRNA levels or subcellular distribution. Cells were treated with PMA or vehicle for the indicated times, and RNA was isolated from total cell lysates (left panel) or from cytosolic and nuclear subcellular fractions (right panel). RNA was analyzed for cyclin D1 message by Northern blotting and phosphorimaging. The lower panel shows ethidium bromide-stained 18 S RNA in corresponding lanes. Data are representative of three independent experiments.

 
Degradation of cyclin D1 occurs via ubiquitin/proteasome-mediated proteolysis, triggered by its phosphorylation on Thr-286 by GSK-3beta (20). An N-terminal (amino acids 29–32) destruction box (RXXL) has also been implicated in regulating proteasomal degradation of cyclin D1 (18). Consistent with the absence of alterations in cyclin D1 stability, PKC activation did not decrease the levels of inhibitory phosphorylation of GSK-3beta on Ser-9 (Fig. 3B, upper panel), and inhibition of GSK-3beta activity with Li+ did not block PMA-induced loss of cyclin D1 (although its accumulation following Li+ treatment confirmed a role for GSK-3beta in normal turnover of cyclin D1 in IEC-18 cells) (middle panel). These data, together with the fact that PKC activation promoted the down-regulation of degradation-resistant cyclin D1 mutants, including the GSK-3beta phosphosite mutant T286A (TA, lower panel) and the destruction box mutants R29Q (RQ) and L32A (LA) (Fig. 3C), confirmed that PKC-induced loss of cyclin D1 protein does not involve acceleration of cyclin D1 turnover.

PKC/PKC{alpha} Signaling Inhibits Translation of Cyclin D1 in IEC-18 Cells
Activation of PKC/PKC{alpha} Inhibits Cyclin D1 Synthesis—[35S]Methionine biosynthetic labeling was performed to determine whether PKC signaling inhibits translation of cyclin D1. A reduction in radiolabeled cyclin D1 was detected in PMA-treated cells relative to vehicle-treated cells at each time point examined (Fig. 4). These data, together with the failure of PMA to affect cyclin D1 mRNA levels or protein stability (Figs. 2 and 3), point to repression of cyclin D1 translation as a mechanism underlying loss of cyclin D1 in PKC agonist-treated cells.


Figure 3
View larger version (23K):
[in this window]
[in a new window]
  		FIGURE 3.
PKC/PKC{alpha} activation does not alter the stability of cyclin D1 protein in IEC-18 cells. A, [35S]Met/Cys pulse-chase analysis of cyclin D1 protein degradation. Cells were labeled with [35S]Met/Cys and, following 1 h of PMA or vehicle treatment, were transferred to chase medium in the presence or absence of PMA. Cyclin D1 was immunoprecipitated at the indicated times of chase, and 35S-labeling was quantified by SDS-PAGE and phosphorimaging (lower panel). [35S]Met/Cys-labeled DLD1 colon carcinoma cells were used as a control for the cyclin D1 immunoprecipitation procedure. B, PKC agonist-induced cyclin D1 down-regulation does not involve GSK-3beta. Upper panel, IEC-18 cells were treated with PMA for the indicated times and subjected to immunoblot analysis of inhibitory phosphorylation of GSK-3beta on Ser-9. Middle panel, immunoblot analysis of cyclin D1 levels in cells pretreated with the GSK-3beta inhibitor LiCl (30 or 50 mM) for 16 h prior to addition of PMA for 2 h. Lower panel, immunoblot analysis of cyclin D1 levels in vehicle- or PMA-treated cells (2 h) expressing FLAG-tagged wild-type (wt) or T286A mutant (TA) cyclin D1. The identity of the slower migrating band as FLAG-tagged cyclin D1 was confirmed by comparison of its mobility with that of FLAG-tagged cyclin D1 immunoprecipitated with anti-FLAG M2 antibody (IP: Flag). C, the RXXL destruction box is not required for PKC-induced loss of cyclin D1. Immunoblot analysis of cyclin D1 levels in whole cell lysates (upper panel) or FLAG immunoprecipitates (lower panel) from cells expressing FLAG-tagged wild-type (wt) or destruction box mutant (R29Q (RQ) or L32A (LA)) cyclin D1 following treatment with PMA or vehicle for 2 h. All data are representative of at least three independent experiments.

