Control by Basal Phosphorylation of Cell Cycle-Dependent, Hormone-Induced Glucocorticoid Receptor Hyperphosphorylation

Jiong-Ming Hu, Jack E. Bodwell and Allan Munck

Department of Physiology Dartmouth Medical School Lebanon, New Hampshire 03756-0001


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS AND DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Mouse glucocorticoid receptors (GRs) are phosphorylated in the N-terminal domain at serine/threonine residues, most lying in consensus sequences for cell cycle-associated kinases. Glucocorticoid agonists, but not antagonists, induce hyperphosphorylation. Phosphorylation of GRs overexpressed in Chinese hamster ovary (CHO) cells is cell cycle-dependent: basal phosphorylation in S phase is one third that in G2/M; glucocorticoids induce hyperphosphorylation in S but not G2/M, paralleling the reported sensitivity in S and resistance in G2/M of proliferating cells to transcriptional activation by glucocorticoids. This parallel led us to investigate what controls hyperphosphorylation. We tested three hypotheses: hyperphosphorylation is controlled by 1) negative charge due to basal GR phosphorylation, being permitted in S by low charge and blocked in G2/M by high charge; 2) presence in S and absence in G2/M of required kinases; 3) availability in S and lack in G2/M of unoccupied phosphorylatable sites. Our results are inconsistent with 2) and 3), but strongly support 1). GR mutants with alanines (A7GR) or glutamates (E7GR) replacing all but one phosphorylated site were overexpressed in CHO cells. Serine 122 remained intact to report GR phosphorylation. Consistent with hypothesis 1, with A7GRs hormone-induced hyperphosphorylation occurred in both S and G2/M (thus revealing kinase activity for hyperphosphorylation of at least serine 122 in both phases), whereas with E7GRs it occurred in neither phase. We conclude that basal GR phosphorylation controls hormone-induced GR hyperphosphorylation by modulating negative charge in the N-terminal domain and could potentially control other cell cycle-dependent GR properties.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS AND DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Steroid hormone receptors are all phosphorylated and undergo hormone-dependent hyperphosphorylation (reviewed in Ref.1). Mouse glucocorticoid receptors (GRs)1 are phosphorylated in the N-terminal domain at serines and one threonine, most within transactivation regions (2). GR hyperphosphorylation is induced by glucocorticoid agonists but not by the antagonist RU486 (3, 4, 5, 6). It occurs within 5–10 min of hormone addition (3, 7), follows receptor activation (7), and yields no new phosphorylated sites (8). The rapidity of hyperphosphorylation compared with the t1/2 for GR dephosphorylation of 90–120 min (7) indicates that hyperphosphorylation is due to accelerated phosphorylation rather than slowed dephosphorylation.

To date, reported effects of steroid hormone receptor phosphorylation on activity are subtle and varied, serving modulatory rather than obligatory functions. For example, phosphorylation of serine 118 in the human estrogen receptor (9) modulates enhancement of transcriptional activation by growth factors (10); mutation of this serine to alanine decreases estrogen-induced transactivation by 15 to 75%, depending on cell type and reporter gene (11, 12). In the chicken progesterone receptor, mutation of serine 530 to alanine reduces transcriptional activity by 70–85% at low, but not at high, hormone concentrations (13). Mutation of serine 211 to alanine reduces transcriptional activity by 25 to 80%, again depending on cell type and reporter gene (14). Comparable effects are seen with vitamin D (15) receptors. Mouse GRs mutated at single and multiple phosphorylated residues have been reported to give hormone-induced activity almost indistinguishable from that of wild type GRs (WTGRs) when transiently transfected into COS cells with a reporter gene under a mouse mammary tumor virus promoter (16). However, when in similar experiments a minimal promoter with simple glucocorticoid response elements is used, most GRs with one or more phosphorylated sites mutated to alanine are 50–75% less active than WTGRs (J. A. Cidlowski, personal communication). These and related GR phosphorylation mutants behave like WTGRs with respect to subcellular localization (17) and superactivation of transcription by cAMP (18) but are much less sensitive to hormone-induced down-regulation (19).

