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The Laboratory of Signal Transduction, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, North Carolina, USA
1Correspondence: 111 T.W. Alexander Dr., Bldg. 101, MD F3–07, Research Triangle Park, NC 27709, USA. E-mail: cidlowski{at}niehs.nih.gov
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: Bcl-2 expression • glucocorticoids • cell shrinkage
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, dimerization (2
, 3)
, and/or proteolytic cleavage (4)
. Although emerging evidence suggests that phosphorylation of Bcl-2 regulates its function, whether Bcl-2 is activated or inactivated by phosphorylation remains controversial (1
, 5)
. Ultimately, the phosphorylation of Bcl-2 may affect its capacity for dimerization with other proapoptotic proteins and/or its stability. For instance, taxol is a chemotherapeutic agent used to treat leukemia and prostatic and nasopharyngeal cancers, and recent studies show that apoptosis induced by taxol is accompanied by Bcl-2 phosphorylation (6
, 7)
. Post-translational modification of Bcl-2, including phosphorylation, has been shown to be associated with its proteosome-dependent degradation (8)
. Bcl-2 has also been suggested to be activated by phosphorylation (9
10
11)
, although opposite roles of phosphorylation in regulating Bcl-2 have been described. This multitude of effects can be accounted for by the multiple phosphorylation sites in a nonconserved loop between BH3 and BH4 domains, which have been proposed to be important for Bcl-2 activity (12
13
14)
. Thus, Bcl-2 phosphorylation status may be modulated by a variety of protein kinases and phosphatases, including those regulated by glucocorticoids (15
16
17)
.
Bcl-2 family proteins have also been proposed to have a role in regulating permeability of the mitochondrial membrane, which is thought to be important during the induction of apoptosis. Some members of the Bcl-2 family have also been shown to be located in endoplasmic reticulum and nuclear envelope (18)
. These molecules may play key roles in regulating Bcl-2 action. Many studies have focused on the interaction between Bcl-2 family members and the permeability transition (PT) pores (19
, 20)
. The PT pores have been proposed to be involved in the change in mitochondrial membrane potential due to ion movement and cytochrome c release (21)
, which in turn activates downstream effector caspases important in the induction of apoptosis. PT pores have been suggested to be regulated by direct interaction with Bcl-2 family proteins (22)
such as Bcl-2, Bcl-XL, and Bax. The 3-dimensional structure suggests that Bcl-2 family proteins are also capable of forming channels in mitochondrial membrane (23
24
25
26)
.
It has been shown that apoptosis induced by glucocorticoids can be blocked by expression of Bcl-2 (27
, 28)
. Since phosphorylation in a nonconserved domain has been suggested to be important in regulating Bcl-2 function (29)
, we have investigated the role of this process in inhibiting glucocorticoid-induced apoptosis. Our results suggest that Bcl-2’s ability to inhibit steroid-induced cell death depends on Bcl-2’s phosphorylation status and suggest that phosphorylation of Bcl-2 on threonine 56 or serine 87 appears to be critical for its anti-apoptotic action. Phosphorylation of Bcl-2 on threonine 74 may play a partial role in Bcl-2’s anti-apoptotic function, whereas phosphorylation of S70 appears to be irrelevant for its anti-apoptotic action in glucocorticoid-induced death of lymphocytes.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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) was used for each reaction (T56A: 5'-CAGCCCCGGGGCACGCGCCCCATCCAG-3' and 5'-CTGGATGGGGCGCGT-GCCCGGGCTG-3'; S70A: 5'-GTCGCCAGGACCGCGCCACTACAGAC-3' and 3'-GTC-TGCAGCGGCGCGGTCCTGGCGAC-5'; T74A: 5'-CTCGCCACTACAGGCCCCGGCTG-CCC-3' and 5'-GGGCAGCC-GGGGCCTGTAGTGGCGAG-3'; S87A: 5'-GCCTGCGCTC-GCCCCGGTGCCACCTG-3' and 5'-CAGGTGGCACCGGGGCGAGCGCAGGC-3'). Each single point mutation was verified by direct DNA sequencing using D-rhodamine terminator kit (Perkin-Elmer, Wellesley, MA).
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Cell transfection
Exponentially growing immature murine thymoma WEHI 7.1 cells were harvested and viability was determined by trypan blue exclusion. Cells were pelleted by centrifugation at 2500 rpm and washed with 5 ml 1x PBS. Thirty microgram SFFV-Neo (WEHI-Neo), SFFV-Bcl-2 wt plasmid (WEHI-Bcl-2 wt) or mutant (WEHI-Bcl-2 T56A, WEHI-Bcl-2 S70A, WEHI-Bcl-2 T74A, or WEHI-Bcl-2 S87A) suspended in 20 µl ddH2O was added to a 0.4 cm gap cuvette. 1.3 x 107 viable cells were resuspended in 300 µl serum-free DMEM and pipetted into the cuvette; plasmid and WEHI 7.1 cell suspension were mixed by gently tapping the cuvette. Cells were electroporated with Bio-Rad Gene Pulser at 230 V and 960 µF. After electroporation, cells were incubated at room temperature for 5 min, then placed in 15 ml DMEM supplemented with 10% heat-inactivated fetal calf serum (FCS). Transfected WEHI 7.1 cells were cultured for 48 h at 37°C in a 7% CO2 humidified environment in normal medium supplemented with 1 mg/ml G418 to select stably transfected cells. Expression of wild-type Bcl-2 protein and its mutants was confirmed by flow cytometric analysis of immunostained cells and Western blot analysis.
Cell culture
WEHI 7.1 cells were stably transfected with SFFV-Neo (WEHI-Neo), SFFV-Bcl-2 wt (WEHI-Bcl-2 wt), or SFFV-Bcl-2 mutants (WEHI-Bcl-2 T56A, WEHI-Bcl-2 S70A, WEHI-Bcl-2 T74A, and WEHI-Bcl-2 S87A). All mutant-derived cell lines were cultured in DMEM supplemented with 10% heat-inactivated FCS as well as 100 I.U./ml penicillin and 70 I.U./ml streptomycin (27)
. The cell cultures were maintained at a density of 1–5 x 105 cells/ml at 37°C in a 7% CO2 humidified environment and passaged every 48 h. Cultures for experiments were initially suspended at 2 x 105 cells/ml in a total volume of 10 ml and viability was > 96%. At the end of each experiment, cell growth was measured by counting cell density in a hemocytometer. Cell death was quantitated by determining cell viability using trypan blue exclusion method.
Immunostaining and flow cytometry
One million cells (WEHI-Neo, WEHI-Bcl-2 wt, WEHI-Bcl-2 T56A, WEHI-Bcl-2 S70A, WEHI-Bcl-2 T74A, or WEHI-Bcl-2 S87A) were harvested and washed with 1 ml 1x PBS. Cells were resuspended in 250 µl of Cytofix/Cytoperm solution (PharMingen, San Diego, CA) and incubated at 4°C for 20 min. Fixed and permeablized cells were pelleted and washed twice in 1 ml 1x Perm/Wash solution (PharMingen, San Diego, CA), then completely resuspended cells in 50 µl of Perm/Wash solution containing 20 µl of phycoerythrin (PE) -conjugated anti-human Bcl-2 monoclonal antibody (30)
. The cells were incubated at 4°C for 30 min in the dark, then pelleted and washed twice in 1 ml 1x Perm/Wash solution. Washed cells were resuspended in staining buffer (Dulbecco’s PBS without Mg2+ or Ca2+, 1% heat-inactivated FCS, 0.09% (w/v) sodium azide, pH: 7.4
7.6, filtered with 0.2 µm filter) before flow cytometric analysis.
Western blot analysis
2 x 107 cells were harvested and washed with 10 ml ice-cold 1x PBS, then lysed in whole cell lysis buffer (50 mM Tris (pH7.5), 150 mM NaCl, 2 mM EDTA, 1 mM EGTA-1% Triton X-100) (31)
containing Complete, Mini protease inhibitor mixture (Roche, Basel, Switzerland). The cell lysate was sonicated on ice with a W-200 sonicator (Heat system-Ultrasonics, Farmingdale, NY) for 40 s. Samples were diluted in 5x Laemmli gel loading buffer to a final concentration of 50 mM Tris-HCl, pH 6.8, 2% sodium dodecyl sulfate (SDS), 0.1% bromphenol blue, 10% glycerol, and 100 mM dithiothreitol. Samples were heated to 100°C for 10 min in a water bath before electrophoresis. Proteins were separated by 12% SDS-polyacrylamide gel electrophoresis and electroblotted overnight onto a 0.1 µm nitrocellulose membrane (Schleicher & Schuell, Keene, NH). Loading equivalency and transfer efficiency were evaluated by Ponceau S staining. Membranes were blocked in Tris-buffered saline (10 mM Tris-HCl, pH 7.4, 154 mM NaCl, 0.05% Tween 20) containing 10% nonfat dry milk (10% TBS-T) for 1 h. Membranes were washed in Tris-buffered saline containing 1% nonfat milk (1% TBS-T) for 15 min and incubated with a mouse monoclonal anti-human Bcl-2 antibody (DAKO, Glostrup, Denmark) in 1% TBS-T (1:250) for 1 h at room temperature. Membranes were washed in 1% TBS-T for 15 min twice and reacted with a horseradish peroxidase-labeled anti-mouse secondary antibody (Amersham Pharmacia Biotech, Buckinghamshire, U.K.) in 1% TBS-T (1:15,000) for 1 h at room temperature. Chemiluminescent detection was performed with ECL reagents (Amersham Pharmacia Biotech) after washing in 1% TBS-T for 15 min twice. Autoradiography of the membranes were processed using hyperfilm-ECL (Amersham Pharmacia Biotech).
Measurement of mitochondrial membrane potential
Changes in mitochondrial membrane potential were monitored using a dual emission fluochrome, 5,5',6,6'-tetrachloro-1,1', 3,3'-tetraethylbenzimidazole carbocyanine iodine (JC-1) (32)
. JC-1 was loaded in 1 ml of cells at a final concentration of 10 µM and incubated at 37°C in a 7% CO2 humidified environment for 30 min. Cells were examined on a Becton-Dickinson FACSort using CELLQuest software by gating FL-1 (530 nm) vs. FL-2 (585 nm) on a contour plot. JC-1 forms aggregates and emits as red-orange fluorescence at a normal mitochondrial membrane potential. This red-orange fluorescence can be detected in the FL-2 channel. Depolarization results in shifting the dye from aggregates to monomers. The accumulation of green monomers can be observed as an increase in FL-1 fluorescence accompanied by a reduction in FL-2 fluorescence. Formation of monomers (FL-1) was used to assess the changes in mitochondrial membrane potential on a single cell basis.
Analysis of apoptosis by flow cytometry
Flow cytometry was used to assess the mode of cell death (33)
that occurred in response to glucocorticoid administration or FCS depletion. For analysis of DNA integrity, cells were pelleted from each culture after cell counting and fixed by addition of cold 70% ethanol with agitation. Cells were stored at 4°C for at least 24 h. Fixed cells were pelleted and washed with 1x PBS, then stained with 20 µg/ml propidium iodide (PI) containing 1 mg/ml RNase (Qiagen, Valencia, CA) in 1x PBS. Stained cells were examined on a Becton-Dickinson FACSort using CELLQuest software (Becton-Dickinson Immunocytometry System, San Jose, CA). A population of cells (10,000 cells per experimental sample) was analyzed by gating on an PI area vs. width dot plot to exclude cell debris and for doublet discrimination.
Cell size was determined by the changes in forward light scattering properties on a Becton-Dickinson FACSort using CELLQuest software. Cells were excited with a 488 nm argon laser and 10,000 cells of each sample were collected. A gate set on each forward scattered light vs. side scattered light dot plot based on control cells was used to distinguish normal and shrunken populations (34)
.
Statistics
All data from experiments showing viability, percent of shrunken cells, and percent subdiploid DNA were analyzed using Student’s t test assuming unequal variances by comparing dexamethasone treatment group to control group. All statistical analysis was performed in Microsoft Excel 98. Statistical significance was defined as P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A) although WEHI-Bcl-2 T56A and WEHI-Bcl-2 T74A exhibited a small population not expressing Bcl-2. Bcl-2 expression levels vary in different cell lines transfected with Bcl-2. This difference probably reflects the fact that some cells are carrying multiple copies of Bcl-2 cDNA or perhaps a difference in the site of integration. Cells from the same experiment were then used to prepare whole cell lysate for Western blotting. S49 Bcl-2 cells (35)
were used as a positive control for Bcl-2 protein. Western blotting demonstrated that all WEHI 7.1 cells transfected with Bcl-2, including wild-type and phosphomutants, displayed the 26 kDa Bcl-2 protein (Fig. 2B
). Variations in Bcl-2 expression in the different cell lines probably reflect unique sites of Bcl-2 cDNA integration in different cell lines and/or the stability of the mutant proteins.
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Effects of Bcl-2 phosphorylation on inhibiting glucocorticoid-induced loss of viability
Dexamethasone decreased the viability of WEHI-Neo cells (Table 1
). Wild-type SFFV-Bcl-2 was transfected into WEHI 7.1 cells. Dexamethasone treatment did not cause a decrease in viability in WEHI-Bcl-2 wt cells (Table 1)
. The role of Bcl-2 phosphorylation in regulating Bcl-2 function was assessed for each Bcl-2 phosphomutant WEHI 7.1 cell line: WEHI-Bcl-2 T56A, WEHI-Bcl-2 S70A, WEHI-Bcl-2 T74A, and WEHI-Bcl-2 S87A (Fig. 1)
. Mutation of threonine 56 and serine 87 to alanine inhibited the ability of Bcl-2 to block dexamethasone-induced loss of viability (Table 1)
. In contrast, mutation of serine 70 to alanine failed to alter the ability of Bcl-2 to block glucocorticoid-induced cell death (Table 1)
. Mutation of threonine 74 to alanine only partially blocked the ability of Bcl-2 to block glucocorticoid-induced cell death (statistically marginal significance, P=0.055). These data implicate that T56 and S87, two known sites of phosphorylation in Bcl-2, are essential for its anti-apoptotic function against glucocorticoid-induced programmed cell death. T74 may play a minor role in Bcl-2’s ability to inhibit glucocorticoid-induced apoptosis.
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Bcl-2 phosphorylation and mitochondrial depolarization during glucocorticoid-induced cell death
By virtue of its predominant subcellular localization in mitochondria, numerous studies have suggested mitochondria as the primary site of Bcl-2 action. Based on this premise, we wished to see whether alterations in the ability of Bcl-2 that we observed with our phosphomutants were reflected by changes in the state of mitochondrial membrane potential. We used the fluorescent dye JC-1 to accomplish this goal. This lipophilic cationic probe exists as aggregates in normal cells with high mitochondrial membrane potential, whereas mitochondrial depolarization leads to the formation of monomers. The dissipation of JC-1 from aggregates (R1 region in Fig. 3
) to monomers (R2 region in Fig. 3
) indicates mitochondrial depolarization and is detected as a reversible shift in light emission from 585 to 530 nm. The statistical significance between different phosphomutants and the control cell line was determined based on the average of percentage of cells in R2 region from three experiments, though we show only a representative contour plot for each cell line.
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As shown in the left-hand column of panels from control cells (Fig. 3)
, Bcl-2 expression of either wild-type or phosphomutants clearly influenced the state of mitochondrial membrane potential, as judged by JC-1 binding (shown by changes from aggregate to mono-mer ratio except in WEHI Bcl-2 S87A cells). Similar to neomycin-resistant cells, Bcl-2 expressors had two populations of mitochondria: those with the same aggregate to monomer ratio as WEHI-Neo cells and a second state reflecting less monomeric JC-1 dye in the R1 region (Fig. 3)
. When WEHI-Neo cells were treated with dexamethasone, mitochondrial depolarization occurred as reflected by the increased percentage of cells in the R2 region of the plots (Fig. 3)
. The expression of Bcl-2 blocked this depolarization in a manner consistent with the viability data. Mutations of Bcl-2 at threonine 56 and serine 87 eliminated Bcl-2’s ability to block depolarization induced by dexamethasone similar to that found in neomycin-resistant cells, whereas mutations at serine 70 did not significantly change Bcl-2 function (Fig. 3)
. The WEHI-Bcl-2 T74A cells were partially resistant to dexamethasone-induced cell death, yet these cells were still sensitive to dexamethasone-induced mitochondrial depolarization, suggesting that mitochondrial depolarization may not accurately reflect cell death induced by dexamethasone. This further supports the notion that these cells remain glucocorticoid responsive despite being resistant to glucocorticoid-induced cell death.