 


Figure 4
View larger version (31K):
[in this window]
[in a new window]
  		FIGURE 4.
PKC/PKC{alpha} signaling inhibits cyclin D1 synthesis. Cells were labeled with [35S]Met/Cys for the indicated times, starting at 45 min of PMA or vehicle treatment. [35S] labeling of immunoprecipitated cyclin D1 was quantified by SDS-PAGE and phosphorimaging. [35S]Met/Cys-labeled MCF7 breast cancer cells were used as a control for the cyclin D1 immunoprecipitation procedure. It should be noted that methionine starvation leads to decreased levels of cyclin D1 protein in control cells, an effect that is associated with inhibition of translation (data not shown and Refs. 16 and 54). Thus, although data from metabolic labeling analysis clearly demonstrate that PKC activation leads to reduced synthesis of cyclin D1, they are likely to underestimate the true extent of this effect. Data are representative of three independent experiments.

 
PKC/PKC{alpha} Activates 4E-BP1, Inhibits Cap-dependent Translation Initiation, and Leads to Sequestration of Cyclin D1 mRNA in 4E-BP1-associated Complexes—To gain insight into the molecular basis for PKC-mediated inhibition of cyclin D1 synthesis, we examined the effects of PKC activation on key regulators of translation initiation, including eIF4E and eIF2{alpha}. eIF4E binds the 5'-cap structure (m7GpppN, where N is any nucleotide) found in the majority of eukaryotic mRNAs and associates with eIF4G and eIF4A to form eIF4F, a translation initiation complex that mediates recruitment of ribosomes to mRNA (31). A major mechanism for control of eIF4E function is through its interaction with a family of translational inhibitory proteins, the eIF4E-binding proteins (4E-BPs), the best characterized of which is 4E-BP1 (31). The hypophosphorylated {alpha} and beta forms of 4E-BP1 compete with eIF4G for binding to eIF4E, thereby preventing eIF4F assembly and inhibiting cap-dependent translation. In contrast, the hyperphosphorylated {gamma} form is unable to bind eIF4E. Immunoblot analysis revealed that PMA treatment leads to rapid hypophosphorylation/activation of 4E-BP1 (Fig. 5A), evident by the appearance of the faster migrating {alpha}/beta forms within 15 min. 4E-BP1 hypophosphorylation was also seen with other PKC agonists including PdBu, Bryo, and DiC8 (Fig. 5B), and the PKC dependence of the effect was confirmed using PKC{alpha}-, PKC{delta}-, and PKC{epsilon}-depleted cells (Fig. 5C). Use of cells depleted of PKC{delta} and -{epsilon}, but not PKC{alpha}, by PMA pulse treatment (7) demonstrated that PKC{alpha} can mediate PMA-induced hypophosphorylation of 4E-BP1 (Fig. 5D), and a requisite role for PKC{alpha} signaling was confirmed using the PKC{alpha}-selective inhibitor Gö6976 (Fig. 5E).

To determine whether the PKC-induced accumulation of 4E-BP1 {alpha}/beta phosphoforms reflected an increase in 4E-BP1 activity, the interaction of 4E-BP1 with eIF4E was examined using cap affinity chromatography. Extracts from IEC-18 cells treated with vehicle or PMA were incubated with Sepharose 4B-immobilized 7-methyl-GTP cap analog to capture eIF4E and its binding partners. Serum-stimulated and serum-starved cells were included as negative and positive controls for 4E-BP1/eIF4E interaction, respectively. Immunoblot analysis of cap analog-bound proteins revealed a striking increase in levels of cap-bound 4E-BP1 {alpha}/beta phosphoforms in PKC agonist-treated cells, comparable with those in serum-starved cells (Fig. 5F). Consistent with inhibition of cap-dependent translation initiation, metabolic labeling studies revealed that PKC agonist treatment results in a reduction in overall protein synthesis in IEC-18 cells (supplemental Fig. S1).


