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Cell Growth & Differentiation Vol. 13, 409-420, September 2002
© 2002 American Association for Cancer Research

Translational Regulation of Cyclin D1 by 15-Deoxy-{Delta}12,14-Prostaglandin J21

Peggy A. Campo, Sonali Das, Chin-Hui Hsiang, Tim Bui, Charles E. Samuel and Daniel S. Straus2

Biomedical Sciences Division and Biology Department, University of California, Riverside, California 92521-0121 [P. A. C., C-H. H., T. B., D. S. S.], and Department of Molecular, Cellular, and Developmental Biology, University of California, Santa Barbara, California 93106 [S. D., C. E. S.]


    Abstract
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
The D-group cyclins play a key role in the progression of cells through the G1 phase of the cell cycle. Treatment of MCF-7 breast cancer cells with the cyclopentenone prostaglandin 15-deoxy-{Delta}12,14-PGJ2 (15d-PGJ2) results in rapid down-regulation of cyclin D1 protein expression and growth arrest in the G0/G1 phase of the cell cycle. 15d-PGJ2 also down-regulates the expression of cyclin D1 mRNA; however, this effect is delayed relative to the effect on cyclin D1 protein levels, suggesting that the regulation of cyclin D1 occurs at least partly at the level of translation or protein turnover. Treatment of MCF-7 cells with 15d-PGJ2 leads to a rapid increase in the phosphorylation of protein synthesis initiation factor eukaryotic initiation factor 2{alpha} (eIF-2{alpha}) and a shift of cyclin D1 mRNA from the polysome-associated to free mRNA fraction, indicating that 15d-PGJ2 inhibits the initiation of cyclin D1 mRNA translation. The selective rapid decrease in cyclin D1 protein accumulation is facilitated by its rapid turnover (t1/2=34 min) after inhibition of cyclin D1 protein synthesis. The half-life of cyclin D1 protein is not significantly altered in cells treated with 15d-PGJ2. Treatment of cells with 15d-PGJ2 results in strong induction of heat shock protein 70 (HSP70) gene expression, suggesting that 15d-PGJ2 might activate protein kinase R (PKR), an eIF-2{alpha} kinase shown previously to be responsive to agents that induce stress. 15d-PGJ2 strongly stimulates eIF-2{alpha} phosphorylation and down-regulates cyclin D1 expression in a cell line derived from wild-type mouse embryo fibroblasts but has an attenuated effect in PKR-null cells, providing evidence that PKR is involved in mediating the effect of 15d-PGJ2 on eIF-2{alpha} phosphorylation and cyclin D1 expression. In summary, treatment of MCF-7 cells with 15d-PGJ2 results in increased phosphorylation of eIF-2{alpha} and inhibition of cyclin D1 mRNA translation initiation. At later time points, repression of cyclin D1 mRNA expression may also contribute to the decrease in cyclin D1 protein.


    Introduction
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Transit of normal mammalian cells through G1 and into the S-phase of the cell cycle requires the action of mitogens and is controlled by CDKs3 that are sequentially activated by cyclins D, E, and A (1, 2, 3, 4, 5) . The D-type cyclins (D1, D2, and/or D3, depending on the cell type) act as sensors for the presence of mitogenic factors. Treatment of quiescent cells with mitogens causes increased expression of D-type cyclins, thereby leading to the activation of CDK4 or CDK6. Activated CDK4 or CDK6 complexes promote progression through G1 phase in two ways. First, these complexes catalyze the first in a series of site-specific phosphorylations of Rb that ultimately inactivate Rb as a transcriptional repressor (6) . Second, these complexes sequester the CDK inhibitors p21CIP1/WAF1 and p27KIP1, thus relieving the inhibitory effect of the CDK inhibitors on the catalytic activity of cyclin E and cyclin A-CDK2 complexes (7) . Evidence suggests that mitogens regulate the expression/activity of cyclin D1/CDK complexes at several levels, including cyclin D1 gene transcription (8, 9, 10) and cyclin D1 translation (11) , nuclear localization and turnover of cyclin D1 protein (12) , and assembly of active cyclin/CDK complexes (13) .

Overexpression of cyclin D1 has been implicated in the etiology of a number of types of human cancer (14) . Increased expression of cyclin D1 is observed in ~50% of invasive primary breast carcinomas by immunohistochemical staining (15) . In about 15–20% of primary breast cancers, the overexpression of cyclin D1 is caused by gene amplification (16) . The molecular mechanism(s) for overexpression of cyclin D1 in the other tumors is unknown, although stabilization of cyclin D1 mRNA (17) and stabilization of cyclin D1 protein (18) have been suggested as possible mechanisms. Overexpression of cyclin D1 mRNA and protein is also observed in 50% or more of breast ductal carcinoma in situ lesions but more rarely in proliferative disease or atypical ductal hyperplasia, providing evidence that cyclin D1 overexpression plays a role during an early stage of tumor development (19) . Results obtained with animal model systems also suggest a role for cyclin D1 in malignant transformation of breast cancer cells (20 , 21) .

The PGs are a family of biologically active molecules having a diverse range of actions, depending on the PG type and cell target. Within this family, PGs of the A and J series, which contain a CP ring system, are potent inhibitors of cell proliferation in vitro and are able to suppress tumorigenicity in vivo (reviewed in Ref. 22 ). The antitumor activity of the CP PGs depends on the presence of an {alpha},ß-unsaturated carbonyl moiety within the CP ring, which reacts avidly with nucleophiles, such as sulfhydryls located on cysteine residues in cellular proteins and glutathione (22) . The CP PGs cause growth arrest in G1 or cell death, depending on the PG dose and characteristics of the target tumor cells (22, 23, 24) . The ability of the CP PGs to cause growth arrest or cell death in a variety of tumor cell lines has raised the possibility that they might be useful for the treatment of human cancer (24 , 25) .

The antineoplastic activity of the CP PGs is thought to be related to their ability to regulate the expression of a variety of stress-induced and cell cycle-related genes (22) . Genes exhibiting increased expression in response to the CP PGs include the genes encoding HSP70 (26) , c-fos (27) , Egr-1 (27) , gadd153 (28) , and the CDK inhibitor p21CIP1/WAF1 (29 , 30) . A number of genes exhibit decreased expression in response to the CP prostaglandins, including the genes that encode c-myc (31) , N-myc (32) , insulin-like growth factor I (30 , 33) , cyclin D1 (29 , 33, 34, 35) , and CDK4 (29) . Previous studies have demonstrated that the CP PG PGA2 causes growth arrest of MCF-7 breast cancer cells and C6 rat glioma cells in the G1 phase of the cell cycle (29 , 33) . In the MCF-7 cells, PGA2 causes a concerted repression of cyclin D1 and CDK4 and induction of the CDK inhibitor p21CIP1/WAF1, events that are causally related to arrest in G1 (29 , 33 , 34) .

Because of the role that cyclin D1 overexpression plays in malignant transformation in breast cancer and other tumors, cyclin D1 is a potential target for the rational design of new drugs to prevent or treat cancer. In the present study, we examined the molecular mechanism for repression of cyclin D1 protein expression by the CP PGs. The results indicate that these compounds cause a very rapid down-regulation of cyclin D1 protein, which precedes a decrease in cyclin D1 mRNA. The most active compound is 15d-PGJ2. The mechanism for rapid down-regulation of cyclin D1 protein is inhibition of cyclin D1 translation, caused at least in part by increased phosphorylation of eIF-2{alpha}.


    Results
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Effect of PGs on Cyclin D1 Protein Expression.
It has been shown previously that treatment of cells with the CP prostaglandin PGA2 results in a rapid decrease in cyclin D1 protein levels (29) . In our initial experiments, we compared the activity of PGA2 with the activity of two other compounds, the CP PG 15d-PGJ2 and the model compound CP. Dose-response experiments performed with cells treated for 1 h with the three agents demonstrated that 15d-PGJ2 was much more active in down-regulating cyclin D1 protein levels than either PGA2 or CP in MCF-7 cells (Fig. 1A)Citation . 15d-PGJ2 was active at the low dose of 3 µM and maximally effective at a dose of 10 µM. The other two compounds also down-regulated cyclin D1 protein but at considerably higher concentrations. Quantitative analysis of the Western blots using laser scanning densitometry yielded estimated IC50s of 5, 290, and 325 µM for 15d-PGJ2, PGA2, and CP, respectively. ß-Actin was used as a control in these experiments and was not affected by the treatment with these compounds.



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Fig. 1. A, dose-response relationships for regulation of cyclin D1 protein by 15d-PGJ2 (top), PGA2 (middle), and CP (bottom). *, location of electrophilic carbon atoms. B, effect of 15d-PGJ2 on cyclin D1 protein expression in HeLa and NIH-3T3 cells. Cultures were treated with DMSO vehicle or 10 µM 15d-PGJ2 for 2 h. Protein extracts were prepared and subjected to Western blotting (upper panel). Results were quantified by scanning densitometry (lower panel). Bars, means of three different cultures; ±, SE. C, dose-response for regulation of cyclin D1 protein by hydrogen peroxide (top, triplicate dishes treated with each concentration), sodium arsenite (middle, triplicate dishes treated with each concentration), and UV light (bottom). MCF-7 cells were treated for 1 h with each compound at the indicated concentrations or with UV as described in "Materials and Methods." Extracts were prepared, and the cyclin D1 and ß-actin protein levels were quantified by Western blotting.

 
15d-PGJ2 is a high-affinity ligand for the nuclear receptor PPAR{gamma} (reviewed in Ref. 22 ). Some biological effects of 15d-PGJ2 are mediated by binding to PPAR{gamma}, and some are independent of the receptor (22) . We have observed previously that MCF-7 breast cancer cells express PPAR{gamma} (36) . To test whether the effect of 15d-PGJ2 on cyclin D1 protein was mediated by PPAR{gamma}, we examined the effect on cyclin D1 protein levels of the thiazolidinedione compound rosiglitazone (BRL49653), a high-affinity PPAR{gamma} ligand that is structurally distinct from 15d-PGJ2 and that lacks the chemically reactive {alpha},ß-unsaturated carbonyl group (37) . Treatment of cells for 1 h with rosiglitazone at concentrations up to 10 µM did not have a significant effect on cyclin D1 protein levels (results not shown), indicating that this effect of 15d-PGJ2 may be independent of PPAR{gamma}. To investigate further the possible involvement of PPAR{gamma} in mediating the repression of cyclin D1 by 15d-PGJ2, we tested the effect of 15d-PGJ2 on cyclin D1 expression in HeLa cells and NIH-3T3 cells. Previous studies have demonstrated that neither cell line expresses detectable levels of PPAR{gamma} protein and that neither exhibits biological responses known to be mediated by PPAR{gamma} (36 , 38 , 39) . Treatment of the HeLa and NIH-3T3 cells with 15d-PGJ2 resulted in decreased expression of cyclin D1 (Fig. 1B)Citation . This result provides additional evidence demonstrating the existence of a PPAR{gamma}-independent pathway for regulation of cyclin D1 by 15d-PGJ2.

Several stress-inducing agents other than 15d-PGJ2 have been shown previously to decrease cyclin D1 protein levels in cultured cells (40 , 41) . We next tested the effect of hydrogen peroxide, sodium arsenite, and UV light on cyclin D1 protein levels in MCF-7 cells (Fig. 1C)Citation . Each of these agents decreased cyclin D1 protein levels but not to the degree nor at such a low dose as 15d-PGJ2. Therefore, all subsequent experiments were performed with 15d-PGJ2.

15d-PGJ2 Blocks Progression of Serum-stimulated Quiescent Cells through G1.
We next determined the effect of 15d-PGJ2 on cell cycle progression of MCF-7 cells (Fig. 2)Citation . Cells were arrested by culture in medium with low serum and then restimulated with fresh medium containing 10% serum with 15d-PGJ2 or DMSO vehicle. FACS analysis was performed to monitor cell cycle phase. Restimulation with medium containing 10% serum and vehicle resulted in synchronous entry into S-phase. Cells stimulated with medium containing 10% serum and 15d-PGJ2 remained arrested in G1 (Fig. 2)Citation .



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Fig. 2. Effect of 15d-PGJ2 on cell cycle reentry of MCF-7 cells. MCF-7 cells were incubated for 40 h in DMEM with 0.5% serum to synchronize cells in G0/G1. At T=0, the medium was changed to fresh medium containing 10% serum with vehicle or 15d-PGJ2 (final concentration, 20 µM). Control and treated cells were processed at various times and subjected to FACS analysis to determine cell cycle phase. A, FACS analysis of cells restimulated with 10% serum plus vehicle (upper) or 10% serum plus 15d-PGJ2 (lower). B, fraction of cells in S-phase at various time points. {bullet}, cells stimulated with 10% serum plus vehicle. {blacksquare}, cells stimulated with 10% serum plus 15d-PGJ2.

 
Time Course of 15d-PGJ2-mediated Reduction in Cyclin D1 Expression.
The time course for the down-regulation of cyclin D1 protein expression by 15d-PGJ2 was next determined by Western immunoblot analysis (Fig. 3)Citation . The effect of 15d-PGJ2 (10 µM) was apparent within 30 min after addition of 15d-PGJ2, with a maximal effect observed at 120 min. Partial recovery of cyclin D1 protein levels was observed at later time points. The partial recovery of cyclin D1 at the later time points may have been attributable to rapid metabolism and inactivation of 15d-PGJ2 mediated by glutathione-S-transferase (22) . In support of this idea, in additional experiments in which a second dose of 15d-PGJ2 (10 µM) was added at the 2-h time point, cyclin D1 protein expression did not recover from the repressed minimum observed at 2 h (data not shown).



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Fig. 3. Time course for the effect of 15d-PGJ2 on cyclin D1 expression. A, MCF-7 cells were treated with 10 µM 15d-PGJ2 for the indicated times, and cyclin D1 and ß-actin protein levels were determined by Western blotting. B, quantitative analysis of the cyclin D1 Western blot was performed using a laser densitometer. Error brackets, SE for three determinations (0 time point) or the range for two determinations (all other time points).

 
Effect of 15d-PGJ2 on Cyclin D1 mRNA Abundance.
Previous studies have demonstrated that PGA2 decreases the steady-state level of cyclin D1 mRNA as well as cyclin D1 protein (29 , 33 , 34) , although in one of these studies, the effect on protein levels appeared to precede the effect on mRNA levels (29) . Treatment of the MCF-7 cells with 15d-PGJ2 (10 µM) led to a time-dependent decrease in cyclin D1 mRNA abundance (Fig. 4A)Citation . This effect was first evident at 2 h, and cyclin D1 mRNA continued to decrease up to 8 h after 15d-PGJ2 addition (Fig. 4A)Citation . The abundance of GAPDH mRNA was unaffected by 15d-PGJ2, indicating that the effect of 15d-PGJ2 on cyclin D1 mRNA was specific. Importantly, no significant effect of 15d-PGJ2 on cyclin D1 mRNA was observed at 1 h after 15d-PGJ2 addition (Fig. 4B)Citation , a time point at which cyclin D1 protein was already decreased by >75% (Fig. 3)Citation . The more rapid effect of 15d-PGJ2 on cyclin D1 protein as compared with cyclin D1 mRNA indicated that 15d-PGJ2 decreased cyclin D1 protein at least partly at a step subsequent to cyclin D1 mRNA production/turnover.



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Fig. 4. Northern blots depicting the effect of 10 µM 15d-PGJ2 on cyclin D1 and GAPDH mRNA levels. A, effect on the quantity of cyclin D1 mRNA in cells treated for 2–8 h with 15d-PGJ2. B, effect in cells treated for 1 h. The major (4 kb) species of cyclin D1 mRNA is shown. C, vehicle-treated controls; T, treated with 10 µM 15d-PGJ2 for the time periods indicated.

 
15d-PGJ2 Alters the Polysome Distribution of Cyclin D1 mRNA.
The rapid disappearance of cyclin D1 protein after the addition of 15d-PGJ2 could be attributable either to inhibition of translation of cyclin D1 mRNA or accelerated degradation of cyclin D1 protein. To determine whether 15d-PGJ2 might decrease the translation of cyclin D1 mRNA, we examined the amount of this mRNA present in the polysomal (i.e., actively translated) fraction in the absence or presence of 15d-PGJ2 (Fig. 5A)Citation . In control cells, part of the cyclin D1 mRNA sedimented with the polysomal fraction and part sedimented with the untranslated fraction near the top of the gradient (Fig. 5ACitation , top panel). After treatment of cells with 15d-PGJ2 (10 µM, 1 h), the polysome-associated mRNA was completely shifted to the more slowly sedimenting region of the gradient, indicating that cyclin D1 translational initiation was severely inhibited (Fig. 5ACitation , lower panel). The results depicted in Fig. 5Citation were obtained 1 h after addition of 15d-PGJ2, at which time decreased size or abundance of cyclin D1 mRNA was not observed (Fig. 4B)Citation . Therefore, increased cyclin D1 mRNA degradation could not account for the results presented in Fig. 5ACitation . ß-Actin mRNA from 15d-PGJ2-treated cells was also shifted to a lighter fraction in the gradient, indicating that ß-actin translation was also inhibited (Fig. 5B)Citation . However, the shift of ß-actin mRNA was not as complete as with cyclin D1 mRNA; part of the ß-actin mRNA was shifted to an intermediate position in the gradient, consistent with the location of mRNA associated with a reduced number rather than completely depleted of ribosomes (Fig. 5BCitation , lower panel). The overall results indicate that 15d-PGJ2 inhibits the translation of cyclin D1 and ß-actin mRNA and are consistent with the possibility that the inhibitory effect of 15d-PGJ2 on ß-actin translation may be less severe than its inhibitory effect on cyclin D1 translation.



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Fig. 5. Polysomal profiles from control MCF-7 cells treated with DMSO vehicle (top) and treated with 10 µM 15d-PGJ2 (bottom). The A254 of each fraction from a sucrose gradient (fraction #1 representing the top of the gradient) was determined ({circ}), and isolated RNA was subject to Northern blot analysis to determine which fractions contained cyclin D1 (A) or ß-actin (B) mRNA ({bullet}). The position of free ribosomes was determined by ethidium bromide staining of the Northern gels. Fraction #3 within this region contained 40S subunits; fractions #4, 5, and 6 contained 60S subunits and 80S ribosomes, incompletely resolved from each other.

 
Cyclin D1 Protein Half-Life in MCF-7 Cells.
As the steady state level of any protein in the cell is equivalent to the amount synthesized minus the amount degraded, it was important to determine the half-life of cyclin D1 protein in MCF-7 cells under our experimental conditions. To do this, CHX was added to stop protein synthesis, and the time course for the disappearance of cyclin D1 was determined (Fig. 6 A)Citation . Cyclin D1 protein is degraded by the proteasome (12 , 42, 43, 44, 45) and calpain protease (46) , and it is known to turn over rapidly. Our results indicated that the half-life of cyclin D1 in MCF-7 cells was ~34 min, in concordance with previously published half-lives (42 , 47, 48, 49) . In contrast, the control protein ß-actin turned over much more slowly in the same cultures (Fig. 6A)Citation . (Linear regression analysis indicated that the slope of the ß-actin decay curve did not differ significantly from zero over the 60-min time course.) We next performed pulse-chase experiments to examine directly the possible effect of 15d-PGJ2 on cyclin D1 protein turnover. The results (Fig. 6B)Citation indicated that 15d-PGJ2 had no significant effect on cyclin D1 protein degradation. The half-life of cyclin D1 protein in control cells was 44.8 min and in 15d-PGJ2-treated cells was 45.2 min. These half-lives are slightly longer but in reasonable agreement with the half-life measured after CHX treatment. Taken together with the results presented above, this result provides evidence indicating that the rapid effect of 15d-PGJ2 on cyclin D1 protein expression is caused by inhibition of cyclin D1 translation rather than acceleration of cyclin D1 protein turnover.



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Fig. 6. A, time course showing effect of CHX on cyclin D1 ({bullet}) and ß-actin ({blacksquare}) protein levels. CHX (final concentration, 100 µg/ml) was added at T=0 to block protein synthesis, and levels of cyclin D1 and ß-actin protein were quantified by Western blotting. Each point represents the mean of three different cultures ± SE. The mean for the zero time point was set at 1.0. Regression analysis indicated that the slope of the cyclin D1 decay curve differed significantly from zero (P < 0.0001), whereas the slope of the ß-actin decay curve did not (P=0.0932). The half-life of cyclin D1 protein was also calculated by regression analysis. B, effect of 15d-PGJ2 on cyclin D1 protein degradation as determined by the pulse-chase method. Cells were pulse-labeled with [35S]methionine for 1 h, washed three times, and transferred to chase medium containing 2 mM unlabeled methionine at T=0 (see "Materials and Methods"). Cyclin D1 protein was immunoprecipitated at the indicated chase times and quantified by SDS gel electrophoresis and autoradiography. The level of labeled cyclin D1 protein at the beginning of the chase (T=0) was set at 1.0. Each data point represents the mean of results obtained in two different experiments. Circles and solid line, control cells; triangles and broken line, cells treated with 15d-PGJ2. The half-life of cyclin D1 protein was calculated by regression analysis.

 
15d-PGJ2 Generates a Predominantly Cytoplasmic Stress.
The results presented above suggested that the rapid down-regulation of cyclin D1 protein expression in cells treated with 15d-PGJ2 (Fig. 3)Citation could be accounted for by severe inhibition of cyclin D1 translation (Fig. 5A)Citation , followed by rapid turnover of cyclin D1 protein (Fig. 6)Citation . One clearly understood molecular mechanism for inhibition of translational initiation in eukaryotic cells is increased phosphorylation of eIF-2{alpha} on serine 51, which results in inhibition of the GTP exchange factor eIF-2B, and ultimately, decreased formation of ternary initiation complexes. A number of agents that induce stress are known to stimulate phosphorylation of eIF-2{alpha} and inhibit translation (50, 51, 52) . These can be roughly divided into two classes. Agents such as sodium arsenite induce stress in the cytoplasm and activate PKR, which phosphorylates eIF-2{alpha} (50 , 51) . Agents such as tunicamycin and thapsigargin induce stress in the ER and activate PERK, which also phosphorylates eIF-2{alpha} (52) . To determine whether 15d-PGJ2 at the concentration used in our experiments (10 µM) induced a stress response and whether this response originated in the cytoplasm or ER, we examined the effect of 15d-PGJ2 on expression of the genes for two molecular chaperones, HSP70 and GRP78. HSP70 is induced primarily by stress in the cytoplasm, whereas GRP78 is induced primarily by stress in the ER. The results (Fig. 7)Citation showed very strong induction by 15d-PGJ2 of HSP70 mRNA and relatively weak and delayed induction of GRP78 mRNA. These results are consistent with a strong cellular stress response to 10 µM 15d-PGJ2, with the stress emanating primarily from the cytoplasm rather than the ER.



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Fig. 7. A, Northern blot analysis showing the effect of 10 µM 15d-PGJ2 on HSP70, GRP78, and GAPDH mRNA levels. C, vehicle-treated controls; T, treated with 10 µM 15d-PGJ2 for the time periods indicated. B, results of experiments presented in A quantified by scanning densitometry. {square}, vehicle controls; {blacksquare}, treated with 15d-PGJ2. Error brackets, SE.

 
15d-PGJ2 Increases Phosphorylation of Translation Initiation Factor eIF-2{alpha}.
We next examined directly the effect of 15d-PGJ2 on eIF-2{alpha} phosphorylation. Sodium arsenite, which is known to activate PKR (50 , 51) and which also down-regulates cyclin D1 (Fig. 1C)Citation , was used as a positive control. The results (Fig. 8)Citation indicated that 15d-PGJ2 did in fact stimulate the phosphorylation of eIF-2{alpha} in MCF-7 cells. This effect was evident at 5 min after addition of 15d-PGJ2 and was maximal at 15 min. The overall results are thus consistent with increased phosphorylation of eIF-2{alpha}, with translational inhibition being the major mechanism for the rapid down-regulation of cyclin D1 by 15d-PGJ2.



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Fig. 8. A, effect of 15d-PGJ2 on phosphorylation of eIF-2{alpha} in MCF-7 cells. Cultures were treated with 10 µM 15d-PGJ2 (T), DMSO vehicle (C), or 100 µM sodium arsenite (A) for the indicated times. Protein extracts were prepared and subjected to Western blotting. Blots were probed with specific antibody for phospho-eIF-2{alpha} (top) and then reprobed with antibody for total eIF-2{alpha} (middle) and ß-actin (bottom). B, results of experiments presented in A quantified by scanning densitometry. Error brackets, range for duplicate determinations.

 
PKR Is Involved in Mediating the Effect of 15d-PGJ2 on eIF-2{alpha} Phosphorylation and Cyclin D1 Expression.
To investigate the possible role of PKR in mediating the induction of eIF-2{alpha} phosphorylation by 15d-PGJ2, we examined the effect of 15d-PGJ2 on eIF-2{alpha} phosphorylation in wild type (Pkr+/+) and PKR-null (Pkr0/0) mouse embryo fibroblast cell lines (Fig. 9)Citation . 15d-PGJ2 (10 µM) treatment significantly increased eIF-2{alpha} phosphorylation in wild-type fibroblasts above the level seen in the absence of the prostaglandin. The increase reached a maximum after 2 h of treatment of the wild-type fibroblasts. In contrast, very little effect was observed in the PKR-null mutant fibroblasts under similar conditions (Fig. 9)Citation . These results suggest PKR involvement in mediating the response to 15d-PGJ2.



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Fig. 9. Effect of 15d-PGJ2 treatment on the phosphorylation of eIF-2{alpha} in wild-type and PKR-null mouse embryo fibroblast cells. A, mouse embryo fibroblast cells, either wild type (Pkr+/+) or PKR-null (Pkr0/0), were treated for the indicated period of time with 10 µM 15d-PGJ2. Phosphorylated eIF-2{alpha} or {alpha}-tubulin protein levels were determined by Western immunoblot analysis. B, Western blots were quantified by scanning densitometry. Error brackets, SE for three independent experiments. {square}, control wild-type cells; {blacksquare}, wild-type cells treated with 15d-PGJ2; {circ}, control PKR-null cells; {bullet}, PKR-null cells treated with 15d-PGJ2.

 
We next tested the effect of 15d-PGJ2 on cyclin D1 protein levels in the wild-type and PKR-null mouse fibroblast cells (Fig. 10)Citation . 15d-PGJ2 treatment significantly reduced cyclin D1 protein levels in the wild-type fibroblasts (Fig. 10)Citation . In three different experiments, the mean decrease in cyclin D1 protein expression was 55% after 2 h and 56% after 3 h treatment of wild-type Pkr+/+ cells with 15d-PGJ2, as compared with vehicle-treated cells. By contrast, under similar conditions, 15d-PGJ2 treatment had relatively little effect on the cyclin D1 protein levels in PKR-null fibroblasts. In these cells, cyclin D1 was decreased by 25% after 2 h and by 15% after 3 h in cells treated with 15d-PGJ2, as compared with vehicle-treated cells. The greater response of cyclin D1 to 15d-PGJ2 in wild-type cells as compared with PKR-null cells was consistently observed in all three experiments. These results, taken together with those presented in Fig. 9Citation , provide evidence that PKR is involved in mediating the effect of 15d-PGJ2 on eIF-2{alpha} phosphorylation and cyclin D1 expression.



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Fig. 10. Time course showing the effect of 15d-PGJ2 treatment on cyclin D1 protein expression in wild-type and PKR-null mouse embryo fibroblast cells. Wild-type (Pkr+/+) or PKR-null (Pkr0/0) cells were treated for the indicated period of time with 20 µM 15d-PGJ2. Cyclin D1 protein levels were determined by Western immunoblot analysis.

 

    Discussion
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Overexpression of cyclin D1 has been implicated in the etiology of a number of types of human cancer including breast cancer (14, 15, 16, 17, 18, 19, 20) . Therefore, cyclin D1 is a potential target for the rational design of new drugs to prevent or treat cancer. The present study confirms and extends previous results demonstrating that PGA2 down-regulates cyclin D1 in tumor cells (29 , 33) . We report here the novel result that 15d-PGJ2 is ~60 times more potent than PGA2 in eliciting this effect. 15d-PGJ2 is a high-affinity ligand for nuclear receptor PPAR{gamma}, and MCF-7 cells have been shown previously to express PPAR{gamma} (22 , 36) . However, the rapid effect of 15d-PGJ2 on cyclin D1 does not appear to be mediated by PPAR{gamma}. In particular: (a) PPAR{gamma} is a regulator of transcription, whereas the effects that we have observed are posttranscriptional; (b) another high-affinity PPAR{gamma} ligand, rosiglitazone, had no effect on cyclin D1; (c) 15d-PGJ2 down-regulated cyclin D1 expression in HeLa and NIH-3T3 cells, which do not express detectable levels of PPAR{gamma}; and (d) CP, which is not a PPAR{gamma} ligand, was active.

Most, if not all, of the PPAR{gamma}-independent biological actions of the CP PGs depend on the presence of the chemically reactive {alpha},ß-unsaturated carbonyl within the CP ring (22 , 30 , 36) . 15d-PGJ2 has been observed previously to be more active than PGA2 with other biological endpoints including repression of nuclear factor-{kappa}B activation (36) . 15d-PGJ2 differs from PGA2 in having two chemically reactive centers rather than one (Fig. 1A)Citation . Thus, it is possible that 15d-PGJ2 could act as a cross-linking agent. Alternatively, it is possible that the higher bioactivity of 15d-PGJ2 is related to higher affinity for some yet-to-be-identified protein target within the cell.

Similar to earlier results obtained with PGA2, we found that 15d-PGJ2 negatively regulates cyclin D1 mRNA levels, and this effect presumably contributes to the down-regulation of cyclin D1 protein expression at times >=2 h after compound addition. The repressive effect of the CP PGs on cyclin D1 mRNA expression is reported to result from transcriptional repression (35) and/or destabilization of cyclin D1 mRNA (34) . However, consistent with an earlier report (29) , we found that 15d-PGJ2 decreased the level of cyclin D1 protein before it decreased the level of cyclin D1 mRNA. The discordant time courses for the changes in cyclin D1 protein and mRNA indicated the existence of an additional mode of regulation at a step subsequent to mRNA production/turnover. Two formal possibilities for this regulation were translational repression or accelerated protein turnover. Our results indicate that translational repression is one mechanism for the regulation of cyclin D1 by 15d-PGJ2. Thus, the effect of 15d-PGJ2 on cyclin D1 resembles the effects of clotrimazole and tunicamycin, which also appear to regulate cyclin D1 translation (40 , 49 , 53) . In contrast, retinoic acid and osmotic shock have been reported to stimulate cyclin D1 protein turnover (43, 44, 45) .

One of the most clearly understood mechanisms for regulation of translation in eukaryotic cells is increased phosphorylation of eIF-2{alpha} (54) . Our results suggest that this modification of eIF-2{alpha} is likely a principal mechanism for translational regulation by 15d-PGJ2. The differential regulation of cyclin D1 as compared with ß-actin protein abundance (Fig. 1)Citation is readily explainable by the more rapid turnover of cyclin D1 protein as compared with ß-actin after inhibition of translation. In particular, the time course for the decrease of cyclin D1 protein in cells treated with 15d-PGJ2 (Fig. 3)Citation is very similar to the time course for its decrease after treatment of cells with the known translation inhibitor, CHX (Fig. 6A)Citation . In contrast, ß-actin turns over very slowly after inhibition of protein synthesis (Fig. 6A)Citation . The polysome profile results also leave open the possibility that cyclin D1 translation may be more severely repressed than ß-actin translation, and that this may also contribute to the observed differential regulation of the steady-state levels of the two proteins. Differential regulation of the translation of reoviral mRNA transcripts by PKR has been described previously (55) .

Although induction of HSP70 and GRP78 by CP PGs has been reported previously (26 , 56) , to our knowledge this is the first study in which both endpoints have been measured with the same compound in the same experiment. Treatment of MCF-7 cells with 15d-PGJ2 resulted in a rapid strong induction of HSP70 gene expression. This effect was clearly apparent at 2 h and was maximal at 4 h. In contrast, the effect on GRP78 was weak and delayed compared with the effect on HSP70. Thus, 15d-PGJ2 induces a stress response, and this response emanates primarily from the cytoplasm rather than the ER.

In considering various hypotheses to explain the repressive effect of 15d-PGJ2 on cyclin D1 translation, we noted that stress-inducing agents such as sodium arsenite have been shown previously to induce phosphorylation of eIF-2{alpha} (Refs. 50 , 51 ; Fig. 8Citation ). Treatment of MCF-7 cells with sodium arsenite also results in decreased levels of cyclin D1 protein (Fig. 1C)Citation . Sodium arsenite has been shown recently to activate PKR (51) ; thus, PKR was a likely candidate enzyme that phosphorylated eIF-2{alpha} in response to 15d-PGJ2. Our findings obtained with the Pkr0/0 fibroblasts, which had an attenuated response to 15d-PGJ2 as compared with wild-type fibroblasts, are consistent with the notion that PKR plays a role in the increase in eIF-2{alpha} phosphorylation and repression of cyclin D1 caused by 15d-PGJ2. An alternative pathway(s) responsible for the weak residual response of the Pkr0/0 cells to 15d-PGJ2 remains to be elucidated. Interestingly, the time course for the increase in eIF-2{alpha} phosphorylation in response to 15d-PGJ2 appeared to be more gradual in mouse embryo fibroblastic cells as compared with the human MCF-7 cells. MCF-7 cells have been reported to express high levels of PKR (57) , and it is possible that this accounts for the more rapid response observed in these cells. Alternatively, the response in the two cell types may be modulated by other factors, such as differences in ability to metabolize 15d-PGJ2 (36) or differences in activities of cellular proteins that modulate PKR function such as PACT or P58, a known regulator of HSP70 (58) .

Tunicamycin, which induces stress in the ER and initiates the unfolded protein response, has been reported previously to repress cyclin D1 translation (49) . This agent is known to activate another eIF-2{alpha} kinase, PERK (52) , located in the ER. In analogy with the results presented here, it is possible that the mechanism for inhibition of cyclin D1 translation by tunicamycin is activation of PERK, followed by phosphorylation of eIF-2{alpha} (49) .

Previous studies of the antineoplastic activity of the PGJ series prostaglandins have focused primarily on {Delta}12-PGJ2 (22 , 59) , the precursor of 15d-PGJ2. McClay et al. (59) have reported synergistic cytotoxic interaction between {Delta}12-PGJ2 and both ionizing radiation and cisplatin. This synergy was observed in a variety of tumor cell lines and resulted in a decrease of ~10-fold in the dose of cisplatin or ionizing radiation needed to kill 50% of the tumor cells. It will be of interest to determine whether 15d-PGJ2 also synergizes with other antitumor agents. A limiting factor for usefulness of most anticancer drugs is toxicity in vivo. A recent study has demonstrated that 15d-PGJ2 (1 mg/kg/day i.p.) is well tolerated in rats (60) . At this dose, anti-inflammatory activity of 15d-PGJ2 was observed in adjuvant-induced arthritis, with no toxic side effects (60) . The low toxicity of 15d-PGJ2 in this study provides a rationale for future testing of the antitumor activity of this compound in vivo.


    Materials and Methods
 TOP
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Materials.
Prostaglandins were obtained from Cayman Chemical Co. and were supplied in methyl acetate. Before using PGA2 and 15d-PGJ2 in experiments, the methyl acetate was evaporated under a stream of argon, and the compounds were redissolved in argon-purged DMSO. 2-Cyclopenten-1-one was purchase from Aldrich Chemical Co. and diluted into DMSO before use. Cycloheximide was purchased from Sigma Chemical Co., and stock solutions were prepared in ethanol at a concentration of 10 mg/ml.

DNA clones were generously provided by the following people: full-length human cyclin D1 cDNA clone (Ref. 61 ; David Beach, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY); hamster GRP78, human HSP70, and GAPDH cDNA clones (Refs. 62 , 63 ; Amy Lee, University of Southern California School of Medicine, Los Angeles, CA); and human ß-actin (Ref. 64 ; Larry Kedes, University of Southern California School of Medicine).

Cell Culture.
MCF-7 cells were obtained from the American Type Culture Collection and maintained as monolayer cultures in DMEM (Cellgro) supplemented with 10% fortified BCS (Cosmic calf serum; HyClone) plus penicillin (100 units/ml) and streptomycin (100 µg/ml). Permanent cell lines derived from embryo fibroblasts of Pkr+/+ and Pkr0/0 mice (65) were generously provided by A. E. Koromilas (Lady Davis Institute, Jewish General Hospital, Montreal, Canada) and cultured in DMEM plus 10% fetal bovine serum and antibiotics.

Western Blot Analysis.
Cells were plated at a density of 700,000 cells/6-cm dish and cultured for 3 days at 37°C. At the beginning of each experiment, cell cultures were washed once with PBS and incubated for 1 h in 0.5% BCS medium. 15d-PGJ2 or other chemical agents were then added to the cultures, and cultures were incubated for an additional 60 min unless otherwise indicated. For treatment with UV, cells were cultured using the same procedure as for treatment with chemicals. The medium was then aspirated, and the cultures were irradiated with UV at the dose indicated, using a Stratalinker UV cross-linking apparatus (Stratagene). Prewarmed (37°C) medium (DMEM with 0.5% BCS and antibiotics) was then added to each culture, and incubation was continued for 60 min at 37°C.

For preparation of extracts for Western blot analysis, cell cultures were transferred to ice, washed once with PBS, and scraped into 1 ml of ice-cold PBS. Cells were then pelleted in a microcentrifuge and suspended in NETN extraction buffer plus protease inhibitors [50 mM Tris (pH 7.6), 0.15 M NaCl, 5 mM EDTA, 0.5% NP40, with 1 µl of 0.4 M benzamidine, 1 µl of 10 mg/ml leupeptin, 1 µl of 10 mg/ml aprotinin, and 1 µl of 250 mM phenylmethylsulfonyl fluoride for every 300 µl of NETN buffer]. For Western blot analysis of eIF-2{alpha} phosphorylation, the following phosphatase inhibitors were also included in the NETN buffer: NaF (20 mM), ß-glycerophosphate (20 mM), and sodium PPi (12 mM). Extracts were clarified by centrifugation in a microcentrifuge, and protein concentration was determined using the Folin/Lowry method (66) . Protein extracts (50 µg of protein) were size fractionated by 10% SDS-PAGE and electroblotted onto nitrocellulose membranes (NitroBind MSI, Westborough, MA), using standard techniques. For detection of cyclin D1, a rabbit polyclonal antibody (Santa Cruz Biotechnology Inc.; SC-718) was used at a 1:400 dilution. For detection of ß-actin, a rabbit affinity-isolated, antigen-specific antibody (Sigma-Aldrich Co.; A-2066) was used at a 1:800 dilution. For detection of total eIF-2{alpha}, a rabbit polyclonal antibody (Santa Cruz; SC-11386) was used at a 1:400 dilution. For detection of phosphorylated eIF-2{alpha}, an antibody specific for phospho-Ser51-eIF-2{alpha} (Biosource International) was used at a 1:1000 dilution (final concentration, 0.5 µg/ml). Antibody directed against {alpha}-tubulin was provided by L. Wilson (University of California, Santa Barbara, CA). The secondary antibody was a peroxidase-labeled, antirabbit IgG antibody, affinity purified made in goat (Vector Laboratories, Inc. or Amersham) at a dilution of 1:10,000. Proteins were then detected with the enhanced chemiluminescence system (SuperSignal; Pierce, Rockford, IL).

Polysomal Profiles.
Polysomal profiles were obtained as described previously (67) , with slight modifications. For preparation of cytoplasmic extracts, MCF-7 cells (three 15-cm cell culture plates) were treated with 15d-PGJ2 (10 µM) or vehicle for 1 h before extraction. The plates were then treated with CHX (100 µg/ml) for 5 min at 37°C, washed with PBS containing CHX (100 µg/ml), and scraped into PBS + CHX. Cells were pelleted by centrifugation, swollen for 2 min in low-salt buffer [LSB; 20 mM Tris (pH 7.5), 10 mM NaCl, 3 mM MgCl2, 1 mM DTT, 50 units of RNAsin (Promega)], and then lysed by the addition of lysis buffer (0.2 M sucrose, 0.1% Triton X-100, in LSB), followed by 10 strokes with a Dounce homogenizer using piston A. The nuclei were pelleted by centrifugation in a microcentrifuge at 15,000 rpm for 30 s. The supernatant corresponding to the cytoplasmic extract was poured into a new centrifuge tube containing 10% heparin (Sigma; 10 mg/ml), 3% 5 M NaCl, and 1 mM DTT. Equal A260 units of cytoplasmic extracts (~80 µg of RNA) from vehicle or 15d-PGJ2-treated cells were applied to a 0.5–1.5 M sucrose gradient (in LSB) layered over a 2 M sucrose cushion, and centrifuged at 36,000 rpm in a Beckman SW41 swinging bucket rotor for 220 min at 4°C. Gradients were fractionated from the top into 1:10 volume of 10% SDS and proteinase K. The absorbance of each fraction at 254 nm was monitored. The RNA from each fraction was extracted with an equal volume of phenol/chloroform, precipitated with isopropanol, and analyzed by Northern blotting to quantify cyclin D1 mRNA.

Determination of Cyclin D1 Protein Half-Life by Pulse-Chase Method.
Cells were metabolically labeled with [35S]methionine for 1 h in pulse medium, methionine, and cystine-free DMEM supplemented with 0.5% BCS, antibiotics, 0.02 mM unlabeled methionine, and 80 µCi/ml Trans35S-label (ICN). Cells were treated with either DMSO vehicle (control) or 10 µM 15d-PGJ2 during the pulse period. After the 1-h labeling period, cultures were flooded with an excess volume of chase medium (DMEM, containing 2 mM unlabeled methionine and supplemented with antibiotics plus 0.5% BCS). The medium was aspirated, and cells were washed three times with chase medium. Prewarmed chase medium was then added to the cells, and incubation was continued at 37°C. For cells treated with 15d-PGJ2, this compound was also present during the chase. Cultures were harvested, and protein extracts were prepared at 0, 20, 40, and 60 min, according to the methods described above for Western blotting. The protein concentration of each extract was determined by the Bradford method using the Bio-Rad protein assay reagent (Bio-Rad, Hercules, CA).

Cyclin D1 antibody was bound to protein A-Sepharose beads by overnight incubation at 4°C. Beads were washed twice with NETN. Aliquots of protein extract (1 mg of protein) were incubated with antibody-conjugated beads or unconjugated beads (negative control) for 2.5 h at 4°C. The beads were washed four times with NETN and once with PBS. Labeled cyclin D1 bound to the beads was removed by boiling the beads in sample buffer and subjected to SDS-PAGE in 10% gels. The gels were fixed in 45% methanol and 10% acetic acid for 20 min and then submerged in 1 M sodium salicylate for 30 min. The gels were washed three times, dried, and subjected to autoradiography. The band corresponding to cyclin D1 was quantified by scanning densitometry.

Northern Blot Analysis.
For Northern blot analysis, total cellular RNA was extracted as described previously (30 , 33) . RNA (15-µg aliquots) was denatured and electrophoresed in 1% agarose gels containing 2.2 M formaldehyde. The RNA was then transferred to nylon filters as described previously (30 , 33) . The integrity of the 18S and 28S bands of the extracted RNA indicated that the RNA had not degraded during the extraction procedure. Even loading of the gels was confirmed by ethidium bromide staining. The DNA probes (gel-purified restriction fragments of cDNA clones) were labeled by random priming with [{alpha}-32P]dCTP (30 , 33) . Filters were prehybridized, hybridized, and washed as described previously (30 , 33) . Results were quantified by scanning autoradiograms with an LKB UltroScan laser densitometer, exercising caution to stay within the linear range of the film (30 , 33 , 34 , 37) .

Flow Cytometry Analysis.
FACS analysis was performed as published previously (33) . Briefly, 8 x 105 cells were plated in 10-cm dishes and grown to 60% confluency in DMEM plus 10% BCS. The culture medium was then aspirated, cells were washed once with PBS, fresh medium with 0.5% BCS was added, and incubation was continued for 40 h at 37°C to synchronize the cells in the G0/G1 phase. Synchronized cells were stimulated to progress through the cell cycle by changing to fresh medium with 10% BCS with vehicle or 15d-PGJ2 (final concentration, 20 µM). Control and treated groups were processed at the time points indicated after the initiation of cell cycle. Cells were trypsinized and resuspended in ice-cold PBS at a density of 2 x 106 cells/ml. Cells (1 ml of suspension) were fixed by adding 2 ml of methanol dropwise with constant mixing and then incubated on ice for 30 min. Cells were then pelleted by centrifugation and resuspended in 500 µl of stain solution (10 mg of propidium iodide, 0.1 ml of Triton X-100, and 3.7 mg of EDTA dissolved in 100 ml in PBS) and 500 µl of heat-treated RNase-A solution (2 mg/ml in PBS) for 30 min at room temperature in the dark. Stained cells were analyzed using a Becton Dickinson Immunocytometry System for the relative content based on red fluorescence levels, and the distribution of cells in different phases of the cell cycle was calculated using CellFIT software (Becton Dickinson).


    Acknowledgments
 
We thank A. Lee, D. Beach, and L. Kedes for providing various plasmids, B. Walter for help with the FACS analysis, and A. Koromilas for providing the wild-type and PKR-null fibroblasts.


    Footnotes
 
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.

1 This research was supported by Department of Defense Breast Cancer Research Program Grant DAMD17-99-1-9102 (to D. S. S.) and NIH Grant AI20611 (to C. E. S.). Back

2 To whom requests for reprints should be addressed, at Biomedical Sciences Division, University of California, Riverside, CA 92521-0121. Phone: (909) 787-5612; Fax: (909) 787-5504; E-mail: daniel.straus{at}ucr.edu Back

3 The abbreviations used are: CDK, cyclin-dependent protein kinase; 15d-PGJ2, 15-deoxy-{Delta}12,14-PGJ2; PG, prostaglandin; CP, cyclopentenone; PPAR, peroxisome proliferator-activated receptor; CHX, cycloheximide; eIF-2{alpha}, eukaryotic initiation factor 2{alpha}; PKR, protein kinase R; FACS, fluorescence-activated cell sorter; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; PERK, PKR-like ER kinase; ER, endoplasmic reticulum; HSP, heat shock protein; GRP, glucose-regulated protein; BCS, bovine calf serum. Back

Received for publication 12/14/01. Revision received 6/17/02. Accepted for publication 6/24/02.


    References
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 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 

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