Caspase 3 is Activated through Caspase 8 instead of Caspase 9 during H<sub>2</sub>O<sub>2</sub>-induced Apoptosis in HeLa Cells
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Abstract:
Oxidative stress is known to be involved in a variety of pathological processes including atherosclerosis, diabetes, and neurodegenerative diseases. Understanding how intracellular signaling pathways respond to oxidative stress will have a significant implication in the therapy of these diseases. In this study, we applied hydrogen peroxide (H2O2) to trigger apoptosis and investigated the dynamic activation of various caspases using a FRET technique. We measured the activation dynamics of caspase 3 and caspase 9 based on two reporter systems, SCAT 3 and SCAT 9. We found that caspase 3 activation was earlier than that of caspase 9 following H2O2 treatment. Caspase 3 was activated rapidly, reaching a maximum in 12±3 min, while the average duration of caspase 9 activation was 21±3 min. When cells were pretreated with Z-LEHD-fmk, a caspase 9 specific inhibitor, caspase 3 activation and apoptosis by H2O2 treatment were little affected, although the caspase 9 activation was completely inhibited. When cells were pretreated with Z-DEVD-fmk, a caspase 3 specific inhibitor, the activation of both caspase 3 and caspase 9, as well as apoptosis, were inhibited. When cells were pretreated with Z-IETD-fmk, a caspase 8 specific inhibitor, the activation of caspase 3 and caspase 9 were significantly delayed. Finally, we found that Bax did not translocate from the cytosol to the mitochondrial membrane during H2O2-induced apoptosis. Our results suggest that, during H H2O2-induced apoptosis, caspase 3 is activated directly through caspase 8 and is not through the mitochondria-dependent caspase 9 activation.Keywords:
Caspase-9
Caspase 8
Caspase 2
Mammalian caspases are a family of cysteine proteases that plays a critical role in apoptosis. We have analyzed caspase-2 processing in human cell lines containing defined mutations in caspase-3 and caspase-9. Here we demonstrate that caspase-2 processing, during cell death induced by UV irradiation, depends both on caspase-9 and caspase-3 activity, while, during TNF-α-dependent apoptosis, capase-2 processing is independent of caspase-9 but still requires caspase-3. In vitro procaspase-2 is the preferred caspase cleaved by caspase-3, while caspase-7 cleaves procaspase-2 with reduced efficiency. We have also demonstrated that caspase-2-mediated apoptosis requires caspase-9 and that cells co-expressing caspase-2 and a dominant negative form of caspase-9 are impaired in activating a normal apoptotic response and release cytochrome c into the cytoplasm. Our findings suggest a role played by caspase-2 as a regulator of the mitochondrial integrity and open questions on the mechanisms responsible for its activation during cell death. Mammalian caspases are a family of cysteine proteases that plays a critical role in apoptosis. We have analyzed caspase-2 processing in human cell lines containing defined mutations in caspase-3 and caspase-9. Here we demonstrate that caspase-2 processing, during cell death induced by UV irradiation, depends both on caspase-9 and caspase-3 activity, while, during TNF-α-dependent apoptosis, capase-2 processing is independent of caspase-9 but still requires caspase-3. In vitro procaspase-2 is the preferred caspase cleaved by caspase-3, while caspase-7 cleaves procaspase-2 with reduced efficiency. We have also demonstrated that caspase-2-mediated apoptosis requires caspase-9 and that cells co-expressing caspase-2 and a dominant negative form of caspase-9 are impaired in activating a normal apoptotic response and release cytochrome c into the cytoplasm. Our findings suggest a role played by caspase-2 as a regulator of the mitochondrial integrity and open questions on the mechanisms responsible for its activation during cell death. phosphate-buffered saline cycloheximide catalytically inactive dominant negative poly(ADP-ribose) polymerase placental alkaline phosphatase tumor necrosis factor fluorescence-activated cell sorting Caspases belong to a conserved family of cysteine proteases playing a critical role in apoptosis and proinflammatory cytokine maturation (1Thornberry N.A. Lazebnik Y. Science. 1998; 281: 1312-1316Crossref PubMed Scopus (6159) Google Scholar). In normal cells, caspases are present as zymogens that are cleaved at aspartic sites during cell death. Enzyme processing generates the active form, which is constituted by a heterodimer consisting of two small and a two large subunits (2Cohen G.M. Biochem. J. 1997; 326: 1-16Crossref PubMed Scopus (4132) Google Scholar). Caspases involved in the apoptotic process can be divided into initiator caspases and effector caspases based on the presence of a large prodomain at their amino terminus region. In general, effector caspases possess short prodomains and are cleaved by initiator caspases, which possess large prodomains. Effector caspases are involved in the cleavage of the death substrates, thus modulating the morphological changes characterizing the apoptotic process (3Salvesen G.S. Dixit V.M. Cell. 1997; 91: 443-446Abstract Full Text Full Text PDF PubMed Scopus (1936) Google Scholar). The long prodomains of the initiator caspases function in inducing dimerization and activation of the proenzymes through interaction with specific adaptor molecules. Caspase-2, caspase-8, caspase-9, and caspase-10 are the long prodomain caspases involved in the apoptotic process (4Earnshaw W.C. Martins L.M. Kaufmann S.H. Annu. Rev. Biochem. 1999; 68: 383-424Crossref PubMed Scopus (2451) Google Scholar). The generation of caspase-deficient mice for initiator caspases has indicated that these enzymes regulate cell death in a tissue- and stimulus-specific fashion (4Earnshaw W.C. Martins L.M. Kaufmann S.H. Annu. Rev. Biochem. 1999; 68: 383-424Crossref PubMed Scopus (2451) Google Scholar). In particular, caspase-9 is the critical player of the apoptotic stimuli acting through mitochondrial dysfunction (5Kuida K. Haydar T.F. Kuan C.Y. Gu Y. Taya C. Karasuyama H. Su M.S. Rakic P. Flavell R.A. Cell. 1998; 94: 325-337Abstract Full Text Full Text PDF PubMed Scopus (1457) Google Scholar, 6Hakem R. Hakem A. Duncan G.S. Henderson J.T. Woo M. Soengas M.S. Elia A. de la Pompa J.L. Kagi D. Khoo W. Potter J. Yoshida R. Kaufman S.A. Lowe S.W. Penninger J.M. Mak T.W. Cell. 1998; 94: 339-352Abstract Full Text Full Text PDF PubMed Scopus (1163) Google Scholar), while caspase-8 is critical for the apoptotic pathways generated by death receptors (7Varfolomeev E.E. Schuchmann M. Luria V. Chiannilkulchai N. Beckmann J.S. Mett I.L. Rebrikov D. Brodianski V.M. Kemper O.C. Kollet O. Lapidot T. Soffer D. Sobe T. Avraham K.B. Goncharov T. Holtmann H. Lonai P. Wallach D. Immunity. 1998; 9: 267-276Abstract Full Text Full Text PDF PubMed Scopus (1032) Google Scholar). Caspase-2-deficient mice showed an apoptotic deficit in the oocytes, following exposure to chemotherapeutic drugs, and in B lymphoblasts following incubation with perforin and granzyme B (8Bergeron L. Perez G.I. Macdonald G. Shi L. Sun Y. Jurisicova A. Varmuza S. Latham K.E. Flaws J.A. Salter J.C. Hara H. Moskowitz M.A. Li E. Greenberg A. Tilly J.L. Yuan J. Genes Dev. 1998; 12: 1304-1314Crossref PubMed Scopus (604) Google Scholar). Additional studies have shown that this initiator caspase is essential for specific apoptotic pathways such as β-amyloid-induced neuron death and salmonella-induced macrophage death (9Troy C.M. Rabacchi S.A. Friedman W.J. Frappier T.F. Brown K. Shelanski M.L. J. Neurosci. 