Figure 5
View larger version (50K):
[in this window]
[in a new window]
  		FIGURE 5.
PKC/PKC{alpha} signaling represses cyclin D1 translation initiation. PKC/PKC{alpha} signaling activates 4E-BP1 in IEC-18 cells. A, immunoblot analysis of 4E-BP1 in cells treated with PMA or vehicle for the indicated times. Arrows indicate the phosphoforms of 4E-BP1. The loading control is a nonspecific band that is not affected by PMA treatment. B, immunoblot analysis of 4E-BP1 in cells treated with 100 nM PMA (P), 1 µM PdBu (PB), 100 nM Bryo (B), or 20 µg/ml of DiC8 (D). C, control. C, PMA-induced activation of 4E-BP1 is PKC dependent. Immunoblot analysis of 4E-BP1 in PMA or vehicle-treated (30 min) IEC-18 cells expressing PKC{alpha}, -{delta}, and -{epsilon} or depleted of these enzymes by prolonged exposure to 1 µM PdBu (PKC-depleted). D, PKC{alpha} signaling is required for PMA-induced 4E-BP1 activation. PKC{delta}/{epsilon}-depleted cells were generated as described under "Experimental Procedures"; selective down-regulation of PKC{delta} and PKC{epsilon} was confirmed by immunoblot analysis of PKC isozymes (left panel). Control and PKC{delta}/{epsilon}-depleted cells were treated with vehicle or PMA for 15 min prior to being subjected to anti-4E-BP1 immunoblot analysis (right panel). E, IEC-18 cells were pretreated with 1µM Gö6976 for 30 min as indicated. PMA or vehicle was then added for 2 h, and cell extracts were subjected to immunoblot analysis for 4E-BP1 and cyclin D1. F, PKC signaling promotes the interaction of 4E-BP1 with the mRNA cap structure. Cells were treated with PMA for the indicated times, serum stimulated by an increase in serum from 5 to 10% for 10 min (Serum STIM.), or serum starved for 22 h (Serum STV.). Cell extracts were incubated with Sepharose-conjugated m7-GTP cap analog, and cap-bound fractions together with corresponding whole cell lysates (WCL) were subjected to immunoblot analysis for 4E-BP1 and eIF4E. M, MW markers. Note the absence of the inactive {gamma} form of 4E-BP1 in cap-bound fractions. G, PKC signaling promotes sequestration of cyclin D1 mRNA in 4E-BP1/eIF4E-associated complexes. 4E-BP1 was immunoprecipitated from control, PMA-treated, serum-stimulated, or serum-starved cells. WCL and immunoprecipitates were subjected to immunoblot analysis for eIF4E and 4E-BP1 (upper panel). N.S., nonspecific band; M, MW markers. Levels of cyclin D1 RNA in immunoprecipitates were analyzed by semiquantitative RT-PCR (middle panel). Parallel immunoprecipitation was performed with anti-PKC{alpha} antibody (confirmed by immunoblot analysis, middle panel) as a negative control for 4E-BP1/cyclin D1 mRNA complex formation. Levels of cyclin D1 mRNA (relative to control) in cell extracts (Input) and immunoprecipitates (I.P.) were also assessed by quantitative real-time RT-PCR (lower panel). This analysis revealed an ~3-fold increase in levels of 4E-BP1-associated cyclin D1 mRNA in PMA-treated and serum-starved cells. All data are representative of three or more independent experiments.

 
To examine the relevance of changes in 4E-BP1 activity to cyclin D1 translation, an immunoprecipitation RT-PCR protocol was used to assess sequestration of cyclin D1 mRNA in 4E-BP1-associated complexes. 4E-BP1 was immunoprecipitated from lysates of IEC-18 cells treated with PMA or vehicle. Immunoblot analysis confirmed immunoprecipitation of 4E-BP1 phosphoforms and increased association of eIF4E with 4E-BP1 in PMA-treated and serum-starved cells (Fig. 5G, upper panel). Semiquantitative RT-PCR analysis of 4E-BP1 immunoprecipitates revealed that PKC activation leads to accumulation of cyclin D1 mRNA in 4E-BP1-associated complexes, comparable with levels seen in serum-starved cells (middle panel). The specificity of the interaction was confirmed by the absence of cyclin D1 mRNA in PKC{alpha} immunoprecipitates. Quantitative real-time RT-PCR (lower panel) determined that PKC activation leads to an ~3-fold increase in the association of 4E-BP1 with cyclin D1 mRNA. As seen in cap binding assays, this effect was comparable with that induced by serum starvation. Thus, the increased activity of 4E-BP1 resulting from PKC activation directly impacts cap-dependent translation of cyclin D1 mRNA in intestinal epithelial cells.

Translation initiation is also regulated by eIF2{alpha}, a factor involved in recruitment of initiator Met-tRNA to the 40 S ribosomal subunit (32). The activity of eIF2{alpha} is negatively regulated by phosphorylation on Ser-51. Immunoblot analysis revealed that PMA treatment has no effect on eIF2{alpha} phosphorylation (Fig. 6), arguing against its involvement in the inhibitory effects of PKC signaling on cyclin D1 in IEC-18 cells.