Bcl-2 expression and cell shrinkage
Since cell shrinkage is a predominant characteristic of programmed cell death, cell size was assessed by flow cytometry in cells cultured in the presence or absence of dexamethasone. Cell shrinkage was measured as a decrease in forward light scatter by flow cytometry (Fig. 4
). Dexamethasone induced a significant increase of the shrunken population in WEHI-Neo cells (Fig. 4)
, whereas cell shrinkage was not induced by dexamethasone treatment in WEHI-Bcl-2 wt cells (Fig. 4)
. Dexamethasone treatment also increased the occurrence of cell shrinkage in WEHI-Bcl-2 T56A, WEHI-Bcl-2 T74A, and WEHI-Bcl-2 S87A cells although to a lesser extent than in WEHI-Neo cells (Fig. 4)
. Mutations at S70 did not abrogate Bcl-2’s protective effect against dexamethasone-induced cell shrinkage (Fig. 4)
whereas mutations in T56, S87, and T74 eliminated the ability of Bcl-2 to block glucocorticoid-induced cell shrinkage (Fig. 4)
. These studies corroborate that T56, T74, and S87 impinge on Bcl-2’s function in inhibiting cell shrinkage and are largely consistent with the changes in mitochondrial membrane potential observed with these mutant forms of Bcl-2.
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Bcl-2 phosphorylation and inhibition of DNA degradation induced by glucocorticoids
DNA degradation has been recognized as an important downstream event that defines the occurrence of apoptosis. We therefore performed flow cytometric analysis of DNA content in cells cultured in the presence or absence of dexamethasone. Under these conditions, a subdiploid peak formation indicates a reduction in propidium iodide fluorescence, which results from DNA fragmentation during apoptosis. Dexamethasone treatment led to loss of DNA content in WEHI-Neo cells (Fig. 5
). Dexamethasone treatment failed to reduce DNA content in WEHI-Bcl-2 wt cells (Fig. 5)
, whereas dexamethasone induced loss of DNA content in the T56, T74, and S87 mutant cell lines (Fig. 5)
. WEHI-Bcl-2 S70A cells did not degrade DNA in response to dexamethasone (Fig. 5)
. These data analyzing DNA degradation in T56A and S87A mutants are consistent with measurement of cell viability by dye exclusion and flow cytometric analysis of cell size and mitochondrial membrane potential, suggesting that T56 and S87 are required for Bcl-2 to be able to inhibit glucocorticoid-induced apoptosis. Moreover, the results examining DNA degradation in T74A mutant are consistent with data from the analysis of mitochondrial membrane potential and cell size. Comparing the measurement of cell viability and flow cytometric analysis of mitochondrial membrane potential, cell size, and DNA content indicates that T74 at least plays a partial role in Bcl-2’s ability to inhibit glucocorticoid-induced apoptosis.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Bcl-2 function is thought to be regulated by changes in its phosphorylation status (12)
. We have developed four Bcl-2 alanine phosphomutants (T56A, S70A, T74A, and S87A) and examined their effects on glucocoticoid-induced apoptosis in WEHI 7.1 cells under conditions similar to our earlier studies with S49 cells (28)
. We show here for the first time that Bcl-2 loses its protective effect against glucocorticoid-induced apoptosis in WEHI-Bcl-2 T56A and WEHI-Bcl-2 S87A cells. T74A is marginally important to the anti-apoptotic effect of Bcl-2. Together, these data suggest that T56, S87, and perhaps T74 are involved in Bcl-2’s anti-apoptotic function.