2000; 4: 1386-1392Crossref Google Scholar, 10Jesenberger V. Procyk K.J. Yuan J. Reipert S. Baccarini M. J. Exp. Med. 2000; 192: 1035-1046Crossref PubMed Scopus (151) Google Scholar). The identification of RAIDD/CRADD as an adaptor protein for caspase-2 has suggested that caspase-2 could be involved also in apoptosis triggered by TNF-α. Recruitment of caspase-2 to the TNF-α receptor is regulated by the interaction between the CARD domain present in the prodomain of caspase-2 and a similar CARD domain present in RAIDD/CRADD (11Ahmad M. Srinivasula S.M. Wang L. Talanian R.V. Litwack G. Fernandes-Alnemri T. Alnemri E.S. Cancer Res. 1997; 4: 615-619Google Scholar, 12Duan H. Dixit V.M. Nature. 1997; 385: 86-89Crossref PubMed Scopus (469) Google Scholar). Different apoptotic insults can cause caspase-2 processing, which generally occurs by two proteolytic steps. A first cleavage at aspartic 316 generates two fragments: one of 32–33 kDa containing the prodomain and the large subunit and a second fragment of 14 kDa containing the small subunit. Subsequent cleavages at Asp152 and Asp330 lead to the formation of the large and the small subunits of 18 and 12 kDa, respectively. The appearance of the 32–33-kDa fragment has been generally used as marker of caspase-2 activation (13Li H. Bergeron L. Cryns V. Pasternack M.S. Zhu H. Shi L. Greenberg A. Yuan J. J. Biol. Chem. 1997; 34: 21010-21017Abstract Full Text Full Text PDF Scopus (168) Google Scholar, 14Colussi P.A. Harvey N.L. Kumar S. J. Biol. Chem. 1998; 273: 24535-24542Abstract Full Text Full Text PDF PubMed Scopus (138) Google Scholar, 15Colussi P.A. Harvey N.L. Shearwin-Whyatt L.M. Kumar S. J. Biol. Chem. 1998; 273: 26566-26570Abstract Full Text Full Text PDF PubMed Scopus (72) Google Scholar). Despite significant evidence for an involvement of caspase-2 in different apoptotic pathways, its specific role is not completely understood. Only recently, a caspase-2 substrate has been identified in golgin-160, thus suggesting that caspase-2 could act as an executioner caspase involved in modulating Golgi integrity (16Mancini M. Machamer C.E. Roy S. Nicholson D.W. Thornberry N.A. Casciola-Rosen L.A. Rosen A. J. Cell Biol. 2000; 149: 603-612Crossref PubMed Scopus (325) Google Scholar). In the present study, we have attempted to further analyze caspase-2 activity and its hierarchy with respect to the common apoptotic pathways. We have used human cell lines containing defined mutations in caspase-9 and caspase-3 to unveil the relationships between the mitochondrial pathway and caspase-2 proteolytic processing in response to different apoptotic triggers. In addition, we have analyzed the ability of caspase-2 to induce cell death in cells defective in caspase-9 activity. Rabbits were immunized with His-tagged caspase-2 fragment (residues 15–436) purified from Escherichia coli transformed with the construct pQE32-caspase-2. Briefly, after induction with isopropyl-1-thio-β-d-galactopyranoside, bacteria where collected by centrifugation at 3500 rpm for 5 min and lysed in 50 mm NaH2PO4, pH 8, 300 mm NaCl, 10 mm imidazole, 1 mg/ml lysozyme, 0.5 mm phenylmethylsulfonyl fluoride. After sonication, the insoluble fraction was resuspended in sample buffer (2% SDS, 10% glycerol, 120 mm Tris-HCl, pH 6.8, 0.005% bromphenol blue, 1% β-mercaptoethanol) and then run on an SDS-10% polyacrylamide gel. The gel was stained with Coomassie Blue R-250 to identify the caspase-2 band, which was then electroeluted. The protein was dialyzed overnight in PBS and purified by nickel chromatography using a His-trap column (Qiagen), eluted with an imidazole buffer (50 mmNaH2PO4, pH 8, 300 mm NaCl, 250 mm imidazole), and then used to immunize rabbits. Purified recombinant caspase-2 was cross-linked to an Affi-Prep 10 column (Bio-Rad) and used to affinity-purify antibody to caspase-2 from rabbit antiserum. The serum was loaded onto the column at a slow flow rate, washed with 10 mm Tris-HCl, pH 7.4, and then washed with a high salt buffer (500 mm NaCl, 10 mmTris-HCl, pH 7.4). Caspase-2 antibodies were eluted using 10 mm glycine, pH 2.5, and neutralized with pH 7.5 with Tris-HCl. To generate pcDNA3HA caspase-2 C303G, the entire coding region of human caspase-2 was amplified from pGDSV7S caspase-2 by polymerase chain reaction. In vitro mutagenesis to substitute Cys303 with Gly was performed as previously described (17Brancolini C. Benedetti M. Schneider C. EMBO J. 1995; 14: 5179-5190Crossref PubMed Scopus (240) Google Scholar). The following set of primers was used: primer A (CATGAATTCATGGCCGCTGACAGGGGACGC), primer B (ATCCAGGCCGGCCGTGGAGAT), primer C (ATCTCCACGGCCGGCCTGGAT), and primer D (CATCTCGAGTCATGTGGGAGGGTGTCCTGG). Caspase-2 C303G cDNA was subcloned in pcDNA3HA asEcoRI/EcoRI and EcoRI/XhoI fragments. Caspase-2 C303G was subcloned from pcDNA3HA caspase-2 C303G as SphI/XhoI fragment in pQE32 to generate a His-tagged caspase-2 construct. To generate pEGFPN1Bid and pBSKBid, the entire coding region of human Bid was amplified by polymerase chain reaction from a human fetal cDNA library. The following set of primers was used: primer E (CATGAATTCTGATGGACTGTGAGGTCAACAAC) and primer F (CATGGATCCCGGTCCATCCCATTTCTGGCTAA). Cells were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum, penicillin (100 units/ml), and streptomycin (100 μg/ml). Cells at 70–80% confluence were treated for 36 h with 1 μg/ml cycloheximide plus 20 ng/ml TNF-α or 10 ng/ml TNF-α (MCF-7pBC3 and MCF-7pBC3mut) or with 1 μg/ml cycloheximide (Sigma) alone as a control. For UV treatment, culture medium was removed, dishes were washed once with PBS1 and UVC (180 J/m2)-irradiated, and fresh medium, containing 10% fetal calf serum, was added to the cells (17Brancolini C. Benedetti M. Schneider C. EMBO J. 1995; 14: 5179-5190Crossref PubMed Scopus (240) Google Scholar). Cells were harvested by scraping with a rubber policeman 15 or 18 h later. When required floating cells were collected separately from the adherent cells. After washes in PBS, cells were resuspended in SDS sample buffer sonicated boiled for 3 min and analyzed by Western blot. Transfections were performed by the calcium phosphate precipitation method. Cell were seeded 24 h before transfection and analyzed 20 h after removal of the precipitates. Cells were transfected with 2 μg of each expression vector together with 200 ng of pEGFPN1 (Invitrogen) to identify transfected cells. Caspase-3 was expressed in bacteria and purified as previously described (18Sgorbissa A. Benetti R. Marzinotto S. Schneider C. Brancolini C. J. Cell Sci. 1999; 112: 4475-4482Crossref PubMed Google Scholar) using the pQE-12 expression system (Qiagen). Density scanning of the ∼20-kDa fragments of the autoprocessed caspases, as evidenced after electrophoretic separation and Coomassie Blue staining, was used to estimate the amount of active enzyme. Purified caspase-7 was obtained from Alexis, and caspase-2 was from Chemicom. The different caspases, poly(ADP-ribose) polymerase (PARP) and Bid, were in vitro translated with 35S using the TNT-coupled reticulocyte lysate system (Promega). 