Loss of PKC/PKC{alpha} Signaling Results in Recovery of the Translational Machinery and Restoration of Cyclin D1 Levels
Members of the PKC family undergo down-regulation following activation (33). In IEC-18 cells, PMA treatment results in progressive down-regulation of phorbol ester-responsive PKC isozymes, albeit with different kinetics (Fig. 7A). PKC{delta} is down-regulated first, followed by PKC{alpha} and then PKC{epsilon}. Activation-induced depletion/desensitization of PKC isozymes was associated with a gradual restoration of cyclin D1 protein expression, first noted at ~3 h of PMA treatment (Fig. 7B). Comparison of the timing of cyclin D1 recovery with the kinetics of PKC isozyme down-regulation demonstrated that the reappearance of cyclin D1 coincides more closely with loss of PKC{alpha} (see 3-h time point), a finding that further supports the role of PKC{alpha} as a key negative regulator of cyclin D1 accumulation in IEC-18 cells. As shown in Fig. 7, A and B, PKC{alpha} expression remains steady for ~2 h of PMA treatment, paralleling the loss of cyclin D1 protein, and levels of the enzyme are considerably reduced by 3 h, coincident with restoration of cyclin D1 levels. In contrast, PKC{delta} is markedly down-regulated by 1 h of PMA addition, and PKC{epsilon} expression is maintained for longer than 4 h in these cells. Loss of PKC/PKC{alpha} signaling was also accompanied by reversal of PMA-induced hypophosphorylation/activation of 4E-BP1 (Fig. 7C, upper panel) and decreased association of the protein with cap analog (lower panel). Thus, PKC down-regulation is accompanied by recovery of the translational machinery and restoration of cyclin D1 levels in IEC-18 cells.


Figure 6
View larger version (18K):
[in this window]
[in a new window]
  		FIGURE 6.
The translation initiation factor eIF2{alpha} is not involved in PKC{alpha}-induced down-regulation of cyclin D1. PMA- or vehicle-treated cells were evaluated for inhibitory phosphorylation on Ser-51 of eIF2{alpha} by immunoblot analysis.

 


Figure 7
View larger version (23K):
[in this window]
[in a new window]
  		FIGURE 7.
Loss of PKC{alpha} signaling is associated with restoration of cyclin D1 levels and recovery of the translational machinery. A, immunoblot analysis of PKC isozyme levels in IEC-18 cells treated with PMA for the indicated times. PKC isozymes are down-regulated following prolonged treatment with PMA. PKC{delta} is down-regulated first, followed by PKC{alpha}. PKC{epsilon} expression is maintained for longer than 4 h in these cells. B, immunoblot analysis of cyclin D1 expression at various times of PMA treatment. C, vehicle-treated cells collected at 0.25 h (first lane) and 3.5 h (last lane). PMA-induced down-regulation of cyclin D1 is maximal at 2–2.5 h, and loss of PKC{alpha} at ~3 h (see panel A) correlates with the initial recovery of cyclin D1 protein levels. C, down-regulation of PKC{alpha} is associated with reversal of 4E-BP1 hypophosphorylation. Upper panel, cells were treated with PMA or vehicle for the indicated times, and WCL were subjected to IB analysis for 4E-BP1. Lower panel, extracts from PMA-treated, serum-stimulated, or serum-starved cells were incubated with Sepharose-conjugated m7-GTP cap analog, and cap-bound fractions were subjected to immunoblot analysis for 4E-BP1 and eIF4E. All data are representative of at least three independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Cyclin D1 levels are low in proliferating intestinal crypt cells and are markedly decreased in post-mitotic cells (3, 34–36). This expression profile points to the existence of mechanisms for tight control of the accumulation of this key G1 progression factor in normal intestinal epithelial cells. Here we have identified PKC{alpha}, an important mediator of growth inhibitory signaling in a variety of biological systems (e.g. Refs. 1, 37, 38), as a negative regulator of cyclin D1 expression in the intestinal epithelium. The involvement of PKC{alpha} in mediating down-regulation of cyclin D1 was established by demonstrating that (a) PKC agonists promote loss of cyclin D1 in IEC-18 cells expressing PKC{alpha} as the only phorbol ester-responsive isozyme (3), (b) the PKC{alpha}-selective inhibitor Gö6976 blocks PMA-induced cyclin D1 down-regulation (Fig. 1), and (c) recovery of cyclin D1 levels closely correlates with loss of PKC{alpha} expression/activity in this system (Fig. 7).