The loss of cell volume is a fundamental and ubiquitous characteristic of apoptosis and expedites separation of the dying cells from adjacent cells. In this study, we assessed the change in cell size in WEHI 7.1 cells expressing wild-type Bcl-2 or its phosphomutants. Our data show that mutations at T56, T74, and S87 abrogate Bcl-2’s ability to inhibit cell shrinkage induced by glucocorticoid treatment. DNA degradation is one of the nearly universal biochemical changes used to monitor apoptosis. We assessed the DNA degradation in WEHI 7.1 cells expressing wild-type Bcl-2 or its phosphomutants to show that this late apoptotic event, similar to the change in cell size, is dependent on the state of phosphorylation of Bcl-2 at T56, T74, and S87. Since Bcl-2 is located primarily in the outer mitochondrial membrane, Bcl-2 is thought to influence the alteration of mitochondrial membrane potential by interaction with PT pores or by its own channel-forming property (37)
. Therefore, we evaluated the change in mitochondrial membrane potential in our model system to dissect the possible mechanism of Bcl-2’s action in its anti-apoptotic activity. Mitochondrial depolarization and apoptosis are consistently induced by glucocorticoids in WEHI-Neo, WEHI-Bcl-2 T56A, and WEHI-Bcl-2 S87A cells, whereas glucocorticoids do not induce mitochondrial depolarization and apoptosis in WEHI-Bcl-2 wt and WEHI-Bcl-2 S70A cells. These results suggest that Bcl-2 phosphorylation plays a role in regulating glucocorticoid-induced apoptosis through mitochondria. However, additional factors may be critical since the WEHI-Bcl-2 T74A cells displayed a shift in mitochondrial potential, cell shrinkage, and DNA degradation in response to dexamethasone despite being partially resistant to death. An alternative explanation of this phenomenon is that mutation of Bcl-2 on T74 may slow down the process of apoptosis induced by dexamethasone that accounts for the relatively small decrease in viability despite the changes in mitochondria, cell shrinkage, and DNA degradation in response to dexamethasone.
Regulation of Bcl-2 family protein activity by post-translational modifications such as phosphorylation has been widely reported in the literature (1
, 9
, 14)
. Bcl-2 is phosphorylated in response to various treatments, but whether this phosphorylation activates or inactivates the anti-apoptotic function of Bcl-2 may be cell type or stimulation specific. The mechanism of Bcl-2’s action is also disputable. Bcl-2 may act on the mitochondria and affect the movement of ions or molecules required for the induction of apoptosis through its channel-forming property or PT pores. However, Bcl-2 resides not only in mitochondrial membrane, but also in nuclear envelope and endoplasmic reticulum (18
, 38
, 39)
. Alternatively, Bcl-2 can be associated with nuclear matrix (40)
, suggesting its possible role in genomic organization, function, and regulation linked through nuclear architecture. In our model system, we have shown that serine/threonine phosphorylation is responsible for Bcl-2’s anti-apoptotic function in dexamethasone-treated lymphoid cells, although the mechanism is uncertain. Furthermore, multiple kinases including Raf-1 (41)
, PKA (16)
, PKC (1
, 42)
, MAPK (8)
, JNK/SAPK (13)
, CDK6 (5)
, and ASK/JNK (14)
have been demonstrated to be involved in phosphorylating Bcl-2. Our data also demonstrated that multiple phosphorylation sites in Bcl-2 are involved in its ability to inhibit glucocorticoid-induced apoptosis and that S87 plays a predominant role in this action. Since the conformation of a protein can be altered by phosphorylation (43)
, the accessibility of phosphorylation sites in Bcl-2 with which protein kinases can be anticipated to be important during Bcl-2 phosphorylation processes (44
, 45)
. We made these mutants by substituting serine or threonine with alanine, which likely leads to conformational change of Bcl-2 and subsequent alteration of protein kinase accessibility (46)
, although these substitutions can also result in changes in Bcl-2’s function in intracellular localization. Thus, the effect of single point mutation can be a combination of modifying the interaction of protein kinases with Bcl-2 and changes in the phosphorylation status of the mutated site per se.
In summary, we have shown for the first time that threonine 56 and serine 87 are required for Bcl-2 to protect cells from glucocorticoid-induced apoptosis. Threonine 74 may play a partial role, but may not be as important as threonine 56 and serine 87, in Bcl-2’s ability to protect the cells from glucocorticoid-induced apoptosis. These results suggest that Bcl-2 may signal via a pathway requiring phosphorylation to interact with glucocorticoids. However, the enzymes responsible for Bcl-2 phosphorylation and the players of Bcl-2-glucocorticoid interaction still need to be elucidated.
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ACKNOWLEDGMENTS |
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Received for publication November 26, 2001.
Revision received February 14, 2002.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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