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Endocrinology Vol. 139, No. 9 3813-3821
Copyright © 1998 by The Endocrine Society

ARTICLES

Glucocorticoid-Induced Apoptosis and Regulation of NF-{kappa}B Activity in Human Leukemic T Cells1

Jyoti Ramdas and Jeffrey M. Harmon

Department of Pharmacology, Uniformed Services University of the Health Sciences, Bethesda, Maryland 20814-4799

Address all correspondence and requests for reprints to: Jeffrey M. Harmon, Ph.D., Department of Pharmacology, Uniformed Services University of the Health Sciences, 4301 Jones Bridge Road, Bethesda, Maryland 20814-4799. E-mail: jharmon{at}mxb.usuhs.mil


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Glucocorticoid-induced apoptosis was investigated in glucocorticoid-sensitive 6TG1.1 and resistant ICR27TK.3 human leukemic T cells. Following glucocorticoid treatment of 6TG1.1 cells, chromatin fragmentation was observed after a delay of 24 h. Fragmentation was not observed in ICR27TK.3 cells containing mutant glucocorticoid receptors (L753F) that are activation-deficient but retain the ability to repress AP-1 activity. Nor was fragmentation observed after treatment with RU38486, indicating that repression of AP-1 activity is not involved. As described in other systems, fragmentation required ongoing protein synthesis. However, inhibition of protein synthesis with cycloheximide anytime during the first 18 h of steroid treatment was as effective in blocking chromatin fragmentation as inhibition for the entire period, suggesting that synthesis of a component with a rapid turnover rate is required. Dexamethasone treatment completely blocked 12-O-tetradecanoylphorbol 13-acetate induction of nuclear factor-{kappa}B (NF-{kappa}B) activity and elicited an increase in the amount of immunoreactive I{kappa}B{alpha} in sensitive 6TG1.1 cells but not in resistant ICR27TK.3 cells. In addition, mild detergent treatment of cell extracts indicated that a substantial amount of cytoplasmic NF-{kappa}B is complexed with I{kappa}B{alpha} or some other inhibitory factor. These results suggest that induction of a labile inhibitory factor such as I{kappa}B{alpha} may contribute to glucocorticoid-induced apoptosis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
APOPTOTIC death of T cells can be initiated by a variety of stimuli including activation of Fas/CD95/ApoI, tumor necrosis factor (TNF)-{alpha}, radiation, DNA damage, oxidative stress, and exposure to glucocorticoids (1, 2, 3, 4, 5). Indeed, the ability of glucocorticoids to induce growth arrest and death of lymphoid cells has resulted in their widespread use in the treatment of multiple myeloma, and various leukemias and lymphomas (6, 7, 8). Analysis of the events which result in cell death, as well as factors that inhibit it, suggest that activation of caspase-3 is a common component of both glucocorticoid and nonglucocorticoid-induced T cell apoptosis (9, 10, 11, 12, 13). However, unlike apoptosis induced by other stimuli, glucocorticoid-induced apoptosis is dependent upon ongoing protein synthesis (14, 15, 16), an observation consistent with the requirement for the synthesis of a new protein, and/or the continuous presence of a protein with a rapid turnover rate. Similarly, the dependence of glucocorticoid-induced apoptosis on an intact transactivating domain in the N-terminus of the glucocorticoid receptor (GR) suggests that induction of gene expression is required for cell death (17).

There is also evidence supporting the hypothesis that glucocorticoid-induced apoptosis is the result of repression of genes whose products are essential for cell growth. c-myc expression is repressed in mouse S49 (18) and human CEM-C7 (19) cells, and transient overexpression of exogenous c-myc blocks cell death (20). In P1798 cells, both c-myc and cyclin D3 levels are repressed by steroid treatment and growth inhibition is absent in stably transfected cells overexpressing both proteins (21). In addition, Jurkat cells expressing mutant GR unable to stimulate transcription are still sensitive to glucocorticoid-induced apoptosis, presumably due to the fact that the mutant receptors retain the ability to repress AP-1 and other glucocorticoid-sensitive activities (22). Similarly, transient expression of fragments of the GR DNA binding domain lacking transactivating activity induce cell death in otherwise resistant CEM cells, suggesting that the repressive activity of the GR, mediated through protein-protein interactions is responsible for cell death (23).

Recently, it has been shown that the activity of NF-{kappa}B, which regulates the expression of numerous cytokine genes, is repressed by glucocorticoids (24, 25, 26, 27, 28). In addition, TNF-{alpha}-induced apoptosis is opposed by high levels of NF-{kappa}B activity (29, 30, 31, 32), suggesting that glucocorticoid-mediated repression of NF-{kappa}B could contribute to the antiinflammatory activity of glucocorticoids and to glucocorticoid-induced apoptosis. Repression of NF-{kappa}B activity has been attributed to direct interaction between the NF-{kappa}B heterodimer and the GR (24, 27, 33, 34, 35, 36, 37). However, glucocorticoids also induce the labile inhibitory protein I{kappa}B{alpha} (25, 26, 36), suggesting that the requirement for protein synthesis in glucocorticoid-induced apoptosis could reflect the need to synthesize this labile inhibitory factor.

To examine the relationship between glucocorticoid-induced apoptosis, protein synthesis, and GR-mediated repression of NF-{kappa}B activity, glucocorticoid-sensitive 6TG1.1 cells were employed. These cells are derived from the clonal human leukemic cell line CEM-C7 (38). In response to glucocorticoid treatment, they are irreversibly arrested in the G1 phase of the cell cycle after a delay of 18–24 h and undergo many of the morphological features characteristic of apoptosis (38, 39, 40). Glucocorticoid-induced chromatin fragmentation and repression of NF-{kappa}B activity were also examined in glucocorticoid-resistant ICR27TK.3 cells that contain mutant, activation-deficient GR that retain the ability to repress AP-1 activity (41, 42). Our results show that glucocorticoid-induced chromatin fragmentation is independent of AP-1 repression and requires the synthesis of a labile protein. In addition, glucocorticoids blocked 12-O-tetradecanoylphorbol 13-acetate (TPA) induction of NF-{kappa}B activity, and elicited an increase in the amount of immunoreactive I{kappa}B{alpha}, suggesting that induction of a labile NF-{kappa}B inhibitory factor may contribute to glucocorticoid-induced apoptosis.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Dexamethasone, cycloheximide, etoposide, and zinc acetate dihydrate were obtained from Sigma Chemical Co. (St. Louis, MO). RU38486 was the generous gift of Roussel UCLAF (Romainville, France). Proteinase K and RNase A were obtained from Boehringer Mannheim (Indianapolis, IN). RPMI 1640 and glutamine were purchased from Life Technologies, Inc. (Gaithersburg, MD). FBS was obtained from JRH Biosciences (Lenexa, KS). Seakem Gold and InCert agaroses were purchased from FMC BioProducts (Rockland, ME). Prestained broad range molecular weight markers for SDS-PAGE were obtained from Bio-Rad Laboratories (Hercules, CA). D-luciferin was obtained from Analytical Luminescence Laboratory (San Diego, CA).

Cell culture
The glucocorticoid-sensitive T cell line 6TG1.1 and its glucocorticoid-resistant derivative ICR27TK.3 have been described previously (38). Cells were maintained in RPMI 1640 tissue culture medium containing 10% FBS and grown at 37 C in a humidified atmosphere containing 5% CO2 as previously described (39).

DNA analysis
For field inversion gel electrophoresis, cells were harvested by centrifugation at 800 x g for 10 min at 4 C. The cell pellet was washed with ice-cold Dulbecco’s PBS and resuspended at a concentration of 8 x 106 cells/ml in cold lysis buffer (0.01 M Tris-HCl, pH 7.5 containing 0.1 M EDTA, and 0.02 M NaCl). Agarose plug molds containing 4 x 106 cells/ml were prepared by mixing 200 µl of cell suspension with an equal volume of 1.6% InCert agarose dissolved in lysis buffer maintained at 42 C. Plugs were incubated at 50 C for 70 h in lysis buffer containing 0.1% N-lauryl sarcosine and 100 µg/ml Proteinase K, and stored at 4 C in 0.5 M EDTA, pH 8.0. Before electrophoresis, plugs were equilibrated 10 mM Tris-HCl, pH 8.0 containing 1 mM EDTA. High molecular weight DNA fragmentation was analyzed by field inversion gel electrophoresis performed at 12 C and 6 volts/cm in 1% Seakem Gold agarose gels buffered with 0.5 x TBE (44.5 mM Tris, 44.5 mM boric acid, 1 mM EDTA). Each run was begun with a 10 min "run-in," followed by a forward to reverse ratio of 3:1, and a linear ramp of 5–50 seconds, for a total of 21 h. DNA was visualized by ethidium bromide staining.

Transient transfection and luciferase reporter assay
Cells were harvested during log phase growth by centrifugation at 200 x g for 10 min at 4 C and resuspended in ice-cold electroporation medium (RPMI 1640 medium containing 25 mM HEPES) at a concentration of 107 cells/ml. Five million cells (0.5 ml) were transferred to a precooled electroporation cuvette (0.4 cm gap, Bio-Rad) to which 10 µg of supercoiled pLTRluc (43) or p4XAP-1-luciferase (44) were added. After incubation for 10 min at 4 C, electroporation was performed with a Bio-Rad Genepulser apparatus (with capacitance extender) at 340 V and 960 µF. Electroporated cells were incubated at 4 C for 10 min and then transferred to 5 ml of culture medium composed of a 1:1 mixture of fresh and conditioned medium containing 10% FBS, preequilibrated for 24 h at 37 C in a humidified atmosphere containing 5% CO2. Cells were allowed to recover for 24 h, then diluted with an equal volume of fresh RPMI 1640 containing 10% FBS. Five milliliters were transferred to a fresh 35-mm well and incubated at 37 C and 5% CO2 for an additional 24 h. After hormone treatment, cells were harvested by centrifugation at 200 x g for 10 min and washed once with ice-cold PBS. Cells were lysed by three cycles of freezing in a dry ice/ethanol bath and thawing at 37 C, followed by suspension in 120 µl of lysis buffer [100 mM potassium phosphate, pH 7.8, containing 0.2% Triton X-100 and 1 mM dithiothreitol (DTT)], and centrifugation at 15,800 x g for 2 min at 4 C. To measure luciferase activity, the supernatant (100 µl) was added to 368 µl of 25 mM glycylglycine buffer (pH 7.8), containing 15 mM MgSO4, 4 mM EGTA, 15 mM potassium phosphate, 1 mM DTT, and 2 mM ATP. The reaction was initiated by addition of 200 µl of 0.25 mM luciferin, and light output was measured for 10 sec using a Laboratory Technologies luminometer (Roselle, IL). Protein concentration was determined by the method of Bradford (45). Luciferase activity was expressed as relative light units per 100 µg protein.

Preparation of nuclear and cytoplasmic extracts
Nuclear and cytoplasmic extracts were prepared as described (46) with minor modification. One to 10 million cells were harvested by centrifugation, and washed once with ice-cold PBS. The cell pellet was frozen on dry ice, allowed to thaw on wet ice, and resuspended in 200 µl of lysis buffer (10 mM HEPES-KOH, pH 7.9, containing 5 mM MgCl2). The suspension was vortexed at high speed for 10 sec and centrifuged at 320 x g for 2 min at 4 C. The supernatant (cytoplasmic extract) was centrifuged for an additional 10 min at 15,800 x g at 4 C, and glycerol added to a final concentration of 20%. Aliquots were frozen and stored at -70 C. The crude nuclear pellet was rinsed twice in a 10 mM HEPES-KOH buffer (pH 7.9) containing 5 mM MgCl2, and 0.1% NP-40, and the final pellet was resuspended in elution buffer (10 mM HEPES-KOH, pH 7.9, containing 420 mM NaCl, and 25% glycerol), vortexed at slow speed for 10 min at 4 C, and centrifuged at 15,800 x g for 15 min at 4 C. The supernatant (nuclear extract) was diluted 3-fold by the addition of 20 mM HEPES buffer, pH 7.9, containing 100 mM KCl, 0.2 mM EDTA, and 20% glycerol, and dialyzed against the same buffer for 2 h at 4 C. The dialysate was centrifuged at 15,800 x g for 10 min at 4 C, and aliquots of the supernatant were frozen and stored at -70 C. All buffers contained aprotinin (20 µg/ml), leupeptin (20 µg/ml), pepstatin (5 µg/ml), phenylmethylsulfonyl fluoride (PMSF, 1 mM), and DTT (1 mM), except the dialysis buffer which contained only PMSF and DTT.

Electrophoretic mobility shift assay (EMSA)
Synthetic complementary oligonucleotides were annealed in 10 mM Tris-HCl buffer (pH 8.0) containing 50 mM NaCl, 10 mM MgCl2 and 1 mM DTT. Double-stranded oligonucleotides were labeled with [{alpha}-32P]dCTP using the Klenow fragment of DNA polymerase I (New England Biolabs, Beverly, MA) and purified by Sephadex G-25 chromatography. The following pairs of complementary oligonucleotides were used (consensus binding site is shown in boldface): NF-{kappa}B, 5'-AGCTCAGAGGGGACTTTCCGAGAG-3' and 5'-AGCTCTCTCGGA-AAGTCCCCTCTG-3' (47, 48); interferon responsive element (IRE), 5'-AGTGATTTCTCGGAAAGAGAG-3' and 5'-AGGCTTCTTTCCGA-GAAAT-3' (49, 50). DNA-protein binding reactions were carried out essentially as described by Bauerle and Baltimore (51). Extracts (5–10 µg protein) were preincubated for 10 min at room temperature in 20 µl of 10 mM Tris-HCl, pH 7.5, containing 50 mM NaCl, 0.5 mM EDTA, 5% glycerol, 1 mM DTT, and 1 µg poly(dI-dC). In some experiments, sodium deoxycholate (0.6%) and NP-40 (1.2%) were also included. For supershift experiments, 1 µl of anti-p50 or anti-p65 rabbit antisera (a generous gift from Dr. Nancy Rice, National Cancer Institute) was added. The binding reaction was initiated by addition of 0.2 ng of 32P-labeled NF-{kappa}B oligonucleotide, and continued for 30 min at room temperature. Samples were resolved on 5% polyacrylamide gels in 0.25 x TBE, and electrophoresed in the same buffer at 6 V/cm at room temperature for 2–3 h. The gels were dried and exposed to x-ray film (Kodak XAR, Eastman Kodak, Rochester, NY) at -70 C.

Immunoblotting
Cytoplasmic extracts, prepared for EMSA as described above, were resolved by SDS-PAGE on 12% polyacrylamide gels and transferred to nitrocellulose filters essentially as previously described (52). I{kappa}B{alpha} protein was visualized by enhanced chemiluminescence (Amersham, Arlington Heights, IL) using a 1:5000 dilution of anti-I{kappa}B{alpha} antibody sc-371(Santa Cruz Biotechnology Inc., Santa Cruz, CA), followed by incubation with a 1:2000 dilution of donkey antirabbit IgG conjugated to horseradish peroxidase. Filters were then incubated with equal portions of luminol and hydrogen peroxide for 1 min, followed by exposure to Hyperfilm (Amersham). For quantification of chemiluminesence, filters were processed using a Bio-Rad Model GS-250 Molecular Imager.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
We have previously shown that treatment of the glucocorticoid-sensitive human T cell line 6TG1.1 with dexamethasone results in irreversible arrest of cells in the G1 phase of the cell cycle and subsequent cell death (39, 40). However, in contrast to the rapid apoptotic response seen in glucocorticoid-treated murine thymocytes and T cell lines, steroid-treated 6TG1.1 cells remain 100% viable for the first 18–24 h of steroid treatment. In addition, if steroid is removed at any time during the first 24 h, no cell cycle arrest or cell death are observed (40). To determine whether inhibition of cell growth and cell death are accompanied by high molecular weight chromatin fragmentation, 6TG1.1 cells were treated with 1 µM dexamethasone for various periods of time, and DNA was analyzed by field inversion gel electrophoresis to resolve fragments in the range of 10 kb to more than 500 kb (53). After 24 h of steroid treatment, there was no discernible difference between the integrity of DNA in treated and untreated cells (Fig. 1Go, lanes 1 and 2). However, 48 h after steroid treatment there was substantial cleavage of DNA to fragments of approximately 50 kb, which was even more pronounced 72 h after treatment (Fig. 1Go, lane 6).



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  		Figure 1. High molecular weight DNA fragmentation in glucocorticoid-sensitive and -resistant cells. Glucocorticoid-sensitive 6TG1.1 cells (lanes 1–6), and glucocorticoid-resistant ICR27TK.3 cells (lanes 7–12) were incubated in the absence (odd numbered lanes) or presence (even numbered lanes) of 1 µM dexamethasone for 24, 48, or 72 h. DNA was analyzed by field inversion gel electrophoresis as described in Materials and Methods. Lanes M1, M2, and M3 contain {lambda} multimers, 1-kb ladder, and high molecular weight DNA markers, respectively.

 
No chromatin fragmentation was observed in steroid-resistant ICR27TK.3 cells (Fig. 1Go) that contain the activation-deficient hGR mutant L753F (41, 42, 54). The absence of chromatin fragmentation in ICR27TK.3 cells does not reflect the absence of endogenous nuclease activity in these cells because treatment with etoposide resulted in significant, dose-dependent high molecular weight chromatin fragmentation (data not shown). We have previously shown that activation-deficient L753F receptors retain the ability to repress AP-1 activity when transfected into CV-1 cells (41). This mutant also retains the ability to repress AP-1 activity in glucocorticoid-resistant ICR27TK.3 cells. When cells were transfected with the AP-1 inducible reporter p4XAP-1-luciferase, which contains four tandem AP-1 elements (44), dexamethasone inhibited TPA induction of luciferase activity to the same extent as in glucocorticoid-sensitive 6TG1.1 cells transfected with the same reporter (Fig. 2Go). This was in sharp contrast to the results obtained when cells were transfected with the glucocorticoid-inducible reporter pLTRluc, where induction of luciferase activity was only seen in glucocorticoid-sensitive 6TG1.1 cells containing wild-type GR (Fig. 2Go). Thus, repression of AP-1 activity is not sufficient for glucocorticoid-induced chromatin fragmentation.



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  		Figure 2. Induction and repression by glucocorticoid receptor mutant L753F. A, 6TG1.1 and ICR27TK.3 cells were transfected with pLTRluc and incubated for 23 h in the absence or presence of 1 µM dexamethasone. Luciferase activity was determined as described in Materials and Methods. Results represent the average of three separate experiments and are presented in terms of relative luciferase activity, with the activity seen in untreated samples set to unity. B, 6TG1.1 and ICR27TK.3 cells were transfected with p4XAP-1-luciferase. Forty-eight hours after transfection, dexamethasone (1 µM) or vehicle (95% EtOH) was added to the cells, followed 7 h later by the addition of TPA (50 ng/ml). Results represent the average of two to three separate experiments and are presented in terms of relative luciferase activity, with the activity seen in the absence of dexamethasone set arbitrarily to 100.

 
Similarly, treatment of glucocorticoid-sensitive or glucocorticoid-resistant cells with RU38486, which elicits GR-mediated repression of AP-1 activity (41, 55), had no effect on chromatin integrity (Fig. 3Go). These results suggest that repression of AP-1 activity does not contribute to cell death in 6TG1.1 cells and are consistent with evidence that the transactivating activity of the GR is required for steroid-induced apoptosis (17). To examine the requirement for synthesis of a steroid-inducible factor, cells were exposed for 48 h to various concentrations of cycloheximide in the presence or absence of 1 µM dexamethasone. Examination of high molecular weight DNA showed that as little as 200 nM cycloheximide effectively inhibited glucocorticoid-induced chromatin fragmentation (Fig. 4Go). This concentration of cycloheximide has been shown to inhibit the synthesis of some glucocorticoid-induced proteins in CEM cells (16). At 500 nM cycloheximide, inhibition of fragmentation was nearly complete. Fragmentation induced by cycloheximide itself was not seen at concentrations below 1 µM (Fig. 4Go, lanes 4, 5, 9, and 10). These results are consistent with previous studies that demonstrated a requirement for new protein synthesis in glucocorticoid-induced apoptosis of murine thymocytes, human CEM cells, and a variety of other susceptible cells (14, 15, 16). However, they cannot differentiate between the need to synthesize a new protein and the need to maintain the concentration of an existing protein that is rapidly turned over. To more completely examine the role of new protein synthesis, cycloheximide was added at various times after the addition of 1 µM dexamethasone, and high molecular weight DNA fragmentation analyzed by field inversion gel electrophoresis 48 h after steroid treatment. The concentration of cycloheximide used (200 nM) was the lowest that effectively inhibited chromatin fragmentation. The results showed that addition of cycloheximide as late as 24 h after addition of dexamethasone was as effective in inhibiting high molecular weight DNA fragmentation as simultaneous addition (Fig. 5Go). Although addition of cycloheximide after the first 24 h was less effective, addition at any time up to 36 h after steroid treatment still reduced the amount of high molecular weight DNA fragmentation compared with cells not treated with cycloheximide at all. Thus, the synthesis of the factor(s) responsible for chromatin fragmentation and cell death may be a relatively late event, or the factor may have a rapid rate of turnover.



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  		Figure 3. RU38486 inhibits dexamethasone-induced high molecular weight fragmentation. 6TG1.1 cells were incubated for 48 h in the absence of hormone (lane 1), in the presence of 1 µM RU38486 (lane 2), 100 nM dexamethasone (lane 3), or 1 µM RU38486 and 100 nM dexamethasone (lane 4). High molecular weight DNA fragmentation was analyzed as described in the legend to Fig. 1Go. Lanes M1, M2, and M3 contained {lambda} multimers, 1-kb ladder, and high molecular weight DNA markers, respectively.

 


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  		Figure 4. Glucocorticoid-induced high molecular weight DNA fragmentation requires protein synthesis. 6TG1.1 cells were incubated with increasing concentrations of cycloheximide in the absence (lanes 1–5) or presence (lanes 6–10) of 1 µM dexamethasone for 48 h. Lanes 1 and 6, no cycloheximide; lanes 2 and 7, 200 nM cycloheximide; lanes 3 and 8, 500 nM cycloheximide; lanes 4 and 9, 1 µM cycloheximide; lanes 5 and 10, 2 µM cycloheximide. High molecular weight DNA fragmentation was analyzed by field inversion gel electrophoresis as described in the legend to Fig. 1Go. Lanes M1, M2, and M3 contained {lambda} multimers, 1-kb ladder, and high molecular weight DNA markers respectively.

 


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  		Figure 5. Kinetics of inhibition of high molecular weight DNA fragmentation. 6TG1.1 cells were incubated in the absence (lanes 1 and 2) or presence (lanes 3–11) of 1 µM dexamethasone for 48 h. Cycloheximide (200 nM) was added 0 (lane 3), 6 (lane 4), 12 (lane 5), 18 (lane 6), 24 (lane 7), 30 (lane 8), 36 (lane 9), or 42 (lane 10) h after the addition of steroid. Lane 2 contains DNA from cells treated for 48 h with cycloheximide alone. High molecular weight DNA fragmentation was analyzed as described in the legend to Fig. 1Go. Lanes M1, M2, and M3 contained {lambda} multimers, 1-kb ladder and high molecular weight DNA markers, respectively.

 
Inhibition of heterodimeric NF-{kappa}B activity is an important component of the antiinflammatory activity of glucocorticoids (26, 28, 34). In addition, it has recently been proposed that this inhibition is the result of induction of labile repressor I{kappa}B (25, 26). To examine glucocorticoid regulation of NF-{kappa}B activity in glucocorticoid-induced apoptosis, cytoplasmic and nuclear extracts prepared from steroid treated and untreated 6TG1.1 cells were used in EMSA to retard the mobility of a 32P-labeled oligonucleotide containing the NF-{kappa}B consensus binding sequence. Basal NF-{kappa}B activity in 6TG1.1 cells was low. However, growth in the presence of 1 µM dexamethasone effectively blocked TPA induction of both nuclear and cytoplasmic NF-{kappa}B activity (Fig. 6AGo). This activity was competed by an excess of unlabeled NF-{kappa}B oligonucleotide, but not an unlabeled oligonucleotide containing an IRE (Fig. 6Go). In addition, the larger of the two TPA-induced complexes was supershifted by antibodies directed against either p65/RelA or p50 (Fig. 6CGo), confirming its identity. The mobility of complex 1 was not altered, nor was DNA binding inhibited, by incubation with anti-hGR antibody 710 (54). This result confirms the specificity of the supershift assay, and demonstrates that the hGR is not associated with cytoplasmic p65/p50 heterodimers. Based on its mobility, the smaller of the two TPA-induced complexes seen in nuclear extracts prepared from TPA-treated cells most probably represents p50/p50 homodimers and appears to be present at higher concentrations in nuclear extracts (Fig. 6Go, A and B), than in cytoplasmic extracts (Fig. 6CGo). Most importantly, dexamethasone did not block TPA induction of NF-{kappa}B activity in glucocorticoid-resistant ICR27TK.3 cells (Fig. 6BGo), suggesting that mutant L753F GR does not inhibit NF-{kappa}B activity.



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  		Figure 6. Dexamethasone inhibition of NF-{kappa}B activity. A, 6TG1.1 cells were incubated for 18 h in the absence (lanes 2, 3, 6, and 7) or presence (lanes 4 and 5 and 8–15) of 50 ng/ml TPA and the absence (lanes 2, 4, 6, 8, 10, 12, 14, and 15) or presence (lanes 3, 5, 7, 9, 11, and 13) of 1 µM dexamethasone. Cytoplasmic and nuclear extracts were prepared as described in Materials and Methods. EMSA was performed using a 32P- labeled NF-{kappa}B probe in the absence (lanes 1–13) or presence of a 50-fold excess of unlabeled NF-{kappa}B (lane 14) or IRE (lane 15) double-stranded oligonucleotide either before (lanes 1–5, and 10–15), or after (lanes 6–9) treatment with sodium deoxycholate (DOC, 0.6%) and 1.2% NP-40. Lane 1 contains labeled probe alone. Arrows to the right of the figure indicate the mobilities of the two NF-{kappa}B-specific complexes. (U, untreated; D, dexamethasone alone; T, TPA-induced; TD, dexamethasone and TPA). B, Nuclear (lanes 2–5) and cytoplasmic (lanes 6–9) extracts were prepared from glucocorticoid-resistant ICR27TK.3 cells incubated for 18 h in the absence (lanes 2,3,6 and 7) or presence (lanes 4, 5, 8, and 9) of 50 ng/ml TPA, and the absence (lanes 2, 4, 6, and 8) or presence (lanes 3, 5, 7, and 9) of 1 µM dexamethasone. EMSA was performed with the same 32P-labeled NF-{kappa}B probe used in A. Lane 1 contains labeled probe alone. Arrows to the right of the figure indicate the mobilities of the two NF-{kappa}B-specific complexes. (U, untreated; D, dexamethasone alone; T, TPA-induced; TD, dexamethasone and TPA). C, EMSA was performed using a 32P-labeled NF-{kappa}B probe incubated in the absence (lanes 1, 5, 7, and 9) or presence (lanes 2–4, 6, 8, and 10) of cytoplasmic extract prepared from TPA-induced 6TG1.1 cells, and the absence (lanes 1, 2, and 5–10) of unlabeled competing oligonucleotide, or in the presence of a 50-fold excess of unlabeled NF-{kappa}B (lane 3) or IRE (lane 4) double-stranded oligonucleotide. Supershifts were performed by incubating either probe alone (lanes 5, 7, and 9) or extracts (lanes 6, 8, and 10) with anti-p50 (lanes 5 and 6), anti-p65 (lanes 7 and 8), or anti-hGR (lanes 9 and 10) antibodies.

 
Mild detergent treatment of nuclear extracts did not increase the amount of retarded complex, indicating that the decrease in nuclear NF-{kappa}B DNA binding activity seen in extracts prepared from glucocorticoid-treated cells is not the result of the presence of inhibitory components in the extracts. In contrast, detergent treatment of cytoplasmic extracts resulted in a substantial increase in the amount of DNA binding activity (Fig. 6AGo), presumably reflecting the presence of inhibitory components. The amount of immunoreactive I{kappa}B{alpha} in cytoplasmic extracts prepared from untreated and steroid-treated 6TG1.1 cells was therefore evaluated. The results revealed a 2- to 2.5-fold increase in the amount of immunoreactive I{kappa}B{alpha} after steroid treatment (Fig. 7Go). Although this increase is small, it was reproducibly seen in each of three independent experiments, and is comparable to that seen in other systems where glucocorticoid induction of I{kappa}B{alpha} antagonizes the activity of TNF-{alpha} and other inflammatory agents (25, 26). No increase in immunoreactive I{kappa}B{alpha}, or decrease in NF-{kappa}B binding activity, was seen in ICR27TK.3 cells after comparable treatment. Because the GR in these cells is capable of nuclear translocation and interaction with other transcription factors (41), it therefore appears that glucocorticoid inhibition of NF-{kappa}B activity in glucocorticoid-sensitive 6TG1.1 cells is, at least in part, the consequence of induction of I{kappa}B{alpha}.



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  		Figure 7. Induction of I{kappa}B{alpha} protein by dexamethasone. 6TG1.1 (lanes 1–4) and ICR27TK.3 (lanes 5–8) cells were incubated for 18 h in the absence (lanes 1, 2, 5, and 6) or presence (lanes 3, 5, 7, and 8) of 50 ng/ml TPA. Dexamethasone (1 µM) was added to both TPA-untreated (lanes 2 and 6), and TPA-induced (lanes 4 and 8) cultures. Cytoplasmic extracts were prepared and analyzed for the presence of immunoreactive I{kappa}B{alpha} protein as described in Materials and Methods. The mobilities of the prestained molecular weight standards (ovalbumin, carbonic anhydrase, soybean trypsin inhibitor, and lysozyme) are shown on the left side of the figure. (U, untreated; D, dexamethasone alone; T, TPA-induced; TD, dexamethasone and TPA).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In contrast to glucocorticoid-induced apoptosis of murine thymocytes and other cells, glucocorticoid-induced apoptosis in human leukemic CEM cells is a relatively slow process. Cell cycle arrest and loss of viability require 24 of continuous steroid treatment (40). However, these cells undergo the same morphological changes, including chromatin fragmentation, seen in glucocorticoid-induced cytolysis of other susceptible cells (16, 38, 39, 40). They therefore provide an ideal system in which to examine the mechanism of glucocorticoid-induced growth inhibition and cell death in the absence of substantial amounts of macromolecular degradation and cell lysis. As has been demonstrated in other systems, steroid induced chromatin fragmentation was dependent upon ongoing protein synthesis. However, it was not necessary to inhibit protein synthesis for the entire period of steroid treatment to block apoptosis; inhibition of protein synthesis at any time before the onset of chromatin fragmentation was sufficient. These results are consistent with the observation that cells remain 100% viable if hormone is withdrawn during the first 18 h of treatment, and suggest that the glucocorticoid-induced component has a rapid rate of turnover.

Glucocorticoid-induced cell death has been proposed to require induction of a "lysis" gene that activates, or supplies a missing component to the apoptotic pathway (56). This hypothesis is supported by the ability of 5-azacytosine to sensitize otherwise resistant cells to glucocorticoid-induced apoptosis (57, 58), and the ability of these cells, when fused to sensitive cells lacking functional GR, to reconstitute an apoptotic response (57, 59). It is also consistent with the contribution of the N-terminal transactivation domain of the GR to steroid-induced apoptosis in mouse S49 cells (17), as well as the lack of glucocorticoid-induced T cell apoptosis in mice expressing an activation-deficient GR that retains the ability to repress collagenase expression (59A ). However, although several glucocorticoid-inducible genes, including the GR gene itself (52, 60) have been identified in lymphoid cells (17, 61, 62, 63, 64, 65, 66), no "lysis" gene has yet been identified.

Alternatively, the ability of glucocorticoids to repress the expression of numerous genes, including c-myc (18, 19, 20), suggests that glucocorticoids repress the activity of a gene(s) necessary for cell survival. This would be consistent with the ability of an activation-deficient GR mutant to inhibit AP-1 activity and c-myc expression, and induce apoptosis in Jurkat cells (22). However, we have shown that a different activation-deficient GR mutant (L753F), which lacks C-terminal ligand-dependent transactivation activity, does not induce apoptosis in CEM cells, although it also retains the ability to repress AP-1 activity (41 and Fig. 2Go). In addition, RU38486 an agonist with respect to repression of AP-1 activity (41, 55), does not induce apoptosis, and completely blocked glucocorticoid-induced chromatin fragmentation. Thus, direct repression of AP-1 activity by GR cannot account for glucocorticoid-induced apoptosis. Indeed, glucocorticoids induce c-jun expression in S49 and CEM cells, suggesting that AP-1 activity may actually be higher in steroid-treated cells (67, 68). Because RU38486 has no immunosuppressive or antiinflammatory activity (69), the ability of RU38486 to repress AP-1 activity also suggests that such repression is either insufficient for, or not involved in, the immunosuppressive or antiinflammatory actions of glucocorticoids.

Many of the antiinflammatory actions of glucocorticoids are mediated through inhibition of NF-{kappa}B (28). In some cases, this inhibition is the consequence of direct interaction between the GR and the p65/p50 heterodimer (24, 27, 33, 34, 35, 36, 37). In other cases inhibition is mediated through induction of the labile NF-{kappa}B inhibitor I{kappa}B (25, 26, 36). Our results indicate that glucocorticoid treatment of 6TG1.1 cells results in both decreased NF-{kappa}B activity and increased immunoreactive I{kappa}B{alpha}. These results are consistent with the ability of a variety of cytokines, including IL-2 and IL-6, whose expression is positively regulated by NF-{kappa}B to inhibit glucocorticoid-induced apoptosis (70, 71, 72, 73, 74, 75), as well as the ability of a dominant-negative I{kappa}B mutant to protect cells from TNF-induced apoptosis (31). They are also consistent with the observation that synthesis of a component with a rapid rate of turnover is necessary for apoptosis and the fact that no repression of NF-{kappa}B activity was seen in steroid-resistant ICR27TK.3 cells where the repressive activity of the GR is retained. Given that c-myc expression is regulated by NF-{kappa}B (76, 77, 78), it is therefore possible that the repressive effects of glucocorticoids seen in susceptible cells are the indirect result of induction of an inhibitory molecule such as I{kappa}B which in turn leads to repression of NF-{kappa}B responsive genes. The identity of the factor(s) induced or repressed by glucocorticoids directly responsible for growth arrest and cell death remain to be determined. However, the results presented here suggest that it may be possible to reconcile evidence supporting induction of gene expression with that supporting repression.


   Footnotes
 
1 This investigation was supported by United States Public Health Service Grant CA-32226, awarded by the National Cancer Institute (to J.M.H.). The opinions or assertions contained herein are the private ones of the authors and are not to be construed as official or reflecting the view of the DoD or the USUHS. Back

Received December 19, 1997.


   References
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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