Down-regulation of cyclin D1 is an early consequence of PKC{alpha} activation in IEC-18 cells, evident by 15–30 min of PKC agonist treatment (Figs. 1, 7). Analysis of the molecular mechanisms underlying the effect excluded several pathways previously implicated in the disappearance of cyclin D1 during cell cycle arrest/cell differentiation. Northern blot analysis revealed no change in cyclin D1 mRNA levels or subcellular distribution during the initial phase of the response (Fig. 2), indicating that PKC-induced down-regulation of cyclin D1 does not involve altered availability of cyclin D1 mRNA (21, 39). Pulse-chase analysis further excluded a role for alterations in protein turnover (20), a conclusion that was substantiated by the ability of PKC/PKC{alpha} to promote down-regulation of GSK-3beta phosphorylation site (T286A) and destruction box (R29Q and L32A) cyclin D1 mutants in IEC-18 cells (Fig. 3).

Instead, our findings provide the first evidence for the ability of PKC{alpha} signaling to repress cyclin D1 synthesis by targeting cap-dependent initiation, a key point of translational control (Fig. 5). Inhibition of cap-dependent initiation is accomplished by rapid, PKC{alpha}-dependent hypophosphorylation/activation of the translational repressor 4E-BP1 and increased association of the {alpha}/beta phosphoforms of the protein with eIF4E (Fig. 5), thus preventing assembly of the eIF4F complex that mediates recruitment of ribosomes to mRNA (31). Accumulation of cyclin D1 mRNA in 4E-BP1-associated complexes provides support for a direct effect on translation of cyclin D1, consistent with findings that active 4E-BP1 represses cyclin D1 synthesis in other systems (40). The marked effects of 4E-BP1 on cyclin D1 synthesis are also consistent with evidence that limited availability of eIF4F complexes selectively impairs translation of several strong growth promoters, including cyclin D1, whose mRNAs are weak for translation due to lengthy and highly structured 5'-untranslated regions (41).

A number of studies have reported effects of PKC signaling on translation initiation factors and modulators, including 4E-BP1 (42–44). However, in direct contrast to our findings, these studies have linked PKC signaling to enhanced translation initiation and phosphorylation/inactivation of 4E-BP1. This discrepancy likely reflects different functions of individual PKC family members. For example, in 293T and glioblastoma cells, inactivation of 4E-BP1 was attributed to the novel PKCs, PKC{delta} and PKC{eta}, respectively (42, 44). The finding that calcium ionophore can modulate the effects of PMA on the translational machinery in lymphocytes (45) suggests that cross-talk between signaling pathways may also underlie some of this discrepancy.

Taken together, the data demonstrate that, during cell cycle withdrawal, PKC{alpha} promotes down-regulation of cyclin D1 protein by engaging translational rather than transcriptional mechanisms, thus ensuring a rapid effect (24, 46, 47). Indeed, loss of cyclin D1 precedes other PKC-induced cell cycle-related events such as induction of p21WAF1/CIP1 and p27KIP1 and down-regulation of cyclin A (3, 7), arguing that targeting cyclin D1 is a key mechanism by which PKC{alpha} signaling promotes cell cycle exit in these cells. Cyclin D1 down-regulation is also an early event during intestinal epithelial growth arrest in situ. Although cyclins D1, E, and A are all expressed in proliferating intestinal crypt cells, only cyclin D1 is down-regulated near the point of growth arrest in the crypt region; changes in the levels of cyclins E and A occur only after the cells have migrated onto the villus (3, 34, 36, 48). Because PKC{alpha} undergoes membrane translocation/activation precisely at the point within intestinal crypts at which cells cease dividing (3, 4, 7, 8), the enzyme is appropriately positioned to promote cyclin D1 loss in vivo.

The finding that PKC{alpha}-induced cell cycle exit is associated with translational repression of cyclin D1 is consistent with data from studies with other G1 arresting signals. For example, blockade of cyclin D1 synthesis is associated with transforming growth factor beta-induced G0/G1 arrest in IEC-6 and RIE non-transformed intestinal cells (11). Given that transforming growth factor beta can activate PKC{alpha} (12), it is tempting to speculate that effects of transforming growth factor beta-signaling on cyclin D1 expression are mediated by PKC{alpha} in the intestinal epithelium. The relevance of translational regulation of cyclin D1 to intestinal homeostasis is highlighted by the fact that overexpression of eIF4E, together with associated up-regulation of cyclin D1 (21, 49), has been implicated in the development of colon tumors (14, 23, 35, 41, 50, 51). In this regard, it is noteworthy that PKC{alpha} is frequently down-regulated/inactivated in colon adenomas and adenocarcinomas as well as in other epithelial cancers (5, 37, 52, 53). Thus, loss of PKC{alpha}-mediated negative regulation of critical translational regulators and cell cycle control molecules is likely to play an important role in epithelial tumorigenesis.