A largely unexplored role for GR phosphorylation is regulation of hormone-induced GR activity through the cell cycle. Synchronized cells with endogenous WTGRs generally are sensitive to glucocorticoids in late G1 and S, but resistant in G2, M, and early G1. Effects measured include induction of alkaline phosphatase activity (20) and epidermal growth factor receptors (21) in HeLa cells, of tyrosine aminotransferase in hepatoma cells (22), and of the metallothionein-I gene in L cell fibroblasts (23). This cell cycle dependence has some specificity for glucocorticoid activity because with cells arrested in G2, transcription of the endogenous metallothionein-I gene is resistant to glucocorticoid induction but remains sensitive to induction by heavy metals (23). In GrH2 cells glucocorticoid resistance in G2 is selective, affecting transactivation from a simple glucocorticoid response element but not repression from a composite glucocorticoid response element (24).

Much evidence links glucocorticoid activity through the cell cycle to GR phosphorylation. Cidlowski and colleagues found that GRs in HeLa cells are more negatively charged in G2 and early G1 than in late G1 and S (21, 25) and proposed that increased GR phosphorylation or glycosylation causes glucocorticoid resistance in G2 (26). Hsu et al. (23) detected alterations in two-dimensional phosphopeptide maps of GRs from L cells arrested in G2, and advanced a similar proposal. Most phosphorylated residues in mouse GRs lie in consensus sequences for cell cycle-associated kinases (2, 27). Furthermore, basal GR phosphorylation is 3-fold higher in G2/M than in S (reflecting a 3-fold higher frequency of phosphorylation of the same set of sites, since no new phosphorylated sites appear in G2/M) (27). Most significantly, hormone-induced hyperphosphorylation nearly doubles GR phosphorylation in S (a glucocorticoid-sensitive phase) but is absent in G2/M (glucocorticoid-resistant phases) (27).

This parallel between cell cycle dependence of hormone-induced GR hyperphosphorylation and of glucocorticoid activity prompted us to determine why GR hyperphosphorylation can occur in S but is blocked in G2/M (27). We have tested the following three hypotheses: hormone-induced hyperphosphorylation is controlled 1) by overall negative charge from basal phosphorylation, being permitted by the relatively low charge in S and blocked by the high charge in G2/M; 2) by the presence in S and absence in G2/M of required kinases; 3) by the presence in S and absence in G2/M of phosphorylatable sites, which are unfilled in S but filled by basal phosphorylation in G2/M. Our results strongly favor the first hypothesis and are inconsistent with the other two.


    RESULTS AND DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS AND DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
To test the first hypothesis — that hormone-induced hyperphosphorylation of GRs is permitted by low negative charge and blocked by high negative charge in the N-terminal domain — we have conducted experiments with two mutant GRs overexpressed in CHO cells. The A7GR mutant has alanines replacing all but one phosphorylated amino acid of the mouse GR, thus limiting the negative charge in the N-terminal domain almost to the level of unphosphorylated GRs. The E7GR mutant has glutamates replacing those same amino acids, raising the overall negative charge in the N-terminal domain to a level comparable to that of wild type GRs phosphorylated at more than half the sites. A single phosphorylated site, serine 122, was left intact in both mutant GRs to serve as reporter of 32P labeling. In the wild type mouse GR, serine 122 is a minor phosphorylated site (2) that undergoes an average degree of hormone-induced hyperphosphorylation in unsynchronized WCL2 cells (8).

Cells stably transfected with the mutant GRs were synchronized in S and G2/M (27). Hormone-induced hyperphosphorylation within each phase was measured by comparing 32P/35S ratios of GRs, purified by immunoprecipitation and SDS-PAGE, from cells that had been incubated with [35S]methionine and [32P]orthophosphoric acid, and during the last hour of labeling were treated with 500 nM triamcinolone acetonide (TA) or were untreated (controls). Incorporation of 35S served to normalize 32P incorporation, with and without TA treatment, to the same amount of GR protein (3, 7, 27).

