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Vol. 16, Issue 9, 4437-4453, September 2005
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B-Crystallin through Inhibition of RAS Activation
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* Hormel Institute, University of Minnesota, Austin, MN 55912;
Department of Molecular Biology, University of Medicine and Dentistry of New Jersey, Stratford, NJ 08084;
College of Life Sciences, Hunan Normal University, Changsha, China 410081; and
# The Burnham Institute, La Jolla, CA 92037
Submitted January 7, 2005;
Revised May 16, 2005;
Accepted June 27, 2005
Monitoring Editor: Carl-Henrik Heldin
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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B-crystallin, a member of the small heat-shock protein family, blocks activation of RAS to inhibit ERK1/2 activation, and greatly attenuates calcimycin-induced apoptosis. Together, our results provide 1) a partial explanation for the lack of spontaneous tumor in the lens, 2) a novel signaling pathway for calcium-induced apoptosis, and 3) a novel antiapoptotic mechanism for B-crystallin. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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). It either occurs as a normal physiological process during animal development (Raff, 1992) or it can be triggered by a number of external signals such as hormone (Wyllie, 1980), 
-irradiation (Strasser et al., 1991), withdrawal of growth factors (Williams et al., 1990) and stress (Li et al., 1995a). Under normal physiological conditions, apoptosis helps to remove excess or unwanted cells or tissues such as vertebrate neurons (Cowan et al., 1984), and larval tissue during metamorphosis (Kerr et al., 1974). In contrast, induced apoptosis can have serious pathological consequences (Carson and Ribeiro, 1993; Thompson, 1995). For example, the bacterial pathogen Shigella flexneri can induce apoptotic death of macrophages, potentially leading to human dysentery (Zychlinsky et al., 1992). Human immunodeficiency virus-1 induces apoptosis of CD4+ T lymphocytes, contributing to development of AIDS (Meyaard et al., 1992).
The lens of the vertebrate eye is a unique organ in that it is nonvascularized and noninnervated (Bloemendal, 1981). It contains only a single layer of epithelial cells that remain quiescent in the central section, divide toward the equatorial area and terminally differentiate into fiber cells in the equatorial region (McAvoy, 1980; Piatigorsky, 1981). This single layer of lens epithelial cells is essential for maintaining the metabolic homeostasis and transparency of the entire lens (Spector, 1995). Induced apoptosis of these lens epithelial cells by various stress factors likely initiates noncongenital cataractogenesis (Li et al., 1995a,b; Li and Spector, 1996; Li, 1997). Another unique feature of the ocular lens is that it never develops spontaneous tumor (Veromann, 1994). The molecular mechanism underlining this phenomenon is largely unknown.
Both biochemical and genetic studies have shown that apoptosis is mediated by different signaling components and controlled by various regulators (Yuan, 1997; Wang, 2001). Among these signaling molecules and apoptotic regulators, the mitogen-activated protein kinases (MAPKs) (Franklin and McCubrey, 2000; Kyriakis and Avruch, 2001; Ballif and Blenis, 2001; Weston and Davis, 2002) and the heat-shock proteins play an important role in regulating stress-induced apoptosis (Xanthoudakis and Nicholson, 2000).
MAPKs participate in diverse cellular functions such as cell proliferation, cell differentiation, and cell death (Kyriakis and Avruch, 2001; Pearson et al., 2001; Weston and Davis, 2002). The importance of MAPK signaling pathways in regulating apoptosis under stress conditions has been widely investigated (Xia et al., 1995; Dickens et al., 1997; Yang et al., 1997; Nemoto et al., 1998; Wang et al., 1998; Zechner et al., 1998; Behrens et al., 1999; Bonni et al., 1999; Hayakawa et al., 1999; Tournier et al., 2000; Liu et al., 2004). Much of the work has supported the general view that among the three types of MAPKs, the ERK1/2-mediated pathway plays an essential role in promoting cell cycle progression and are generally involved in proliferative signaling and cell survival (Xia et al., 1995; Wang and Reed, 1998; Blalock et al., 2000; Hoyle et al., 2000). On the other hand, c-Jun-NH2-terminal kinases (JNKs) and p38 kinase, activated by extracellular stress signals, are involved in apoptosis (Xia et al., 1995; Yang et al., 1997; Nemoto et al., 1998; Wang et al., 1998; Zechner et al., 1998; Behrens et al., 1999; Hayakawa et al., 1999; Tournier et al., 2000). The survival function of ERK1/2 is further demonstrated by the fact that inhibition of ERK signaling leads to increased sensitivity of ovarian cancer cell lines to cisplatin-induced apoptosis (Hayakawa et al., 1999; Persons et al., 1999). In contrast to these previous studies, we demonstrate here that activation of RAF/MEK/ERK signaling pathway alone mediates calcimycin-induced apoptosis of lens epithelial cells. Calcimycin also activates both p38 kinase and JNK2. However, activation of these two kinases does not seem to be involved in calcimycin-induced apoptosis. Downstream of ERK activation, p53 plays a critical role. It positively regulates expression of the proapoptotic factor Bax (Oltvai et al., 1993) and promotes the mitochondrial death pathway. Thus, our results provide a novel signaling pathway for calcium-triggered apoptosis. Our finding that the stress-activated RAF/MEK/ERK signaling pathway, instead of promoting survival, mediates stress-induced apoptosis also provides a partial explanation for the absence of the spontaneous tumor in the ocular lens.