   FOOTNOTES
 
* This work was supported in part by National Institutes of Health Grants DK60632, DK54909, and CA16056 and by a grant from the Mae Stone Goode/Roswell Park Alliance Foundation. 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

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. S1. Back

1 Both authors contributed equally to this work. Back

2 Supported by National Institutes of Health Postdoctoral Fellowship CA113048 [GenBank] . Back

3 To whom correspondence should be addressed: Dept. of Pharmacology and Therapeutics, Roswell Park Cancer Inst., Elm and Carlton Sts., Buffalo, NY 14263. Tel.: 716-845-5766; Fax: 716-845-8857; E-mail: jennifer.black{at}roswellpark.org.

4 The abbreviations used are: PKC, protein kinase C; Bryo, bryostatin 1; DiC8, 1,2-dioctanoyl-sn-glycerol; 4E-BP1, eIF4E-binding protein 1; eIF4E, eukaryotic initiation factor 4E; PMA, phorbol 12-myristate 13-acetate; PdBu, phorbol 12,13-dibutyrate; RT-PCR, reverse transcription PCR. Back


   ACKNOWLEDGMENTS
 
We thank Joseph Marinaro, Laura Kunneva, and Michael Fox for expert technical assistance, Dr. Olga Leontieva for help with PKC isozyme analysis, and Drs. E. Berleth, G. Das, and A. Karpf for critical review of the manuscript. We also thank Dr. C. Sherr for providing FLAG-tagged cyclin D1 constructs.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Black, J. D. (2000) Front Biosci. 5, D406–D423[Medline] [Order article via Infotrieve]
  2. Di Mari, J. F., Mifflin, R. C., and Powell, D. W. (2005) Gastroenterology 128, 2131–2146[CrossRef][Medline] [Order article via Infotrieve]
  3. Frey, M. R., Clark, J. A., Leontieva, O., Uronis, J. M., Black, A. R., and Black, J. D. (2000) J. Cell Biol. 151, 763–778[Abstract/Free Full Text]
  4. Saxon, M. L., Zhao, X., and Black, J. D. (1994) J. Cell Biol. 126, 747–763[Abstract/Free Full Text]
  5. Verstovsek, G., Byrd, A., Frey, M. R., Petrelli, N. J., and Black, J. D. (1998) Gastroenterology 115, 75–85[CrossRef][Medline] [Order article via Infotrieve]
  6. Jiang, Y. H., Aukema, H. M., Davidson, L. A., Lupton, J. R., and Chapkin, R. S. (1995) Cell Growth Differ. 6, 1381–1386[Abstract]
  7. Frey, M. R., Saxon, M. L., Zhao, X., Rollins, A., Evans, S. S., and Black, J. D. (1997) J. Biol. Chem. 272, 9424–9435[Abstract/Free Full Text]
  8. Clark, J. A., Black, A. R., Leontieva, O. V., Frey, M. R., Pysz, M. A., Kunneva, L., Woloszynska-Read, A., Roy, D., and Black, J. D. (2004) J. Biol. Chem. 279, 9233–9247[Abstract/Free Full Text]
  9. Ko, T. C., Beauchamp, R. D., Townsend, C. M., Jr., Thompson, E. A., and Thompson, J. C. (1994) Am. J. Surg. 167, 14–19[CrossRef][Medline] [Order article via Infotrieve]
  10. Kurokowa, M., Lynch, K., and Podolsky, D. K. (1987) Biochem. Biophys. Res. Commun. 142, 775–782[CrossRef][Medline] [Order article via Infotrieve]
  11. Ko, T. C., Yu, W., Sakai, T., Sheng, H., Shao, J., Beauchamp, R. D., and Thompson, E. A. (1998) Oncogene 16, 3445–3454[CrossRef][Medline] [Order article via Infotrieve]
  12. Sakaguchi, M., Miyazaki, M., Sonegawa, H., Kashiwagi, M., Ohba, M., Kuroki, T., Namba, M., and Huh, N. H. (2004) J. Cell Biol. 164, 979–984[Abstract/Free Full Text]
  13. Sherr, C. J. (1996) Science 274, 1672–1677[Abstract/Free Full Text]
  14. Weinstein, I. B. (2000) Carcinogenesis 21, 857–864[Abstract/Free Full Text]
  15. Filmus, J., Robles, A. I., Shi, W., Wong, M. J., Colombo, L. L., and Conti, C. J. (1994) Oncogene 9, 3627–3633[Medline] [Order article via Infotrieve]