1 μl of eachin vitro translated protein was incubated with increasing amounts of caspase-3 or caspase-7 in 15 μl of the appropriate buffer (final volume) for 1 h at 37 °C. Reactions were terminated by adding one volume of SDS gel loading buffer and boiling for 3 min. For Western blotting, proteins were transferred to 0.2-μm pore-sized nitrocellulose (Schleicher & Schuell) using a semidry blotting apparatus (Amersham Pharmacia Biotech) (transfer buffer: 20% methanol, 48 mm Tris, 39 mm glycine, and 0.0375% SDS). After staining with Ponceau S, the nitrocellulose sheets were saturated for 1 h in Blotto-Tween 20 (17) (50 mm Tris-HCl, pH 7.5, 200 mm NaCl, 5% nonfat dry milk, and 0.1% Tween 20) and incubated overnight at room temperature with the specific antibody: anti-caspase-2, anti-actin, and anti-p85 PARP fragment (Promega). Blots were then rinsed three times with Blotto-Tween 20 and incubated with peroxidase-conjugated goat anti-rabbit (Sigma) or goat anti-mouse (Sigma) for 1 h at room temperature. The blots were then washed four times in Blotto-Tween 20, rinsed in phosphate buffer saline, and developed with Super Signal West Pico, as recommended by the vendor (Pierce). For indirect immunofluorescence assays, transfected cells were fixed with 3% paraformaldehyde in PBS for 1 h at room temperature. Fixed cells were washed with PBS and 0.1 m glycine, pH 7.5, and then permeabilized with 0.1% Triton X-100 in PBS for 5 min. The coverslips were treated with anti-cytochrome c (Promega) or with anti-cytochrome c oxidase, diluted in PBS for 1 h in a moist chamber at 37 °C. They were then washed with PBS twice, followed by incubation with tetramethylrhodamine isothiocyanate-conjugated anti-mouse (Sigma) for 30 min at 37 °C. Nuclei were evidenced by Hoechst staining. Cells were examined by epifluorescence with a Zeiss Axiovert 35 microscope or with a LEICA TCS laser scan microscope equipped with a 488 λ argon laser and a 543 λ helium neon laser. We produced an antibody against E. coli expressed caspase-2 fragment in order to study caspase-2 activation during cell death. The antiserum was purified on a caspase-2 affinity column and tested for caspase-2 detection by Western blot. 293 cells were transfected with HA-tagged catalytic inactive caspase-2 as positive control. Anti-caspase-2 antibody recognizes a single band migrating at around 48 kDa, showing a similar electrophoretic mobility to the band detected with the anti-HA antibody in caspase-2-transfected cells. The intensity of the band detected by the anti-caspase-2 antibody was dramatically increased in caspase-2-transfected cells (Fig.1 a). Caspase-2 processing during apoptosis can be followed by the appearance of different proteolytic fragments. A first cleavage at aspartic 316 generates two fragments: a first fragment of 32–33 kDa containing the prodomain and the large subunit and a second fragment of 14 kDa containing the small subunit. Subsequent cleavages at Asp152 and Asp330 generate the large and the small subunits of 18 and 12 kDa, respectively (13Li H. Bergeron L. Cryns V. Pasternack M.S. Zhu H. Shi L. Greenberg A. Yuan J. J. Biol. Chem. 1997; 34: 21010-21017Abstract Full Text Full Text PDF Scopus (168) Google Scholar). To confirm the specificity of the anti-caspase-2 antibody, we investigated caspase-2 processing by Western blot during apoptosis. MDA cells were UV-irradiated, and cellular lysates were prepared 24 h later. In UV-irradiated cells, a band migrating at ∼33 kDa was detected, the appearance of which parallels the rate of cell death in the culture as confirmed by β-catenin processing (19Brancolini C. Sgorbissa A. Schneider C. Cell Death Differ. 1998; 5: 1042-1050Crossref PubMed Scopus (60) Google Scholar). This band is similar in size to the previously identified intermediate processed form of caspase-2, p33. The small and large subunits of caspase-2 were detected only after long exposure of the blot (Fig. 1 b). In conclusion, this evidence indicates that the produced antibody specifically recognizes caspase-2 both in normal and apoptotic cells. We next investigated the appearance of the 33-kDa form of caspase-2 in different cell lines when apoptosis was induced by UV irradiation. As shown in Fig. 1 c, the processed form of caspase-2 was detected upon apoptosis induction in most of the cell lines tested, with the exception of the MCF-7 cells. The anti-caspase-2 antibody also showed cross-species reactivity, since it detected a band of 50 kDa in the cellular lysates of murine fibroblasts NIH 3T3 that is converted in a p33 form upon induction of apoptosis (data not shown). To study the hierarchy of caspase-2 processing in respect to different caspases during apoptosis in vivo, we decided to use cell lines containing defined mutations in specific caspases. We took advantage of the IMR90 cells transformed with E1A oncogene and containing a dominant negative form of caspase-9 (caspase-9 DN) (20Fearnhead H.O. Rodriguez J. Givek E-E. Guo W. Kobayashi R. Hannon G. Lazebnik Y. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 13664-13669Crossref PubMed Scopus (159) Google Scholar). As a control IMR90 fibroblasts expressing E1A alone were used. It has been previously demonstrated that IMR90-E1A cells expressing the caspase-9 DN do not show all of the "classical" apoptotic features when challenged by etoposide treatment, whereas in these cells cytochrome c release from mitochondria was reported as normal (20Fearnhead H.O. Rodriguez J. Givek E-E. Guo W. Kobayashi R. Hannon G. Lazebnik Y. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 13664-13669Crossref PubMed Scopus (159) Google Scholar). We also used the human MCF-7 breast carcinoma cell line, which is devoid of caspase-3 due to the functional deletion of theCASP-3 gene (21Janicke R.U. Sprengart M.L. Wati M.R. Porter A.G. J. Biol. Chem. 1998; 273: 9357-9360Abstract Full Text Full Text PDF PubMed Scopus (1720) Google Scholar). This cell line was used to reintroduce wild type caspase-3 or its catalytic inactive point-mutated derivative caspase-3 (CI) as a control (22Faleiro L. Lazebnik Y. J. Cell Biol. 2000; 151: 951-960Crossref PubMed Scopus (207) Google Scholar). We used FACS analysis to confirm that the different cell lines selected showed different susceptibility to enter cell death by apoptosis following UV irradiation. UV-irradiated IMR90-E1A cells underwent apoptosis as evidenced by cell morphology, chromatin condensation, and detachment from the adhesion substrate. Levels of apoptosis were assessed by FACS analysis using propidium iodide staining (Fig.2 a). Analysis of DNA content (Fig. 2) revealed a significant increase of cells with sub-G1 DNA content, indicative of apoptosis (19Brancolini C. Sgorbissa A. Schneider C. Cell Death Differ. 1998; 5: 1042-1050Crossref PubMed Scopus (60) Google Scholar), when IMR90-E1A cells were UV-irradiated. On the contrary, IMR90-E1A cells containing caspase-9 DN were impaired in entering efficiently apoptosis following UV irradiation, as evidenced by FACS analysis. Indeed, in these cell lines, floating cell fragments could be detected upon UV irradiation (data not shown). Reintroduction of wild type caspase-3 in MCF-7 cells renders these cells susceptible to apoptosis following UV irradiation, whereas cells expressing a catalytic inactive form of caspase-3 were partially resistant to UV-dependent cell death (Fig. 2 b). Moreover, when MCF-7 caspase-3 CI cells were UV-irradiated, some cell debris was detected floating in the medium (data not shown). Having selected two human cell lines containing defined mutations in caspase-3 and caspase-9, we analyzed the relationships between caspase-2 processing and the above mentioned caspases by inducing cell death with two different stimuli: UV irradiation and TNF-α. As mentioned before, some dead cells floating in the medium were observed also in the case of UV-irradiated IMR90-E1A-caspase-9 DN that were resistant to apoptosis as judged by FACS analysis. These dead cells might be indicative of a necrotic or "frustrated apoptotic" response that occurs following extensive DNA damage and/or mitochondrial dysfunction. In our studies, we have focused our attention on the status of caspase-2 in the population of dead cells that can be isolated as nonadherent cells (Fig. 3, D) from the population of still viable cells (Fig. 3, V). It is important to note that the population of nonadherent cells was consistently reduced in cells expressing the catalytic inactive caspases (data not shown), but in our Western analysis the total amount of protein lysates loaded for each sample was normalized. In apoptotic IMR90-E1A cells, almost all caspase-2 was processed to the p33 form, as shown in Fig. 3 a, while in floating cells from IMR90-E1A-caspase-9 DN (D), caspase-2 processing was largely impaired. The same lysates were also analyzed for PARP processing by using an antibody specific for the cleaved form. Processing of PARP was observed in both cell lines although the presence of a dominant negative form of caspase-9 clearly reduced the amount of cleaved PARP detectable. Actin was used as loading control. We next analyzed the dependence of caspase-2 processing on caspase-9 in response to the activation of a different apoptotic pathway. IMR90-E1A and IMR90-E1A-caspase-9 DN were treated with TNF-α in the presence of cycloheximide, and Western blot analysis was performed using lysates prepared from TNF-α/CHX- and CHX-treated or untreated cells. Nonadherent dead cells (Fig. 3, D) and adherent viable cells (V) were harvested separately. In TNF-α-triggered cell death, despite of the presence of a dominant negative form of caspase-9, caspase-2 was normally processed, thus generating the p33 fragment (Fig. 3 b). PARP processing during apoptosis induced by TNF-α was independent from caspase-9 activity. Here again, detection of actin was used as a loading control. The previous experiment clearly shows that caspase-2 processing following UV irradiation is dependent on caspase-9. Since caspase-9 activates caspase-3 in response to DNA damage (20Fearnhead H.O. Rodriguez J. Givek E-E. Guo W. Kobayashi R. Hannon G. Lazebnik Y. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 13664-13669Crossref PubMed Scopus (159) Google Scholar), we next analyzed if caspase-3 is critical for caspase-2 processing in vivo. Human MCF-7 cells were UV-irradiated, and 18 h later nonadherent dead cells (Fig. 3, D) and adherent viable (V) cells were harvested and combined, and lysates were prepared for Western analysis. As shown in Fig. 4 a, in UV-irradiated MCF-7 wild type caspase-3, a consistent fraction of caspase-2 could be detected as the p33-processed form, while in MCF-7 caspase-3 CI cells, caspase-2 processing was undetectable. Processing of PARP was reduced but still detectable in MCF-7 caspase-3 CI as described above in IMR90-E1A caspase-9 DN. We next analyzed the dependence of caspase-2 processing on caspase-3 under TNF-α-induced apoptosis. MCF-7 wild type caspase-3 and caspase-3 CI cells were treated with TNF-α in the presence of cycloheximide for 36 h, and Western blot analysis was performed. Nonadherent dead (Fig. 4, D) and adherent viable (V) cells were harvested separately. Caspase-2 was fully processed when apoptosis was induced in wild type caspase-3 MCF-7 cells treated with TNF-α and CHX. In apoptotic MCF-7 caspase-3 CI cells, caspase-2 processing was largely impaired. PARP was similarly processed in both cell lines, thus confirming the induction of cell death. Having demonstrated that caspase-3 is the critical enzyme for an efficient processing of caspase-2 in vivo, we wanted to analyze the ability of caspase-3 to cleave caspase-2 in vitro with respect to other long and short prodomain caspases involved in the apoptotic response. In vitro proteolytic assays using recombinant caspases-3 were performed. Full-length caspase-1, caspase-2, caspase-3, caspase-6, caspase-7 caspase-8, and caspase-9 cDNA were in vitrotranslated and then incubated with purified caspase-3 (Fig.5). Treatment with an increasing amount of purified caspase-3 for 30 min at 37 °C leads to the processing of all of the tested caspases with the exception of caspase-1. Among the different long prodomain caspases, some processing of caspase-2 to generate the p18/p12 active form was detectable after incubation with only 0.01 ng of caspase-3, while the full processing of the p48 form was detectable when 500 ng of caspase-3 were used. In contrast, caspase-9 and caspase-8 were weak substrates of caspase-3 and only partially processed after incubation with 10 ng of the recombinant caspase-3. Among the different short prodomain caspases analyzed, caspase-7 was the most efficiently cleaved by caspase-3, and partial processing was detectable after incubation with 10 ng of recombinant caspase-3. PARP, a well defined substrate of caspase-3 (4Earnshaw W.C. Martins L.M. Kaufmann S.H. Annu. Rev. Biochem. 1999; 68: 383-424Crossref PubMed Scopus (2451) Google Scholar, 23Nicholson D.W. Ambereen A. Thornberry N.A. Vaillancourt J.P. Ding C.K. Gallant M. Gareau Y. Griffin P.R. Labelle M. Lazebnik Y.A. Munday N.A. Raju S.M. Smulson M.E. Yamin T.T., Yu, V.L. Miller D.K. Nature. 1995; 376: 37-38Crossref PubMed Scopus (3796) Google Scholar), was used as a control under the same conditions. From this analysis, we can suggest that caspase-2, among the different caspases analyzed, is the best substrate for caspase-3. Residual caspase-2 processing was observed when apoptosis was induced by TNF-α in MCF-7 cells defective for caspase-3 activity. Since caspase-3 and caspase-7 show overlapping cleavage consensus sequences and share many death substrates in vivo(4Earnshaw W.C. Martins L.M. Kaufmann S.H. Annu. Rev. Biochem. 1999; 68: 383-424Crossref PubMed Scopus (2451) Google Scholar), we asked whether caspase-7 was able to cleave caspase-2 in anin vitro assay. In vitro translated caspase-2 was incubated with increasing amounts of purified caspase-7 for 60 min at 37 °C. As shown in Fig.6 a, 1 μg of caspase-7 was required for the full processing of caspase-2, while partial caspase-2 processing was observed after incubation with 10 ng of caspase-7. Under the same experimental conditions, full processing of PARP was observed after incubation with 10 ng of purified caspase-7. This analysis suggests that caspase-7 can cleave caspase-2 in vitro,although with a lower efficiency when compared with PARP. It is possible that caspase-7 is responsible for the limited proteolytic processing of caspase-2 during apoptosis in MCF-7 caspase-3 CI cells (25Germain M. Affar E.B. D'Amours D. Dixit V.M. Salvesen G.S. Poirier G.G. J. Biol. Chem. 1999; 274: 28379-28384Abstract Full Text Full Text PDF PubMed Scopus (394) Google Scholar). We therefore analyzed if caspase-7 was activated when these cells were treated with TNF-α/CHX. We have analyzed by Western blot caspase-7 expression, as shown in Fig.6 b, with an antibody that recognizes the caspase-7 proenzyme migrating at around 34 kDa. Disappearance of the 34-kDa band was considered as evidence for caspase-7 processing. Nonadherent apoptotic and adherent nonapoptotic cells were harvested separately. Caspase-7 was processed in MCF-7 wild type caspase-3 cells during apoptosis, and its processing was reduced but still detectable in apoptotic MCF-7 containing catalytic inactive caspase-3. The dependence of caspase-2 processing on caspase-3 argues that caspase-2 acts as an effector caspase and that its activation is a downstream event in the proteolytic cascade triggering cell death. On the other hand, caspase-2 possesses a prodomain, which is a marker for caspases acting at the apex of the proteolytic cascade, and its overexpression can trigger cell death by apoptosis, possibly as a consequence of caspase-2 activation following prodomain-mediated oligomerization (14Colussi P.A. Harvey N.L. Kumar S. J. Biol. Chem. 1998; 273: 24535-24542Abstract Full Text Full Text PDF PubMed Scopus (138) Google Scholar, 15Colussi P.A. Harvey N.L. Shearwin-Whyatt L.M. Kumar S. J. Biol. Chem. 1998; 273: 26566-26570Abstract Full Text Full Text PDF PubMed Scopus (72) Google Scholar, 26Butt A.J. Harvey N.L. Parasivam G. Kumar S. J. Biol. Chem. 1998; 273: 6763-6768Abstract Full Text Full Text PDF PubMed Scopus (98) Google Scholar, 27Wang L. Miura M. Bergeron L. Zhu H. Yuan J. Cell. 1994; 78: 739-750Abstract Full Text PDF PubMed Scopus (801) Google Scholar). If caspase-2 induces apoptosis simply by acting as an effector caspase, its ability to induce cell death should be independent from mutations in the regulative caspases such as caspase-9. Therefore, we have investigated the ability of caspase-2 to efficiently induce cell death when overexpressed in cells impaired in caspase-9 activity. IMR90-E1A caspase-9 DN and IMR90-E1A were co-transfected with caspase-2 and GFP used as a reporter, and the appearance of apoptotic cells was scored 24 h after transfection. As shown in Fig. 7 a, the apoptotic response triggered by caspase-2 overexpression in IMR90-E1A caspase-9 DN was impaired
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客观：为了处于网膜的损坏调查 Caspase-3 的角色，在老鼠由轻暴露引起了。方法：到视网膜的光射损伤被坚持的暴露导致到照明(紧张：30 000 +/- 50 勒克司) 操作，为 30 的显微镜在 Sprague-Dawley 老鼠的右眼纪录。视网膜的病理学的变化被观察在下面光并且在不同时间点的电子显微镜学，是 6 个小时，在轻暴露以后的 1， 3， 7，和 15 天。视网膜细胞的 Apoptosis 被流动血细胞计数分析。Caspase-3 的活动被使用 Caspase-3 试金工具包评估。同时， Caspase-3 朊酶的表示与西方的污点分析被决定。结果：考试结果光并且传播电子显微镜显示出视网膜的内部、外部的片断的那浮肿，特别在内部片断内的线粒体，变得明显在轻暴露以后的 6 个小时。变化与增加的时间一起被败坏。discoidalvalve 的 Thestructures 同时在外部片断分裂了。混乱安排原子核， karyopycnosis，并且在外胞核层变瘦被观察。网膜的颜料上皮几乎消失在以后的舞台期间。与 PI 相结合的 Annexin-V 的染色的结果证明 apoptotic 房间的比例与时间增加了。然而，在第 7 白天(82.7%) 和第 15 白天(80.4%) 之间的比例没显示出有效差量。在它开始下降以前， Caspase-3 与时间的消逝变得显著地活跃，它在第 7 白天在第 6 小时从 0.02 增加了到 9.8 的山峰。西方的污点在 7thday 和第 15 白天检测了 Caspase-3 的活跃形式的表情。结论：光敏电阻器房间的 Apoptosis 显著地涉及轻损坏， Caspase-3 朊酶可以在老鼠在轻暴露以后在视网膜的 apoptotic 过程起一个重要作用。
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Despite two decades of research, the role of caspase-2 in physiology and disease is still poorly understood and controversial. This Cell Science at a Glance article provides an overview of the proposed functions and possible modes of action and regulation of caspase-2. In addition, we will highlight recent findings that may lead to a more comprehensive understanding of this highly conserved protease.Caspase-2, formerly known as ICH-1 (Wang et al., 1994) or NEDD-2 (Kumar et al., 1994), is a highly conserved but functionally poorly defined member of the caspase family, a group of cystein-driven aspartate-directed endopeptidases that are involved in cell death, inflammation and differentiation (Li and Yuan, 2008). Caspase-2 contains an N-terminal caspase recruitment domain (CARD), followed by a large subunit containing the active site (p19) and a small subunit (p12). Thus, caspase-2 is most similar to caspase-9, the initiator of the intrinsic pathway of apoptosis (Li and Yuan, 2008). However, in contrast to 'conventional' initiator caspases, such as caspase-9 or the apical caspase of extrinsic apoptosis, caspase-8, caspase-2 does not process apoptosis effectors that need to be cleaved by initiators for their activation, such as caspase-3, -6 or -7 (Guo et al., 2002; Van de Craen et al., 1999), suggesting that caspase-2 has other substrates (see supplementary material Table S1 and poster). Functionally, caspase-2 has been implicated in the regulation of cell death that is induced by metabolic imbalance, DNA damage, endoplasmic reticulum (ER) stress, mitotic catastrophe and others (reviewed by Krumschnabel et al., 2009; Vakifahmetoglu-Norberg and Zhivotovsky, 2010). The precise function of caspase-2, that is whether it functions as an initiator or effector caspase, is still unknown, and studies on caspase-2 activation are confounded by the fact that caspase-2 is a substrate for caspase-8 as well as for caspase-3 (Paroni et al., 2001; Van de Craen et al., 1999).Similar to other initiator caspases, caspase-2 is activated by proximity-induced oligomerization and trans-cleavage in vitro, and ectopic overexpression is sufficient for its activation in cells (Butt et al., 1998; Read et al., 2002). Endogenous caspase-2 becomes activated as part of different signaling pathways (see poster), some of which depend on the DNA damage-induced formation of the PIDDosome, a complex composed of the CARD- and death domain (DD)-containing adapter protein RAIDD (also known as CRADD) and the p53-inducible DD-containing protein PIDD (also known as LRDD) (Tinel and Tschopp, 2004). However, the significance of this platform in vivo has been challenged by genetic experiments that have demonstrated that PIDD and RAIDD are dispensable for caspase-2 activation (Kim et al., 2009; Manzl et al., 2009), suggesting that alternative modes of caspase-2 activation exist.One such alternative mechanism for caspase-2 activation is possibly mediated by caspase-8 in the death-inducing signaling complex (DISC), which is formed upon CD95 (also known as FAS) receptor clustering (Lavrik et al., 2006; Olsson et al., 2009). The role of caspase-8-mediated caspase-2 activation is, however, controversial, because this event does not necessarily contribute to apoptosis downstream of CD95 (Lavrik et al., 2006). Hence, DISC-dependent caspase-2 activation might be a 'bystander' effect due to its incidental co-recruitment together with that of 'non-canonical' DISC components, such as RAIDD (Duan and Dixit, 1997) or receptor-interacting protein 1 (RIP1) (Ahmad et al., 1997).Other alternative caspase-2 activation mechanisms have been described, including caspase-2 dimerization and subsequent auto-proteolysis, which is induced by K+ efflux in response to bacterial pore-forming toxins (Imre et al., 2012) or heat-shock-triggered protein aggregation (Tu et al., 2006). Furthermore, alternative mRNA splicing (Wang et al., 1994), post-translational modifications, including N-terminal acetylation (Yi et al., 2011) and phosphorylation, appear to impact on and fine-tune these activating events (see poster). The variety of activation mechanisms suggest that caspase-2 becomes activated under diverse conditions of cellular stress, as discussed below.Caspase-2 has been implicated in the induction of cell death by pathogenic bacteria, such as Brucella, Staphylococcus aureus and Salmonella (Chen et al., 2011; Chen and He, 2009). Caspase-2-deficient murine macrophages are protected from Salmonella-induced cell death and show decreased processing of caspase-1, a major driver of inflammation upon infection (Jesenberger et al., 2000). Similarly, cells lacking caspase-2 are also protected from cell death induced by Staphylococcus aureus α-toxin, but the underlying mechanisms that are responsible for escape from cell death have not yet been defined (Imre et al., 2012).In addition to bacterial infections, caspase-2 activation has also been described in the apoptotic response to viral infections by the single-stranded (ss) RNA Rhabdoviridae, such as Maraba virus or vesicular stomatitis virus (VSV). Both viruses are known to induce a strong ER or unfolded protein stress response (UPR), which can lead to apoptosis (Mahoney et al., 2011). Interestingly, by keeping protein levels of RAIDD low, the ER-stress response machinery appears to be able to delay caspase-2-dependent apoptosis upon infection (Mahoney et al., 2011). Under conditions of sustained ER stress, however, the ER stress response factor IRE1α (also known as ERN1), an ER transmembrane kinase-endoribonuclease (RNase), promotes the rapid degradation of microRNAs that target caspase-2 mRNA. This, in turn, causes a rapid induction of caspase-2 protein expression that might contribute to the induction of apoptosis (Upton et al., 2012). Mechanistically, apoptosis induction by caspase-2 has been linked to its cleavage of the pro-apoptotic BCL2 family protein BID into its active form tBID (Guo et al., 2002), and thymocytes or mouse embryonic fibroblasts (MEFs) from Bid−/− mice have been reported to resist ER-stress-dependent apoptosis (Upton et al., 2008). However, as the microRNAs that are inactivated by IRE1α also repress multiple pro-apoptotic BCL2 family proteins that have been previously implicated in ER-stress induced apoptosis, such as BIM (miR-17) or PUMA (miR-125b) (Le et al., 2011; Xiao et al., 2008), it will be important to assess the functional significance of caspase-2 induction in response to ER stress.Most reports implicate caspase-2 in controlling cell death in response to DNA damage (reviewed by Krumschnabel et al., 2009). But although it has been shown that caspase-2 and the PIDDosome, as well as BID, become activated in response to DNA damage, genetic analysis in mice failed to reveal an essential function for any of these genes in normal physiology or in response to DNA damage (Kaufmann et al., 2007; Manzl et al., 2009; our unpublished data). Although the possibility of compensatory events taking place in knockout mice cannot be entirely excluded, these data suggest that these genes act redundantly to contribute to the robustness of the DNA damage response.Such a redundant mechanism has been identified in p53-mutated zebrafish, in which caspase-2 controls cell death in response to DNA damage downstream of the DNA damage response pathway. As in other organisms, inactivation of p53 function causes resistance to radiation in Danio rerio, which can be overcome by inhibition of the checkpoint kinase 1 (Chk1) (Sidi et al., 2008). The functional abrogation of Chk1 results in the failure of the G2-M checkpoint and premature entry into mitosis, leading to aberrant mitosis and apoptosis after DNA damage (see below). Under these conditions, cell death has been found to be independent of several well-known apoptosis regulators, including caspase-3, -8 and -9, and could not be inhibited by overexpression of BCL2, but requires caspase-2 and the PIDDosome components PIDD and RAIDD (Ando et al., 2012; Sidi et al., 2008). Activation of caspase-2 under these conditions (i.e. irradiation upon inhibition of Chk1 in the absence of functional p53) appears to be controlled by ataxia telangiectasia mutated (ATM) kinase, both in zebrafish and in human cancer cell lines (Ando et al., 2012; Sidi et al., 2008). ATM can directly phosphorylate PIDD, triggering the assembly of the PIDDosome and activation of caspase-2 (Ando et al., 2012). Although epistasis clearly posits that Chk1 acts as an inhibitor of the ATM-dependent activation of the PIDDosome, the mechanistic nature of this inhibition remains to be understood. Elucidating this mechanism is important to improve our understanding of the cellular responses upon DNA damage in the presence or absence of functional p53 and might help to predict whether the application of pharmacological Chk1 inhibitors will radiosensitize tumor cells with a given genetic makeup (reviewed by Garrett and Collins, 2011).In addition to the proapoptotic protein BID, only a few other caspase-2 substrates with a link to cell death regulation have been identified so far (see supplementary material Table S1 and poster), including the p53-related transcription factor p63 (Jeon et al., 2012). In response to DNA damage, ΔNp63, a splice variant that inhibits the proapoptotic transcriptional activity of full length p63 (the TAp63 isoform) by dimerization in a dominant-negative manner, is modified by the ubiquitin-like protein insulin-stimulated gene 15 (ISG15), which is required for its subsequent cleavage and inactivation by caspase-2. This cleavage leads to the nuclear export of the inhibitory fragment, allowing TAp63 to induce transcription of relevant target genes like proapoptotic PUMA and NOXA (also known as BBC3 and PMAIP1, respectively) (Jeon et al., 2012). In a xenograft tumor model, no difference in growth was observed between cells expressing either wild-type ΔNp63, a version of TAp63 resistant to caspase-2 cleavage, or a version of TAp63 that is defective for ISG15 conjugation. Upon DNA damage, however, wild-type cells are more sensitive (Jeon et al., 2012), suggesting that TAp63 has an important role in the proapoptotic response to DNA damage, and that this requires caspase-2 (see poster).Caspase-2 is the sole caspase that is able to translocate from the cytosol into the nucleus upon activation, corroborating the hypothesis that it might act as a nuclear protease to orchestrate the response to DNA damage (Baliga et al., 2003; Colussi et al., 1998). The role of nuclear localization is still debated (reviewed by Krumschnabel et al., 2009) and sophisticated imaging techniques have revealed that the bulk of caspase-2 activation in response to various stressors, including DNA damage (Bouchier-Hayes et al., 2009), is in the cytoplasm. Whereas the augmentation of p53 signaling that is mediated by caspase-2 appears to rely on the cleavage of the p53 inhibitor MDM2 in the cytoplasm (Oliver et al., 2011), as discussed below, processing of ΔNp63 conceivably occurs in the nucleus, suggesting that caspase-2 acts in the cytoplasm to control p53 and in the nucleus in the p63 pathway (see poster).Cell death associated with an abnormal mitosis is often referred to as mitotic catastrophe. Confusingly, a consensus for the definition of 'mitotic catastrophe' is missing and the term carries various meanings (reviewed by Vitale et al., 2011). Caspase-2 has been linked to cell death that is induced by aberrant mitosis in cancer cells that transiently arrest in prometaphase after cell fusion (Castedo et al., 2004) and to cell death by abnormal mitosis that has been induced by microtubule-interfering agents, such as vincristine and Taxol (Ho et al., 2008), but the underlying mechanisms remain to be fully defined. Several lines of evidence support the notion that caspase-2 can trigger cell death that is induced by abnormal mitosis, but how this 'abnormality' is sensed and translated into caspase-2 activation is still unknown. One possible mechanism is the inhibition of caspase-2 auto-cleavage (by phosphorylation of the conserved residue Ser340 located within the linker region connecting its large and small subunit) by cyclin-dependent kinase 1 (CDK1) (Andersen et al., 2009). Mitotic arrest is likely to affect the propensity of caspase-2 to become activated: the degree of its phosphorylation might be altered by the balance of CDK1 and protein phosphatase 1 (PP1), which can dephosphorylate Ser340 (Andersen et al., 2009) to counteract the inhibitory phosphorylation by CDK1. Recently, caspase-2-deficient MEFs have been shown to accumulate micronuclei and hyperploidy faster than wild-type cells, indicating that they are genetically unstable (Dorstyn et al., 2012). Whether caspase-2 deficiency causes micronucleation because of its postulated role in the DNA damage checkpoint or because of its role in response to mitotic errors remains to be established.Despite the evidence that caspase-2 controls cell death in response to DNA damage, caspase-2-deficient mice develop without gross phenotypes (O'Reilly et al., 2002). Mice lacking caspase-2 show signs of premature aging (Zhang et al., 2007) and a higher number of oocytes (Bergeron et al., 1998). The premature aging phenotype appears to be caused by increased oxidative stress in aged caspase-2-deficient mice (Shalini et al., 2012), which was found to correlate with decreased expression levels of the cysteine sulfinyl-reductases sestrin-2 and -3, reduced activity of the FOXO transcription factors FOXO1 and FOXO3a and/or increased activation of p53 and p21 (encoded by Cdkn1a) (Shalini et al., 2012). In contrast to aged cells, MEFs that have been derived from young Casp2−/− mice show an impaired activation of p53 and p21, leading to evasion of senescence (Dorstyn et al., 2012). Therefore, the increased p53 activation noted in aged animals might be a consequence of changes that are induced by lack of caspase-2 in young mice that lead to accumulation of damage over time and subsequently increased p53 activity, which no longer is co-regulated by caspase-2 function. This phenomenon might be driven by the lack of an augmentative function of this protease on p53 stability (see below).Caspase-2 is crucial for oocyte apoptosis in the mouse and in Xenopus laevis (Bergeron et al., 1998; Nutt et al., 2005). Work based on Xenopus oocytes and egg extracts has shown that pentose-phosphate pathway (PPP)-driven generation of NADPH appears to be critical for oocyte survival (Nutt et al., 2005). Engagement of the PPP by addition of glucose 6-phosphate could prevent oocyte death by inhibiting caspase-2 due to its phosphorylation at Ser135 (Ser164 in human). This event is catalyzed by calcium/calmodulin-dependent kinase II (CaMKII; encoded by CAMK2G) and leads to the subsequent binding of caspase-2 to 14-3-3ζ, which prevents caspase-2 activation (Nutt et al., 2005). Nutrient depletion promotes the release of caspase-2 from 14-3-3ζ, which, in turn, allows the dephosphorylation of Ser135 by PP1, relieving caspase-2 inhibition and causing cell death (Nutt et al., 2009). The ability of 14-3-3ζ to interact with and inhibit caspase-2 in oocytes and human cancer cell lines (Andersen et al., 2011) is regulated by acetylation. The control of the acetylation status of 14-3-3ζ is exerted partly through the activity of the deacetylase sirtuin 1 (SIRT1) that can respond to glucose 6-phosphate by an unknown mechanism (Andersen et al., 2011). Intriguingly, whereas the inhibition of SIRT1 in cancer cells is sufficient to drive caspase-2 release from 14-3-3ζ, it has been reported to not be sufficient to trigger cell death in the absence of another proapoptotic stimulus, for instance the microtubule poison Taxol (Andersen et al., 2011). How the metabolic control of caspase-2 intersects with other pathways that lead to caspase-2 activation is an important but still unresolved question.On the basis of its reported proapoptotic properties, caspase-2 is considered to act as a tumor suppressor (Kumar et al., 1995). Consistently, reduced expression of caspase-2, which is encoded on human chromosome 7q in a region that is frequently lost in human tumors, has been reported in acute myeloid leukemia (AML) and acute juvenile lymphoblastic leukemia (ALL) patients that are refractive to therapy and have a poor prognosis (Holleman et al., 2005; Johansson et al., 1993; Mrózek, 2008). Reduced caspase-2 protein levels have also been reported in a meta-analysis of published data sets for Burkitt lymphoma (BL), mantle cell lymphoma (MCL), chronic lymphocytic leukemia (CLL) and hairy cell leukemia, as well as in solid tumors, such as hepatocellular carcinoma, gastric and ovarian cancer, as well as metastasizing tumors of the brain (Kumar, 2009; Ren et al., 2012).Consistent with a tumor suppressive function of caspase-2, MEFs derived from Casp2−/− mice that are transformed with the oncoproteins E1A and Ras show increased proliferation in vitro and are more tumorigenic when they are transplanted into nude mice (Ho et al., 2009). These cells also show an impaired arrest in response to irradiation, as well as decreased cell death (Ho et al., 2009). Failure to arrest the cell division cycle arrest in response to DNA damage (Ho et al., 2009) might lead to increased genomic instability and chromosomal aberrations, which can promote tumorigenesis. Consistent with this idea, it has been found that lymphomagenesis caused by overexpression of Myc, which also promotes DNA damage, is accelerated in Casp2−/− mice (Ho et al., 2009; Manzl et al., 2012). Surprisingly, however, in other models of DNA-damage-induced tumorigenesis, for example in 3-methyl-cholantrene-driven fibrosarcomas or fractionated irradiation-driven thymic lymphomas, loss of caspase-2 does not accelerate tumorigenesis, which argues against a general role of caspase-2 in tumor suppression (Manzl et al., 2012). It remains to be determined whether caspase-2 can suppress other forms of oncogene-driven tumor formation, in addition to those mediated by Myc.The tumor suppressive capacity of caspase-2 has also been linked to its role in regulating cell death upon oncogenic stress because the strong selective pressure to inactivate the p53 pathway in Myc-induced B cell lymphoma (Ho et al., 2009) is reduced in the absence of caspase-2 (Manzl et al., 2012). This can be explained by an intrinsic defect in p53 activation or stabilization in developing B cells in such mice (Dorstyn et al., 2012; Ho et al., 2009), and could be due to the loss of caspase-2-dependent cleavage of the E3 ligase MDM2, which converts it from an inhibitor into an activator of p53 (Oliver et al., 2011). Additionally, caspase-2 might also have a direct, but less-well understood, effect on the translation of Cdkn1a mRNA (which encodes p21) (Sohn et al., 2011).In summary, the involvement of caspase-2 in cell death initiation appears to be restricted to a rather limited number of cellular conditions, such as p53 deficiency, aberrant mitosis or nutrient deprivation. Despite the high degree of conservation, caspase-2 is much less understood in terms of its regulation and function when compared with other caspases. It remains to be fully explored whether caspase-2 indeed acts simultaneously as an initiator and effector of apoptosis, an idea that is compatible with its proposed ancestral function. The basic cell death machinery that operated with only one caspase might have been replaced by more sophisticated mediators of cell death over the course of evolution, rendering caspase-2 largely redundant today. Of note, in some paradigms, we now believe caspase-2 does not exert a direct pro-death role, neither as initiator nor as an effector, but instead acts as 'damage sensor' mediating limited proteolysis for signaling, and that this reflects a more general phenomenon in caspase-2 biology. Caspase-2 might, therefore, be embedded in intricate cellular signaling pathways that allow cells to initiate apoptosis as the last resort in contrast to catastrophic signals, which require cellular suicide to be rapidly committed for the sake of the entire organism. Caspase-2 research will therefore remain challenging, but it is likely to unveil important cellular response pathways to different harmful processes, such as DNA damage, oxidative or metabolic stress and possibly others, that are yet to be linked to caspase-2 activity.
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Members of the caspase family of cysteine proteases coordinate the highly disparate processes of apoptosis and inflammation. However, although hundreds of substrates for the apoptosis effector caspases (caspase-3 and caspase-7) have been identified, only two confirmed substrates for the key inflammatory protease (caspase-1) are known. Whether this reflects intrinsic differences in the substrate specificity of inflammatory versus apoptotic caspases or their relative abundance in vivo is unknown. To address this issue, we have compared the specificity of caspases-1, -3, and -7 toward peptide and protein substrates. Contrary to expectation, caspase-1 displayed concentration-dependent promiscuity toward a variety of substrates, suggesting that caspase-1 specificity is maintained by restricting its abundance. Although endogenous concentrations of caspase-1 were found to be similar to caspase-3, processed caspase-1 was found to be much more labile, with a half-life of ∼9 min. This contrasted sharply with the active forms of caspase-3 and caspase-7, which exhibited half-lives of 8 and 11 h, respectively. We propose that the high degree of substrate specificity displayed by caspase-1 is maintained through rapid spontaneous inactivation of this protease. Members of the caspase family of cysteine proteases coordinate the highly disparate processes of apoptosis and inflammation. However, although hundreds of substrates for the apoptosis effector caspases (caspase-3 and caspase-7) have been identified, only two confirmed substrates for the key inflammatory protease (caspase-1) are known. Whether this reflects intrinsic differences in the substrate specificity of inflammatory versus apoptotic caspases or their relative abundance in vivo is unknown. To address this issue, we have compared the specificity of caspases-1, -3, and -7 toward peptide and protein substrates. Contrary to expectation, caspase-1 displayed concentration-dependent promiscuity toward a variety of substrates, suggesting that caspase-1 specificity is maintained by restricting its abundance. Although endogenous concentrations of caspase-1 were found to be similar to caspase-3, processed caspase-1 was found to be much more labile, with a half-life of ∼9 min. This contrasted sharply with the active forms of caspase-3 and caspase-7, which exhibited half-lives of 8 and 11 h, respectively. We propose that the high degree of substrate specificity displayed by caspase-1 is maintained through rapid spontaneous inactivation of this protease.
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Satratoxins, which are produced Stachybotrys species, have been recognized as potential agents associated with sick building syndromes. In spite of the potential importance of satratoxins, the molecular mechanism induced by them remains unknown. Recently, we have found that satratoxin G induced potent apoptosis in HL-60 human promyelotic leukemia cells and both caspase-8 and caspase-9 are activator of caspase-3. However, the detail activation profile of caspases and role of the other caspases including caspase-6, caspase-7 and a novel initiator caspase, caspase-2 remain to be clear. Here we report detail caspase cascade in satratoxin G-induced apoptosis in HL-60 cells. Caspase assay with DEVD-AMC, a fluorogenic substrate, and western blot analysis of caspase-7 suggested that caspase-3 was involved, but casapase-7 was not directly involved in satratoxin-induced apoptosis. We investigated in-depth time courses of caspase activation. Caspase-3 was activated after 1 hour from satratoxin treatment. Activation of caspase-2 was observed apparently as early as 1 h. Caspase-9 was also activated significantly at the 1 hour time point. Caspase-6 and caspase-8 were activated at the 3-hour time point. Western blot analysis showed that cleavage of bid and activation of caspase-12 were not involved in satratoxin-mediated apoptosis. These findings suggest that the initial activators of caspase-3 are caspase-2 and casapse-9 in satatoxin-induced apoptosis. Because caspase-8 and casapse-9 were activated independently, caspase-8 is also involved in activation of caspase-3 in the late stage of the apoptosis. Moreover, caspase-6 appears to be associate with activation of caspase-8. Both caspase-7 and caspase-12 may be not potential caspases in the apoptosis.
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Caspase belongs to proteolytic enzymes family closely related to cell apoptosis,which exists in most metazoan cells as its preemzyme and plays an important role in apoptosis. Under the influence of apoptosis signals,caspase first activates initiator caspase, resulting in cascade reaction of caspase,and then splits specific substrates by executioner caspase activated to induce cell apoptosis. So the activation of caspase was the central component for inducing apoptosis, and there are different activating mechanisms between initiator caspase and executioner caspase.
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