Figure 1Go shows that A7-GRs, in contrast to WTGRs, become hyperphosphorylated in both S and G2/M. This is the outcome predicted by the first hypothesis, because with A7-GRs the negative charge in the N-terminal domain is limited throughout the cell cycle by an almost complete lack of phosphorylatable sites and therefore cannot be raised through basal phosphorylation to levels postulated to block hyperphosphorylation of WTGRs in G2/M.



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Figure 1. Hormone-Induced Hyperphosphorylation of A7GR Mutant Glucocorticoid Receptors in Cells Synchronized in S Phase and G2/M

Cells with stably transfected A7GRs were synchronized and then metabolically labeled with [32P]orthophosphate and [35S]methionine for 4 h. During the last hour of labeling, they were treated with 500 nM TA, or not treated (controls). The A7GRs were immunopurified, separated on SDS-PAGE, and analyzed for 32P and 35S incorporation as described in Materials and Methods. 32P incorporation was normalized to 35S incorporation, which within each phase of the cell cycle is proportional to GR protein (7). Bars represent means and ranges from two experiments. The high values for S and G2/M are from one experiment, and the low values are from the other experiment. Controls are set at 100%.

 
An additional conclusion from the results in Fig. 1Go is that the kinase necessary for hyperphosphorylation of serine 122 is present and active in both S and G2/M, because hyperphosphorylation occurred in both these phases. Thus, at least with respect to serine 122, we can rule out the second hypothesis, namely, that absence of hormone-induced hyperphosphorylation in G2/M is due to absence of the required kinase activity.

The conclusions drawn from the results in Fig. 1Go assume that no major new sites, i.e. sites that are not present in WTGRs, are phosphorylated in the A7GRs. The HPLC phosphopeptide maps in Fig. 2Go validate this assumption by establishing that no significant phosphorylated sites other than those present in WTGRs appear in A7GRs, either before or after hormone treatment. The top map, of WTGRs from unsynchronized cells, is included for comparison. It was obtained by our current procedures (8, 28), which differ slightly from those used originally (2). Locations are given of previously identified phosphopeptides (2) and of a peak we have recently identified tentatively with a tryptic peptide containing serine 412.



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Figure 2. HPLC Phosphopeptide Maps of WTGR from Unsynchronized Cells, of A7GRs from TA-Treated or Untreated Cells Synchronized in S or G2/M, and of WTGRs from Cells Synchronized in G2/M

The upper and lower maps of WTGRs are included for comparison. The lower one is from a previous study (27). Both maps were obtained with GRs purified from WCL2 cells metabolically labeled with [32P]orthophosphate for 4 h (27). The middle four maps were obtained with A7GRs from synchronized cells that, as for the experiments described in Figs. 1Go and 3Go, were labeled with [32P]orthophosphate for 4 h, and treated or not during the last hour with 500 nM TA. Labeled A7GRs were immunopurified and trypsinized, and HPLC phosphopeptide mapping analyses were performed as described in Materials and Methods. Vertical scales of 32P incorporation differ between maps: maximum incorporation per fraction for the top map was about 300 cpm, for the middle four maps about 50 cpm, and for the bottom map about 3000 cpm.

 
The next four maps in Fig. 2Go are of A7GRs purified from cells synchronized in S and G2/M, untreated and treated with TA. Background noise is relatively high because the phosphorylation level of serine 122 is low (2, 8). As expected, the only previously identified peak in these maps, around fraction 180, corresponds in position to the peptide in WTGRs (top map) containing serine 122 (2). The peak or peaks around fractions 50–60 match one in the WTGR that is still unidentified. In the original HPLC maps (2) it was a minor peptide that was not sequenced. Since then, with different procedures (8, 28) and preparations of trypsin, it has become more prominent, as can be seen in the top map (WTGR). From the present results it could represent another peptide containing serine 122, differing from that near fraction 180 in completeness of trypsin digestion. Whether that assignment is correct or not, however, affects none of our conclusions. Some peaks that are slightly above background noise level appear in the maps of the TA-treated A7GR; they account for insignificant amounts of 32P labeling.

Figure 3Go shows that with E7GRs no hyperphosphorylation occurs in either S or G2/M. This, again, is the outcome predicted by the first hypothesis, since throughout the cell cycle the glutamate residues in the E7GR contribute a high, fixed negative charge in the N-terminal domain, which cannot be lowered to a level comparable to that postulated to permit hyperphosphorylation of wild type GRs in S phase. These results also provide evidence against the second hypothesis, because it fails to predict absence of hyperphosphorylation in S.



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Figure 3. Hormone-Induced Hyperphosphorylation of E7GR Mutant Glucocorticoid Receptors in Cells Synchronized in S Phase and G2/M Cells with stably transfected E7GRs were treated, and the results analyzed, as described in the legend for Fig. 1Go. Bars represent means and ranges from two experiments. The high value for S and low value for G2/M are from one experiment, and the low value for S and high value for G2/M are from the other experiment. Controls are set at 100%.

 
Two independent lines of evidence argue against the third hypothesis — that wild type GRs are not hyperphosphorylated in G2/M because all sites are already fully phosphorylated under basal conditions. First, without ad hoc assumptions this hypothesis would not explain either the occurrence of hyperphosphorylation of A7GRs in G2/M (Fig. 1Go), or the lack of hyperphosphorylation of E7GRs in S (Fig. 3Go).

Second, the HPLC phosphopeptide map of WTGRs from cells synchronized in G2/M (bottom map in Fig. 2Go) is practically indistinguishable from the maps of WTGRs from unsynchronized cells (top map in Fig. 2Go) and of WTGRs from cells synchronized in S (27). It gives no indication that in G2/M, all sites on the WTGRs are fully phosphorylated. If they were fully phosphorylated, they would be equally labeled after reaching steady state with 32P. Rates of GR phosphorylation and dephosphorylation (3, 7, 8, 28) indicate that the 4-h incubations with 32P used here will label GRs to 70–80% of steady state, sufficient to give HPLC-labeling patterns closely approximating those at steady state. The sum of 32P in peaks for individual sites from HPLC maps of unsynchronized WTGRs obtained after 14 h incubation with 32P (2), as well as from the top HPLC map in Fig. 2Go, show, for example, that the ratio of steady state labeling of serine 234 to serine 315 is about 4. The same ratio is found from HPLC maps of WTGRs from cells synchronized in G2/M (bottom map of Fig. 2Go) and synchronized in S (27), despite the fact that basal GR phosphorylation is about 3 times higher in G2/M than in S (27). If all sites in WTGRs from G2/M were filled, the ratio would be about 1. Even if all contributions to each site were not included in the sums, substantial differences between the ratios for WTGRs in S and G2/M would still be expected. Thus, absence of hormone-induced hyperphosphorylation of WTGRs in G2/M cannot be explained by all sites being fully phosphorylated at basal levels.

We are left, then, with the conclusion that hormone-induced hyperphosphorylation of GRs is modulated by overall negative charge in the N-terminal domain. If that conclusion is valid it follows that qualitatively, at least, the charge carrier is not critical, since negative charges carried in E7GRs by glutamates (Fig. 3Go) can block hyperphosphorylation as effectively as those in WTGRs carried by phosphates (27). Quantitatively there may be differences, however: E7GR molecules are negatively charged at all seven mutated sites, whereas WTGR molecules on average are phosphorylated, and therefore negatively charged, at only a fraction of those sites. The region from about residues 195 to 260, which includes the most heavily phosphorylated sites in WTGRs (2), is strongly acidic even without phosphorylation (29), suggesting that phosphorylation modulates negative charge above and below a threshold.

Whether particular phosphorylated sites or combinations of sites are more effective than others in blocking hyperphosphorylation is not known. The fact that the same set of sites on WTGRs is phosphorylated throughout the cell cycle and after hormone treatment, despite large changes in overall phosphorylation levels (8, 27), and that the sites are neither cell type-specific (2, 7) nor, apparently, species-specific (30), is consistent with a concerted role of the phosphorylated sites to modulate overall negative charge in the N-terminal domain. How, in turn, the high negative charge in G2/M might block hormone-induced hyperphosphorylation is an open question. Possible mechanisms include the alternatives that the negative charge hinders hormone-induced activation of GRs (which, as mentioned earlier, appears to be required for hyperphosphorylation), or causes activated GRs to be sequestered beyond reach of kinases, or interferes with proper contact between the N-terminal domain of the GR and the active sites of kinases.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS AND DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Reagents and Buffers
TA was obtained from Steraloids (Wilton, NH), [3H]TA from New England Nuclear (Boston, MA), and [32P]orthophosphoric acid (carrier free) and [35S]methionine (>1000 Ci/mmol) from ICN Biomedicals (Irvine, CA). Ham’s nutrient mixture F12 was from JRH Biosciences (Denver, PA), and DMEM was from GIBCO BRL (Gaithersburg, MD). Methylated trypsin was from Promega Biotec (Madison, WI). Iron-supplemented calf serum and hydroxyurea were from Sigma (St. Louis, MO). Other reagents and buffers have been described previously (2, 28).

Cells and Cell Culture
Cells with A7GRs and E7GRs were grown at 37 C with 5% CO2 in DMEM (4.5 g glucose/liter) containing 10% charcoal-stripped calf serum supplemented with 39.5 mg proline/liter and 100 nM methotrexate. WCL2 cells were grown in the same medium but with 3 µM methotrexate.

Charcoal-Stripped Serum
A suspension of dextran-coated charcoal was prepared as described previously (31), except that 10-fold higher concentrations of charcoal and dextran were used, and PBS was substituted for MgCl2. The pellet obtained by centrifuging 25 ml of the suspension at 10,000 x g for 10 min was resuspended with 250 ml of iron-supplemented calf serum and incubated for 30 min at 23–25 C. After centrifugation at 10,000 x g for 10 min, the serum was decanted from the charcoal, then treated once more with dextran-coated charcoal before sterile filtering through a 0.2-µm filter.

Mutation of GRs
Mouse GR phosphorylation mutants with phosphorylated sites changed to either alanine or glutamate were produced by the Kunkel method (32, 33) as described previously (28). Two mutant GRs, A7GR and E7GR, were generated for this study. In the A7GR, serines 150, 212, 220, 234, 315, 412, and threonine 159 were changed to alanine. Serine 122 was not mutated: it remained as a reporter of phosphorylation. In the E7GR the same sites were changed to glutamate, again leaving serine 122 intact.

Generation of Stably Transfected Cell Lines
The general procedures have been described by Hirst et al. (34). Briefly, mutant GR cDNA and a selectable dhfr cDNA (pSV2dhfr) were cotransfected into DG44 cells [a Chinese hamster ovary cell line missing both alleles of the dhfr gene (35)] by calcium phosphate precipitation. After 48 h the cells were exposed to different concentrations of methotrexate starting from 100 nM. Only cells that express dhfr survive in the presence of methotrexate. Clones expressing relatively high numbers of GRs were selected by [3H]TA binding assay (27). Those used for the present studies had approximately 600,000 A7GRs and about 800,000 E7GRs per cell. SDS-PAGE and Western blot analyses showed that the mutant GRs had the same molecular mass (~100 kDa) as WTGRs and that no other receptor forms were present. Binding affinities of the mutant GRs for TA were indistinguishable from that of wild type GRs.

Cell Synchronization and Metabolic Labeling
Procedures used were those described for synchronization of WCL2 cells (27). Briefly, cells with mutant GRs were incubated under serum deprivation conditions (0.1% calf serum) for 48 h at 37 C, which partially synchronized them in G0/G1. They were then synchronized in late G1/early S phase by incubation in 1 mM hydroxyurea for 24 h. To lower phosphate and methionine levels for labeling, during the last hour they were incubated in phosphate-free, low methionine (5.4 mg/liter) medium with 1 mM hydroxyurea. Hydroxyurea was washed out (time = 0), and the cells were divided equally among four flasks (~4 x 106 cells per flask), to each of which was added [35S]methionine in phosphate-free, low methionine labeling medium. Culture in this medium (as compared with medium with normal methionine and phosphate) did not affect cell progression through the cell cycle.

[32P]orthophosphoric acid was added to one pair of flasks at 0.5 h for S phase labeling and to the other pair at 6.5 h for G2/M labeling. TA (500 nM) was added to one of each pair of flasks 3 h later. After 1 h the cells were harvested and the cell pellets frozen with liquid nitrogen in FTT buffer (freeze-thaw buffer + 0.2% Triton X-100).

For cell cycle analysis, in parallel with the cells used for labeling, some cells were cultured in labeling medium without radioactivity and incubated for various times, after which they were harvested and fixed in ethanol. To measure relative DNA content the cells were treated with RNase A, stained with propidium iodide (36), and analyzed by flow cytometry (FACScan, Becton Dickinson, Franklin Lakes, NJ). The synchronization procedure yielded approximately 85% of cells in S phase from 0 to 5 h after release from hydroxyurea, and 85% in G2/M from 8 to 12 h after release.

Purification and HPLC Phosphopeptide Mapping of GRs
As described for WTGRs (2, 28), A7GRs and E7GRs were extracted and purified by immunopurification and SDS-PAGE. To determine the magnitude of hormone-induced hyperphosphorylation, the total 32P incorporation into GRs was normalized to total 35S incorporation, the latter providing a relative measure of GR protein (3). The purified A7GRs and E7GRs were digested with trypsin, and the resulting phosphopeptides were analyzed by HPLC as detailed previously (2, 28).


    ACKNOWLEDGMENTS
 
We thank Dr. Margaret Hirst for the kind gift of WCL2 cells, Dr. Lawrence Chasin for generously providing the DG44 cells, and Ms. Fiona Swift for excellent technical assistance.


    FOOTNOTES
 
Address requests for reprints to: Allan Munck, Department of Physiology, Dartmouth Medical School, Lebanon, NH 03756-0001, Phone: 603-650-7734, Fax: 603-650-6130.

This research was supported by Research Grants DK-03535, DK-47329, and DK-45337 from the NIH and by the Norris Cotton Cancer Center Core Grant CA-23108. J.-M. H. was supported by a predoctoral fellowship from the Norris Cotton Cancer Center.

1 Abbreviations and trivial names used: A7GR, mutant GR with all but one phosphorylated site mutated to alanine; E7GR, mutant GR with all but one phosphorylated site mutated to glutamate; hsp90, approximately 90-kDa heat shock protein; G1 phase, gap 1 phase of cell cycle; S phase, DNA synthesis phase of cell cycle; G2, gap 2 phase of cell cycle; M, mitosis phase of cell cycle; G2/M, gap 2 and mitosis phases of cell cycle; RU486, 17ß-hydroxy-11ß,4-dimethylaminophenyl-17{alpha}-propynyl estra-4,9-diene-3-one; TA, tri-amcinolone acetonide, 9{alpha}-fluoro-11ß,16{alpha},17{alpha},21-tetrahydroxypregna-1,4-diene-3,20-dione-16,17-acetonide; dhfr, di-hydrofolate reductase. Back

Received for publication October 29, 1996. Revision received December 2, 1996. Accepted for publication December 5, 1996.


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 RESULTS AND DISCUSSION
 MATERIALS AND METHODS
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