  16. Guo, Y., Harwalkar, J., Stacey, D. W., and Hitomi, M. (2005) Oncogene 24, 1032–1042[CrossRef][Medline] [Order article via Infotrieve]
  17. Amanatullah, D. F., Zafonte, B. T., Albanese, C., Fu, M., Messiers, C., Hassell, J., and Pestell, R. G. (2001) Methods Enzymol. 333, 116–127[Medline] [Order article via Infotrieve]
  18. Agami, R., and Bernards, R. (2000) Cell 102, 55–66[CrossRef][Medline] [Order article via Infotrieve]
  19. Brewer, J. W., Hendershot, L. M., Sherr, C. J., and Diehl, J. A. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 8505–8510[Abstract/Free Full Text]
  20. Diehl, J. A., Zindy, F., and Sherr, C. J. (1997) Genes Dev. 11, 957–972[Abstract/Free Full Text]
  21. Rousseau, D., Kaspar, R., Rosenwald, I., Gehrke, L., and Sonenberg, N. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 1065–1070[Abstract/Free Full Text]
  22. Sansom, O. J., Reed, K. R., van de Wetering, M., Muncan, V., Winton, D. J., Clevers, H., and Clarke, A. R. (2005) J. Biol. Chem. 280, 28463–28467[Abstract/Free Full Text]
  23. Hulit, J., Wang, C., Li, Z., Albanese, C., Rao, M., Di Vizio, D., Shah, S., Byers, S. W., Mahmood, R., Augenlicht, L. H., Russell, R., and Pestell, R. G. (2004) Mol. Cell. Biol. 24, 7598–7611[Abstract/Free Full Text]
  24. Rajasekhar, V. K., and Holland, E. C. (2004) Oncogene 23, 3248–3264[CrossRef][Medline] [Order article via Infotrieve]
  25. Bateman, N. W., Tan, D., Pestell, R. G., Black, J. D., and Black, A. R. (2004) J. Biol. Chem. 279, 12093–12101[Abstract/Free Full Text]
  26. Leontieva, O. V., and Black, J. D. (2004) J. Biol. Chem. 279, 5788–5801[Abstract/Free Full Text]
  27. Caron, S., Charon, M., Cramer, E., Sonenberg, N., and Dusanter-Fourt, I. (2004) Mol. Cell. Biol. 24, 4920–4928[Abstract/Free Full Text]
  28. Fisher, T. S., Taggart, A. K., and Zakian, V. A. (2004) Nat. Struct. Mol. Biol. 11, 1198–1205[CrossRef][Medline] [Order article via Infotrieve]
  29. Kazanietz, M. G. (2000) Mol. Carcinog. 28, 5–11[CrossRef][Medline] [Order article via Infotrieve]
  30. Spinella, M. J., Freemantle, S. J., Sekula, D., Chang, J. H., Christie, A. J., and Dmitrovsky, E. (1999) J. Biol. Chem. 274, 22013–22018[Abstract/Free Full Text]
  31. Gingras, A. C., Raught, B., and Sonenberg, N. (1999) Annu. Rev. Biochem. 68, 913–963[CrossRef][Medline] [Order article via Infotrieve]
  32. Clemens, M. J. (2004) Oncogene 23, 3180–3188[CrossRef][Medline] [Order article via Infotrieve]
  33. Parker, P. J., Bosca, L., Dekker, L., Goode, N. T., Hajibagheri, N., and Hansra, G. (1995) Biochem. Soc. Trans. 23, 153–155[Medline] [Order article via Infotrieve]
  34. Zhang, T., Nanney, L. B., Luongo, C., Lamps, L., Heppner, K. J., DuBois, R. N., and Beauchamp, R. D. (1997) Cancer Res. 57, 169–175[Abstract/Free Full Text]
  35. Rosenwald, I. B., Chen, J. J., Wang, S., Savas, L., London, I. M., and Pullman, J. (1999) Oncogene 18, 2507–2517[CrossRef][Medline] [Order article via Infotrieve]
  36. Chandrasekaran, C., Coopersmith, C. M., and Gordon, J. I. (1996) J. Biol. Chem. 271, 28414–28421[Abstract/Free Full Text]
  37. Tibudan, S. S., Wang, Y., and Denning, M. F. (2002) J. Investig. Dermatol. 119, 1282–1289[CrossRef][Medline] [Order article via Infotrieve]
  38. Detjen, K. M., Brembeck, F. H., Welzel, M., Kaiser, A., Haller, H., Wiedenmann, B., and Rosewicz, S. (2000) J. Cell Sci. 113(Pt 17), 3025–3035[Abstract]
  39. Page, K., Li, J., Corbit, K. C., Rumilla, K. M., Soh, J. W., Weinstein, I. B., Albanese, C., Pestell, R. G., Rosner, M. R., and Hershenson, M. B. (2002) Am. J. Respir. Cell Mol. Biol. 27, 204–213[Abstract/Free Full Text]