The heat-shock proteins are important regulators against apoptosis (Xanthoudakis and Nicholson, 2000). These factors seem to protect cells from induction of apoptosis through different mechanisms. Hsp90 seems to inhibit apoptosome formation (Pandey et al., 2000a). Hsp70 can interact with BAG-1 (Takayama et al., 1997) and inhibit apoptosis by preventing recruitment of procaspase-9 to the Apaf-1 apoptosome complex (Beere et al., 2000). Hsp70 is also able to function downstream of caspase activation (Jaattela et al., 1998). Hsp27 prevents cell death at different signaling steps. First, it interacts with cytochrome c to prevent activation of procaspase-9 (Garrido et al., 1999; Bruey et al., 2000). It also binds to caspase-3 and modulates the activity of caspase-3 (Pandey et al., 2000b). Finally, a recent study demonstrates that Hsp27 can abrogate the apoptotic pathway through regulation of Bid intracellular distribution and protection of F-actin integrity (Paul et al., 2002). Both Hsp27 and -crystallins are closely related family members (Ingolia and Craig, 1982). Compared with Hsp27, the antiapoptotic mechanisms for -crystallins remain largely unknown until our recent study in which we showed that B-crystallin is able to interact with the precursors of caspase-3 to repress its activation (Mao et al., 2001). Similar results are reported from another laboratory (Kamradt et al., 2001). More recently, we also have demonstrated that -crystallin is able to sequester the translocation of Bax and Bcl-XS, proapoptotic members of the Bcl-2 family, from cytoplasm into mitochondria to prevent staurosporin-induced apoptosis (Mao et al., 2004). To further explore the antiapoptotic mechanism of -crystallin, we have examined its ability to block calcimycin-induced apoptosis. Here, we demonstrate that B-crystallin can repress calcimycin-induced activation of RAF/MEK/ERK pathway. By blocking RAS activation by RAS-GRF2, B-crystallin suppresses the RAF/MEK/ERK pathway and abrogates calcimycin-induced apoptosis. Therefore, our results also provide a novel antiapoptotic mechanism for B-crystallin.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Cell Culture
The rabbit lens epithelial cells (RLECs), N/N1003A, were grown in Eagle's minimum essential medium containing 10% rabbit serum as we described previously (Mao et al., 2001).
Expression Constructs and Stable Cell Lines
The stable transfected cell lines, pEGFP-N/N1003A (expressing only the green fluorescence protein, GFP, from the vector), and pEGFP-mB-N/N1003A (expressing the fusion protein of GFP and mouse B-crystallin) were established and maintained as described previously (Li et al., 2001). The stable cell lines carrying either the p53 expression construct pCMV-p53 or its corresponding vector pCMV-neo were described previously (Miyashita and Reed, 1995). The antisense-p53 construct was created by first subcloning the HindIII-EcoRI fragment of p53 into pBluescript SK(+/-) vector and then isolate the EcoRI-SalI fragment and ligated back into pCMV-Neo vector. The mouse Bax cDNA was kindly provided by Dr. Stanley Korsmeyer (Oltvai et al., 1993). This cDNA was subcloned from pBluescript-SK into pCI-neo vector at the EcoRI site. The antisense-Bax construct was created with opposite insertion of the cDNA into pCI vector. The RAF and RAS dominant negative mutants (Rosen et al., 1994; Mischak et al., 1996) were obtained from the BD Biosciences Clontech. The ERK dominant negative mutant was described previously (Huang et al., 1999). These constructs were amplified in DH-5 and purified by two rounds of CsCl ultracentrifugation (Ausubel et al., 2002) or by Plasmid Maxiprep kit (catalog no. 732-6130; Bio-Rad, Hercules, CA). Transfection of N/N1003A cells was performed using electroporation with a BTX Electro Cell manipulator as we described previously (Xiang et al., 2002; Wang et al., 2005) or with Lipofectamine 2000 from the Invitrogen (Feng et al., 2004). The transfected cells were then subjected to neomycin (G418) selection for 4-6 wk before stable lines of individual clones were obtained.
Pretreatment or Treatment of Various Types of Cells by Dimethyl Sulfoxide (DMSO), Calcimycin, UO126, SP600125, PD169316, and Cyclosporine A
The parental N/N1003A and various transfected cells (Table 1) were grown to 100% confluence in minimal essential medium (MEM) containing 10% rabbit serum in the absence (for N/N1003A) or presence (for various transfected cell lines) of 400 µg/ml G418. Then, MEM containing 10% serum plus 0.01% DMSO (mock) or 5 µM calcimycin (experiment) was used to replace the culture media for the required period of incubation (as indicated in the figure legends and Table 1). For pretreatment, the MEM containing 10% serum plus 0.01% DMSO (mock), 50 µM UO126 (MEK1/2 inhibitor), 1 µM SP600125 (JNK1/2 inhibitor), 2 µM PD169316 (p38 inhibitor), or 250 nM cyclosporine A (PP-2B inhibitor) were used to replace the culture media, for a total of 6 h of incubation before subjected to treatment by 0.01% DMSO (mock) or 5 µM calcimycin (experiment) for the required time. After treatment, all samples were collected for analysis of apoptosis, protein and enzyme activity, or reporter gene activity.
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Cell Flow Cytometry Analysis
The percentage of apoptotic cells was determined by cell flow cytometry, which was conducted as previously described (Ormerod, 1994) using the Annexin V-FITC apoptosis detection kit from MBL International (Watertown, MA). Briefly, cells were harvested with 0.025% trypsin/5 mM EDTA in phosphate-buffered saline (PBS), and 2.5% fetal bovine serum in PBS was added to the cells as soon as they were released from the culture dishes. Then, the cells were transferred to a centrifuge tube, washed with PBS, and incubated with Annexin V-FITC plus propidium iodide (PI) following the protocol included in the kit. Cells were analyzed on a BD Biosciences FACS Calibur flow cytometer (Beckman Coulter, Fullerton, CA), with the fluorescein isothiocyanate (FITC) signal in FL1 and the PI signal in FL2. Intact cells were gated in the FSC/SSC plot to exclude small debris. Cells in the lower right quadrant of the FL1/FL2 dot plot (labeled with annexin-FITC only) are considered to be in early apoptosis, and cells in the upper right quadrant (labeled with annexin-FITC and PI) are in late apoptosis. The quantitative results of apoptosis presented were averaged from three independent experiments. The SD was shown in each figure and Table 1.
DNA Fragmentation
The apoptotic nature of cell death was determined using Hoechst staining and DNA fragmentation as described previously (Li et al., 1998, 2001; Mao et al., 2001, 2004; Huang et al., 2005).
Membrane Protein Cross-linking
pEGFP-N/N1003A and pEGFP-mB-N/N1003A cells were grown to 100% confluence and then treated with 5 µM calcimycin (experiment) or 0.01% DMSO (control) for 10 min and then further incubated in PBS with 5 µM calcimycin (experiment) or 0.01% DMSO (control) plus 2 mM dimethyl-3,3'-dithiobispropionimidate(2HCl) (the cross-linking agent; Pierce Chemical, Rockford, IL) for 30 min. After washing four times in PBS, cells were harvested in hypotonic buffer (50 mM HEPES, pH 7.4, 1 mM CaCl2, 1 mM phenylmethylsulfonyl fluoride, and 1x protease inhibitor cocktail) for extraction of both membrane and cytosol proteins as described previously (Brennan and Lin, 1996). Both membrane and cytosol proteins were used for Western blot as described below.
Isolation of Mitochondria
After treatment with 5 µM calcimycin (experiment), or 0.01% DMSO (control), N/N1003A (with or without UO126-pretreatment) and various transfected cells were harvested for isolation of cytosol and mitochondria as previously described previously (Eskes et al., 2000; Mao et al., 2004). The proteins from both mitochondria and cytosol were subjected to Western blot analysis as described below.
Protein Preparation and Western Blot Analysis
The total proteins were prepared from N/N1003A or various transfected cells as described previously (Mao et al., 2001 and 2004). Membrane proteins were isolated as described previously (Brennan and Lin, 1996). Western blot analysis was conducted as described previously (Li et al., 2001, 2003). Various antibodies and their concentrations used for Western blot analysis were described previously (Li et al., 2003; Mao et al., 2004; Feng et al., 2004; Wang et al., 2005).
GTP Loading Assays
GTP loading assays were conducted according to Rosen et al. (1994). pEGFP-N/N1003A and pEGFP-mB-N/N1003A cells were metabolically labeled in PO4-free DMEM (DMEM) in the presence of 0.5 mCi/ml [32P]orthophosphate for 4 h. Then, 0.01% DMSO or 5 µM calcimycin was added into these labeling cells and incubated for 30 min. After stimulation, the cells in each dish were washed once with cold PBS, and lysed on the plate with 250 µl of lysis buffer (20 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM MgCl2, 1% Triton X-100) containing 2 µg of anti-RAS monoclonal antibody (Oncogene Science, Cambridge, MA). Extracts were scraped into Eppendorf tubes and drawn 15 times through a 0.22-gauge needle, and 1:10 volume of PBS/1% bovine serum albumin/10% charcoal (Sigma) slurry was added. Extracts were rocked at 4°C for 1 h and centrifuged 5 min at 12,000 x g, and 200 µl of the supernatants was mixed with 40 µl of proteins A/G agarose. The mixture was rocked at 4°C for another hour and then washed three times with lysis buffer and once with PBS. Pellets were resuspended in 40 µl of 1 M KH2PO4, pH 3.4, and incubated at 85°C for 3 min. Samples were centrifuged at 12,000 x g for 5 min, and 20 µl of supernatant was decanted to a new tube and stored at -20°C overnight. Samples were centrifuged again the next day, and 10 µl was decanted for spotting, 1 µl at a time, onto 20 x 20-cm PEI-cellulose thin layer chromatography plates (sigma). Plates were developed in 1 M KH2PO4, pH 3.4, for 2 h.
Assay of Caspase-3 Activity
The caspase-3 activity in various cell lines with or without pretreatment followed by treatment with calcimycin was assayed as we described previously (Li et al., 2001; Mao et al., 2001, 2004).
Analysis of Transient Gene Expression
For reporter gene activity, the construct of chloramphenicol acetyltransfeRASe (CAT) reporter gene driven by either the human bax gene promoter or the same promoter with the p53 sites mutated (Miyashita and Reed, 1995), together with the control construct expressing 
-galactosidase (Xiang et al., 2002), was introduced into pCMV vector-, pCMV-p53- and pCMV-antisense-p53-transfected cells as described above. The transfected cells were grown in 100-mm culture dishes for 24 h and then treated with either 0.01% DMSO or 5 µM calcimycin for additional 6 h before harvested for assays of -galactosidase and CAT activities as described previously (Ausubel et al., 2002; Xiang et al., 2002).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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,b; Li and Spector, 1996). To explore the signaling pathways for stress-induced apoptosis, we studied calcium-induced apoptosis in rabbit lens epithelial cells (RLECs), N/N1003A. Calcimycin, a calcium mobilizer, is known to induce calcium influx and apoptosis of different types of cells (McConkey et al., 1992; Orrenius et al., 2003). As shown in Figure 1, A and D, cell flow cytometry analysis revealed that treatment of N/N1003A cells with 5 µM calcimycin induced time-dependent apoptosis of N/N1003A cells. The apoptotic nature was verified by DNA fragmentation assay (Figure 1B, lane 3). N/N1003A cells treated with 0.01% DMSO (a solvent for calcimycin) displayed very few apoptotic cells (our unpublished data).
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), we next examined whether these kinases are involved in calcimycin-induced apoptosis. As shown in Figure 1F, left, treatment of N/N1003A cells with calcimycin induces steady activation of ERK1/2 within the first 5 h of treatment, and by 10 h of treatment, the activation slightly decreased. To demonstrate that activation of ERK1/2 was necessary for calcimycin-induced apoptosis, N/N1003A cells were pretreated with 50 µM UO126, an inhibitor of MEK1/2 kinases, for 6 h followed by treatment with 5 µM calcimycin for 0-36 h. As shown in Figure 1F, right, pretreatment with UO126 completely blocked activation of ERK1/2. When ERK1/2 activation was blocked, the calcimycin-induced apoptosis was substantially blocked (Figure 1, A and E, and Table 1, rows 1 and 2). This inhibition of calcimycin-induced apoptosis was further verified by DNA fragmentation assay (Figure 1B, lane 4). As control, N/N1003A cells pretreated with 50 µM UO126 followed by treatment with 0.01% DMSO displayed very few apoptotic cells (Figure 1A, B, lane 2, and C). Together, these results indicate that activation of ERK1/2 is necessary for calcimycin-induced apoptosis.
To further confirm that activation of ERK1/2 is the key event in calcimycin-induced apoptosis, we next introduced a dominant negative mutant ERK2 (Huang et al., 1999) into N/N1003A cells. Western blot analysis of the total ERK1/2 proteins and their activities from either ERK mutant or vector-transfected cells revealed that the dominant negative mutant clearly interfered with calcimycin-induced activation of ERK1/2 (Figure 1G). Cell flow cytometry analysis demonstrated that inhibition of ERK activity by the dominant negative mutant substantially blocked calcimycin-induced apoptosis (Table 1, row 8). In contrast, the vector-transfected cells underwent similar apoptosis as parent cells did after calcimycin treatment (Table 1, row 5). The cells treated with 0.01% DMSO displayed only background apoptosis (our unpublished data). These results further confirm that activation of ERK1/2 is necessary in mediating calcimycin-induced apoptosis.
Calcimycin-induced Activation of MEK1/2 and RAF1 but Not p38 and JNK MAP Kinases Is Necessary for Calcimycin-induced Apoptosis
The UO126 inhibition experiments described above (Figure 1F) indicated that MEK1/2 activities were necessary for ERK1/2 activation and thus critical for calcimycin-induced apoptosis. Next, we determined the activation of MEK1/2 by calcimycin and also the importance of such activation in calcimycin-induced apoptosis. As shown in Figure 2A, left, calcimycin induced steady activation of MEK1/2. Inhibition of MEK1/2 activation by a dominant negative mutant RAF1 (Mischak et al., 1996) (Figure 2A, right) substantially suppressed calcimycin-induced apoptosis (Table 1, row 7). Next, we examined the role of RAF activation in calcimycin-induced apoptosis. The RAF kinase is normally activated by the small GTP binding protein, RAS (Hingorani and Tuveson, 2003). Under calcimycin treatment, RAF1 was steadily activated within the first 10 h (Figure 2B, left). Inhibition of RAF1 activation by a dominant negative mutant RAS (Rosen et al., 1994) (Figure 2B, right) also greatly suppressed calcimycin-induced apoptosis (Table 1, row 6). Together, our results showed that activation of RAF/MEK/ERK signaling pathway is necessary for calcimycin-induced apoptosis.
Because an early study (Lotem et al., 1999) revealed that activation of p38 kinase was necessary for calcimycin-induced apoptosis of mouse myeloid leukemic cells, we next examined the possible involvement of p38 kinase activation in calcimycin-induced apoptosis of N/N1003A cells. As shown in Figure 2C, left, calcimycin induced a steady activation of p38 kinase within the first 10 h of treatment. However, inhibition of p38 kinase activation by 2 µM PD169316 (Figure 2C, right) displayed little effects on calcimycin-induced apoptosis (Table 1, row 3). Calcimycin also induced a transient activation of JNK2 after 1-h treatment (Figure 2D, left). Inhibition of JNK2 activation by the pharmacological drug SP600125 (Figure 2D, right) did not interfere with calcimycin-induced apoptosis (Table 1, row 4). Together, these data demonstrated that calcimycin-induced RAF/MEK/ERK pathway alone mediates calcium-induced apoptosis in RLECs.
The Tumor Suppressor p53 Is the Key Regulator Downstream of ERK1/2 Activation during Calcimycin-induced Apoptosis
Previous studies have shown that p53 is an important regulator of apoptosis in the ocular lens. Knockout of p53 (-/-) completely suppresses the Rb-deficiency-induced apoptosis in mouse lens (Morgenbesser et al., 1994). To explore the role of p53 in calcimycin-induced apoptosis, we first examined the possible change in p53 expression after calcimycin treatment. As shown in Figure 3A, left, Western blot analysis revealed the steady up-regulation of p53 protein after treatment with 5 µM calcimycin for 5-10 h. Inhibition of ERK1/2 activation by UO126 almost completely abrogated calcimycin-induced up-regulation of p53 although UO126 pretreatment increased the background p53 expression (Figure 3A, right). These results suggest that downstream of ERK1/2 activation, p53 may be necessary for calcimycin-induced apoptosis. To further test the role of p53 in calcimycin-induced apoptosis, we have either overexpressed p53 through the expression construct, pCMV-p53 or inhibited p53 expression using an antisense construct, pCMV-antisense-p53 (Figure 3B). The cells overexpressing p53 were more sensitive to calcimycin-induced apoptosis (Table 1, row 9). In contrast, cells with p53 expression repressed are much more resistant to calcimycin-induced apoptosis (Table 1, row 10). Thus, the presence or absence of p53 is directly linked to the level of calcimycin-induced apoptosis.
The Tumor Suppressor p53 positively Regulates Expression of Bax to Deliver the Calcimycin-induced Death Signal
To explore how p53 may regulate calcimycin-induced apoptosis, we next examined the expression of Bax, a downstream target gene of p53 (Miyashita and Reed, 1995). As shown in Figure 3C, Bax expression in N/N1003A cells is directly associated with p53 level. Calcimycin treatment of N/N1003A cells induced up-regulation of both p53 and Bax (Figure 3C, lane 1). Overexpression of p53 enhanced Bax expression in the absence and presence of calcimycin treatment (Figure 3C, lane 2). On the other hand, inhibition of p53 expression greatly attenuated Bax expression without or with calcimycin treatment (Figure 3C, lane 3). To confirm that p53 directly regulates Bax in N/N1003A cells, we next introduced a CAT reporter gene driven by a Bax promoter which contains one perfect p53 binding site and three imperfect p53 binding sites or by the same expression construct with the p53 binding site mutated (Miyashita and Reed, 1995) into N/N1003 Cells. After transfection, the transfected cells were grown in normal culture medium for 24 h followed by treatment with 0.01% DMSO or 5 µM calcimycin for 6 h. At the end of treatment, both 0.01%dimethyl sulfoxide and 5 µM calcimycin-treated Bax-CAT-transfected cells were harvested for examination of reporter gene activity. As shown in column 1 of Figure 3D, compared with DMSO, calcimycin induced more than threefold increase in CAT activity from Bax-CAT-transfected cells. In contrast, such induction was not observed in the cells transfected by the same construct with the p53 binding sites mutated (Figure 3D, column 5). These results indicate that p53 can directly regulate Bax in N/N1003A cells. Overexpression of p53 induced a six- and ninefold CAT activity in the absence or presence of calcimycin treatment (Figure 3D, column 3). In contrast, inhibition of p53 expression blocked the calcimycin-induced CAT activity (Figure 3D, column 4). The CAT activity generated from the reporter construct with p53 binding site mutated was close to background and displayed little difference in different cell lines with or without calcimycin treatment (Figure 3D, columns 5-8). To determine that Bax up-regulation played an important role in calcimycin-induced apoptosis, we next overexpressed mouse Bax in N/N1003A cells. As shown in the row 11 of Table 1, cells overexpressing Bax, as those overexpressing p53, were more sensitive to calcimycin-induced apoptosis than the vector-transfected cells. Inhibition of Bax expression with antisense-bax RNA also attenuated calcimycin-induced apoptosis (Table 1, row 12). Together, these results demonstrate that Bax is an important downstream target of p53 during calcimycin-induced apoptosis.
Calcimycin-induced Release of Cytochrome c and Activation of Caspase-3 Can Be Substantially Blocked by the ERK Inhibitor UO126
Because Bax promotes apoptosis through interactions with antiapoptotic regulators of the Bcl-2 family in mitochondria (Oltvai et al., 1993; Gross et al., 1999) and induces release of cytochrome c (Eskes et al., 1998; Jürgensmeier et al., 1998; Pastorino et al., 1999), we predicted that p53-regulated Bax expression would alter the permeability of mitochondria in the calcimycin-treated cells and thus allow release of cytochrome c. To test this possibility, we isolated cytosol and mitochondria from both DMSO and calcimycin-treated cells without or with UO126 pretreatment. As shown in Figure 3E, Western blot analysis revealed that calcimycin treatment indeed led to leakage of mitochondria because a substantial amount of cytochrome c was released from the mitochondria to cytoplasm in calcimycin-treated cells but not in DMSO-treated cells (Figure 3E, left). Pretreatment with UO126 greatly attenuated this release of cytochrome c (Figure 3E, right). As a result of differential release of cytochrome c in RLECs without or with pretreatment of UO126, differential caspase-3 activation was observed. As shown in Figure 3F, caspase-3 activity was up-regulated approximately ninefold by calcimycin in N/N1003A cells. In contrast, a <3-fold up-regulation of caspase-3 activity was observed in UO126-pretreated N/N1003A cells after calcimycin treatment. Thus, UO126 prevention of ERK1/2 activation induced by calcimycin also repressed the downstream apoptotic events.
Calcimycin-induced Apoptosis in N/N1003A Cells Is Greatly Attenuated in Cells Expressing the Exogenous Mouse B-Crystallin
During calcimycin-induced apoptosis, we observed distinct down-regulation of B-crystallin (Figure 4A). Because B-crystallin is an antiapoptotic regulator (Li et al., 2001; Mao et al. 2001, 2004), we predicted that overexpression of B-crystallin may prevent calcimycin-induced apoptosis. As shown in Figure 4B, overexpression of the fusion protein of GFP and mouse B-crystallin in N/N1003A cells provided resistance to calcimycin-induced degradation of both the endogenous B-crystallin, and the fusion protein of GFP and mB-crystallin (Figure 4B). When the viability of the vector-transfected cells (pEGFP-N/N1003A) and the mouse B-crystallin-transfected N/N1003A cells (pEGFP-mB-N/N1003A) were compared, it was found that >90% of the vector-transfected cells were dead or undergoing apoptosis after 36-h treatment by 5 µM calcimycin. In contrast, <40% of the mouse B-crystallin expression cells were apoptotic after the same length of treatment by 5 µM calcimycin (Figure 4C). The apoptotic nature of these cells was verified by DNA fragmentation. As shown in the lane 3 of Figure 4D, an 18-h treatment caused distinct DNA fragmentation laddering in the vector-transfected N/N1003A cells. However, DNA fragmentation ladder was hardly detectable at this stage from the B-crystallin expression cells (Figure 4D, lane 5 from the left). Hoechst staining further confirmed the above-mentioned results (our unpublished data). Treatment of either vector-transfected cells or mouse B-transfected cells with 0.01% DMSO induced very few apoptotic cells (our unpublished data).
B-Crystallin Suppresses Calcimycin-induced Activation of ERK1/2
Next, we investigated the possible mechanism by which B-crystallin prevents calcimycin-induced apoptosis. Knowing that inhibition of ERK1/2 leads to blockage of calcimycin-induced apoptosis, we first tested whether B-crystallin could abrogate calcimycin-induced activation of ERK1/2. As shown in Figure 5A, left, Western blot analysis revealed that although calcimycin also induced steady activation of ERK1/2 in vector-transfected cells as in the parent N/N1003A cells, calcimycin-induced ERK1/2 activation was not observed in pEGFP-mB-N/N1003A cell (Figure 5A, right two panels). These results indicate that B-crystallin prevent calcimycin-induced apoptosis through suppression of calcimycin-induced activation of ERK1/2.
B-Crystallin Blocks RAS-GRF2-mediated RAS Activation to Suppress ERK Activation
The fact that in B-crystallin expression cells, calcimycin-induced activation of ERK1/2 (Figure 5A), p38 and JNK2 (our unpublished data) was all inhibited indicates that B-crystallin may acts on the initial steps of RAS/RAF/MEK/ERK signaling pathway. For this reason, we investigated the effect of B-crystallin on RAS activation. Both pEGFP-N/N1003A and pEGFP-mB-N/N1003A cells were label with [32P]orthophosphate and then treated with DMSO and 5 µM calcimycin. Cell extracts were prepared from labeled and treated cells, and the RAS protein in each sample was immunoprecipitated. After that, the levels of GTP-bound RAS and GDP-bound RAS were determined with TLC. As shown in Figure 5B, DMSO treatment of both vector and B-crystallin expression cells only induced background levels of RAS activation as reflected by the faint spots of labeled GTP released from RAS-GTP. Treatment of the vector-transfected cells with 5 µM calcimycin induced substantially activation of RAS. However, such activation of RAS was not observed in the B-crystallin expression cells. Thus, B-crystallin blocks RAS activation to suppress activation of the downstream MAP kinases after treatment by calcimycin.
To explore how B-crystallin may block the activation of RAS, we examined the upstream regulators of RAS. The RAS small GTPases are normally activated by the quanine nucleotide exchange factors (GEFs), which induce the dissociation of GDP to allow association of GTP (Cullen and Lockyer, 2002). On response to growth factor stimulation, two common RAS GEFs, son-of-sevenless 1 (Sos 1) and 2 (Sos 2), are activated to transfer from cytosol to the plasma membrane, and there they activate RAS (Campbell et al., 1998). However, under calcium treatment, RAS is activated by other factors, the RAS-guanine-nucleotide-releasing factor 1 (RAS-GRF1) and 2 (RAS-GRF2) (Cullen and Lockyer, 2002). Because RAS-GRF1 is mainly expressed in the nerve system and RAS-GRF2 is widely expressed in many tissues (Farnsworth et al., 1995; Fam et al., 1997), we therefore examined the activation of RAS by RAS-GRF2 after calcimycin treatment. As shown in Figure 6A, left, after calcimycin treatment, the 135-kDa RAS-GRF2 was transferred from cytosol (CS) to plasma membrane (MF). In the B-crystallin expression cells, however, the translocation of the RAS-GRF2 from cytosol to plasma membrane was largely inhibited (Figure 6A, right). To further confirm that the RAS-GRF2 translocated to plasma membrane activated RAS, we performed in vivo cross-linking after calcimycin treatment as described in the Materials and Methods. If the RAS-GRF2 indeed activated RAS after calcimycin treatment, it would bind to RAS and thus could be cross-linked together to form a complex. As showed in Figure 6B, left, calcimycin treatment induced formation of the RAS-GRF2 and RAS complex in the vector-transfected N/N1003A cells as demonstrated by the Western blot analysis. In contrast, in the B-crystallin expression cells, RAS-GRF2 activation of RAS was largely inhibited as reflected by the fact that majority of RAS and RAS-GRF2 were not coupled together (Figure 6B, right). These results demonstrated that in N/N1003A cells, calcimycin activated RAS mainly through RAS-GRF2, and B-crystallin negatively regulates activation of RAS through suppression of RAS-GRF2 translocation.
B-crystallin Prevents Calcimycin-induced Up-Regulation of p53 and Bax, Release of cytochrome c, and Activation of Caspase-3
As described above, inhibition of ERK1/2 activation by UO126 prevents p53 up-regulation (Figure 4A). In B-crystallin-transfected N/N1003A cells, calcimycin-induced p53 up-regulation was completely abrogated (Figure 7A, right and top). Moreover, calcimycin-induced Bax up-regulation was also suppressed (Figure 7A, right and middle). Thus, by repressing activation of ERK1/2, B-crystallin also suppressed calcimycin-induced up-regulation of p53 and Bax.
Because B-crystallin suppressed calcimycin-induced upregulation of p53 and Bax, we predicted that B-crystallin might preserve the integrity of mitochondria and thus prevent release of cytochrome c. To test this possibility, again, we isolated cytosol and mitochondria from both DMSO and calcimycin-treated vector- and B-crystallin-transfected cells. As shown in Figure 7B, Western blot analysis of the proteins from these different samples revealed that calcimycin treatment led to malfunction of mitochondria in vector-transfected cells and a substantial amount of cytochrome c was released from the mitochondria to cytoplasm in calcimycin-treated cells but only barely detectable in DMSO-treated cells (Figure 7B, left). Expression of B-crystallin greatly attenuated this release of cytochrome c (Figure 7B, right). As a result of differential release of cytochrome c in vector- and B-crystallin-transfected cells, differential caspase-3 activation was observed. As shown in Figure 7C, caspase-3 activity was up-regulated >8.5-fold in the vector-transfected N/N1003A cells. In contrast, <2-fold up-regulation of caspase-3 activity was observed in mouse B-crystallin-transfected N/N1003A cells after calcimycin treatment (Figure 7C). Together, through repression of ERK1/2 activation induced by calcimycin, B-crystallin prevents activation of the downstream apoptotic events.
Overexpression of p53 Attenuates the Protective Function of B-Crystallin
To further confirm that p53 is a key component downstream of ERK1/2 activation during calcimycin-induced apoptosis, we overexpressed p53 in B-crystallin-transfected N/N1003A cells. Overexpression of p53 in B-crystallin-transfected N/N1003A cells attenuated the B-crystallin functions in suppressing expression of Bax (Figure 8A, right), preventing release of cytochrome c (Figure 8B, right), and inhibiting of caspase-3 activation (Figure 8C). As a result, overexpression of p53 in B-crystallin-transfected N/N1003A cells enhanced apoptosis (Table 1, row 16).
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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,b, 1992; Asai et al., 1999; Orrenius et al., 2003). After calcimycin treatment, an increase in the intracellular calcium is a necessary event in the induction of apoptosis (McConkey et al., 1992; Orrenius et al., 2003). After calcium increase, several downstream signaling pathways are apparently activated in different types of cells. One pathway activated by calcium to mediate apoptosis is through activation of the protein phosphatase-2B (PP-2B) (Asai et al., 1999; Wang et al., 1999). Activated PP-2B has been reported to dephosphorylate Bad (Wang et al., 1999), an important proapoptotic member of the Bcl-2 family (Yang et al., 1995; Gross et al., 1999). Another pathway seems to be mediated by the p38 kinase. In mouse myeloid leukemic cells, Lotem et al. (1999) have demonstrated that activation of p38 kinase is necessary for calcimycin-induced apoptosis. How activation of p38 kinase signals the downstream apoptotic events is not defined. In the present study, we demonstrate that calcimycin induces activation of ERK1/2 (Figure 1F), p38 kinase, and also JNK2 (Figure 2, C and D). Activation of p38 kinase and JNK2 is not necessary for calcimycin-induced apoptosis in RLECs because inhibition of p38 kinase activation by PD169316 and JNK2 activation by SP600125 does not interfere with calcimycin-induced apoptosis (Table 1), which is apparently different from that observed in mouse myeloid leukemic cells (Lotem et al., 1999). Furthermore, inhibition of PP-2B activation by cyclosporine A does not block calcimycin-induced apoptosis (our unpublished data), suggesting that an alternative pathway is activated. Indeed, when activation of ERK1/2 is blocked by UO126 or a dominant negative mutant ERK, calcimycin-induced apoptosis is mostly abrogated. Moreover, upstream of ERK activation, the ERK activation kinases, MEK1/2 and MEK1/2 activation kinase, RAF1 also were activated after calcimycin treatment. Activation of both MEK1/2 and RAF1 is necessary for calcium-induced apoptosis of lens epithelial cells based on the inhibition studies with dominant negative mutant RAF1. RAF1 is activated by RAS because a dominant negative mutant RAS interfered with RAF1 activation. Thus, our data show that the RAS-activated RAF/MEK/ERK pathway alone mediates calcium-induced apoptosis.
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). Elevation of intracellular calcium through various stimuli also induces activation of the proline-rich tyrosine kinase 2 (Pyk2) (Lev et al., 1995; Dikic et al., 1996). In neuronal and hematopoietic cells, activated Pyk2 binds to Src, which allows the formed complex to interact with Sos, and thus induces the calcium-dependent activation of RAS (Avraham et al., 2000). RAS also can be activated by the RAS-guanine-nucleotide-releasing factor 1 (RAS-GRF1) (Martegani et al., 1992; Shou et al., 1992; Farnsworth et al., 1995) and 2 (RAS-GRF2) (Fam et al., 1997). Although RAS-GRF1 is mainly expressed in the nerve system, our data suggests that in N/N1003A cells, calcimycin treatment leads to RAS activation mainly through RAS-GRF2. Calcimycin not only induces translocation of the RAS-GRF2 from cytosol to plasma membrane but also causes the direct binding of RAS-GRF2 to RAS (Figure 6). Moreover, B-crystallin that substantially blocks calcimycin-induced activation of RAS also suppresses the translocation of RAS-GRF2. Thus, through activation of the calcium-triggered RAS-GRF2, calcium can indirectly activated RAS in rabbit lens epithelial cells.
Once ERK1 and ERK2 are activated, the key downstream event is up-regulation and activation of p53 as evidenced from several observations. First, calcimycin induces tremendous up-regulation of the endogenous p53 protein in N/N1003A cells. Regulation of p53 expression by RAS-activated RAF/MEK/ERK pathway has recently been observed in human fibrosarcoma cell line HT1080 (Agarwal et al., 2001). The importance of p53 in mediating calcimycin-induced apoptosis is further demonstrated by the fact that overexpression of human p53 in N/N1003A cells enhanced the sensitivity of calcimycin-induced apoptosis. In contrast, down-regulation of p53 through antisense technology attenuated calcimycin-induced apoptosis.
10-fold more than that in 