  40. Jiang, H., Coleman, J., Miskimins, R., and Miskimins, W. K. (2003) Cancer Cell Int. 3, 2[CrossRef][Medline] [Order article via Infotrieve]
  41. De Benedetti, A., and Graff, J. R. (2004) Oncogene 23, 3189–3199[CrossRef][Medline] [Order article via Infotrieve]
  42. Aeder, S. E., Martin, P. M., Soh, J. W., and Hussaini, I. M. (2004) Oncogene 23, 9062–9069[CrossRef][Medline] [Order article via Infotrieve]
  43. Herbert, T. P., Kilhams, G. R., Batty, I. H., and Proud, C. G. (2000) J. Biol. Chem. 275, 11249–11256[Abstract/Free Full Text]
  44. Kumar, V., Pandey, P., Sabatini, D., Kumar, M., Majumder, P. K., Bharti, A., Carmichael, G., Kufe, D., and Kharbanda, S. (2000) EMBO J. 19, 1087–1097[CrossRef][Medline] [Order article via Infotrieve]
  45. Miyamoto, S., Kimball, S. R., and Safer, B. (2000) Biochim. Biophys. Acta 1494, 28–42[Medline] [Order article via Infotrieve]
  46. Rajasekhar, V. K., Viale, A., Socci, N. D., Wiedmann, M., Hu, X., and Holland, E. C. (2003) Mol. Cell 12, 889–901[CrossRef][Medline] [Order article via Infotrieve]
  47. Campo, P. A., Das, S., Hsiang, C. H., Bui, T., Samuel, C. E., and Straus, D. S. (2002) Cell Growth Differ. 13, 409–420[Abstract/Free Full Text]
  48. Andreu, P., Colnot, S., Godard, C., Gad, S., Chafey, P., Niwa-Kawakita, M., Laurent-Puig, P., Kahn, A., Robine, S., Perret, C., and Romagnolo, B. (2005) Development 132, 1443–1451[Abstract/Free Full Text]
  49. Rosenwald, I. B., Kaspar, R., Rousseau, D., Gehrke, L., Leboulch, P., Chen, J. J., Schmidt, E. V., Sonenberg, N., and London, I. M. (1995) J. Biol. Chem. 270, 21176–21180[Abstract/Free Full Text]
  50. Dua, K., Williams, T. M., and Beretta, L. (2001) Proteomics 1, 1191–1199[CrossRef][Medline] [Order article via Infotrieve]
  51. Tan, A., Bitterman, P., Sonenberg, N., Peterson, M., and Polunovsky, V. (2000) Oncogene 19, 1437–1447[CrossRef][Medline] [Order article via Infotrieve]
  52. Neill, G. W., Ghali, L. R., Green, J. L., Ikram, M. S., Philpott, M. P., and Quinn, A. G. (2003) Cancer Res. 63, 4692–4697[Abstract/Free Full Text]
  53. Zhu, Y., Dong, Q., Tan, B. J., Lim, W. G., Zhou, S., and Duan, W. (2005) Cancer Res. 65, 4520–4524[Abstract/Free Full Text]
  54. Fafournoux, P., Bruhat, A., and Jousse, C. (2000) Biochem. J. 351, 1–12[CrossRef][Medline] [Order article via Infotrieve]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 Mol. Biol. CellHome page
R. Tapia, M. Huerta, S. Islas, A. Avila-Flores, E. Lopez-Bayghen, J. Weiske, O. Huber, and L. Gonzalez-Mariscal
Zona Occludens-2 Inhibits Cyclin D1 Expression and Cell Proliferation and Exhibits Changes in Localization along the Cell Cycle
Mol. Biol. Cell, February 1, 2009; 20(3): 1102 - 1117.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
L. Guan, K. Song, M. A. Pysz, K. J. Curry, A. A. Hizli, D. Danielpour, A. R. Black, and J. D. Black
Protein Kinase C-mediated Down-regulation of Cyclin D1 Involves Activation of the Translational Repressor 4E-BP1 via a Phosphoinositide 3-Kinase/Akt-independent, Protein Phosphatase 2A-dependent Mechanism in Intestinal Epithelial Cells
J. Biol. Chem., May 11, 2007; 282(19): 14213 - 14225.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
281/21/14596    most recent
M601959200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Hizli, A. A.
Right arrow Articles by Black, J. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Hizli, A. A.
Right arrow Articles by Black, J. D.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2006 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement