|Home | About | Journals | Submit | Contact Us | Français|
Caspase-2 is an initiator caspase, activated in response to heat shock and other stressors that induce apoptosis. Activation of caspase-2 requires induced proximity resulting after recruitment to caspase-2 activation complexes, such as the PIDDosome. We have adapted bimolecular fluorescence complementation (BiFC) to measure caspase-2 induced proximity in real time, in single cells. Non-fluorescent fragments of the fluorescent protein Venus that can associate to reform the fluorescent complex were fused to caspase-2 allowing visualization and kinetic measurements of caspase-2 induced proximity after heat shock and other stresses. This revealed that the caspase-2 activation platform occurred in the cytosol and not in the nucleus in response to heat shock, DNA damage, cytoskeletal disruption and other treatments. Activation, as measured by this approach, in response to heat shock, was RAIDD-dependent and upstream of mitochondrial outer membrane permeabilization. Furthermore we identify Hsp90α as a key negative regulator of heat shock-induced caspase-2 activation.
The initiator caspases-8, -9 and -10 are responsible for cleavage and activation of the executioner caspases-3, and -7 that orchestrate apoptosis. Unlike executioner caspases, initiator caspases are activated by dimerization (Boatright et al., 2003; Donepudi et al., 2003). Upon dimerization, initiator caspases undergo auto-cleavage that stabilizes the active enzyme (Pop et al., 2007). Initiator caspases are generally activated by recruitment to large molecular weight protein complexes or “activation platforms” that induce the proximity of caspase molecules, facilitating dimerization (Boatright and Salvesen, 2003). For example, caspase-8 is activated when bound to the death inducing signaling complex (DISC) following death receptor ligation. In a similar fashion, caspase-9 is activated by the Apaf-1 apoptosome formed upon cytochrome c release from the mitochondria (Creagh et al., 2003).
Caspase-2 is also an initiator caspase but specific details of how, where and when it is activated remain unclear. Unlike other initiator caspases, caspase-2 does not directly activate executioner caspases but instead acts via mitochondrial outer membrane permeabilization (MOMP) to induce apoptosis (Guo et al., 2002). The caspase-2 activation platform appears to be a complex of proteins termed the PIDDosome, consisting of PIDD, the adaptor protein RAIDD and caspase-2 (Tinel and Tschopp, 2004). RAIDD binds and induces dimerization of the pro-form of caspase-2 via interaction between the caspase recruitment domain (CARD) present in both proteins (Duan and Dixit, 1997). A second caspase-2 activation platform has been identified to contain PIDD, caspase-2 and the nuclear serine/threonine kinase DNA-PKcs and does not require RAIDD (Shi et al., 2009). Caspase-2 was initially considered to be a component of the TNF receptor complex and hence has been reported to be activated independently of the PIDDosome through certain death receptor pathways (Droin et al., 2001; Duan and Dixit, 1997; Lavrik et al., 2006; Wagner et al., 2004). Indeed, studies with PIDD-deficient cells suggest that PIDD is not essential for caspase-2 activation (Manzl et al., 2009), suggesting that alternate caspase-2 activation platforms may exist.
Caspase-2 deficient mice have a mild apoptotic phenotype, the defining feature of which is an excess number of oocytes (Bergeron et al., 1998). However, caspase-2 deficient cells have been shown to be more resistant to apoptosis induced by cytoskeletal disruptors, such as vincristine and cytochalasin D (Ho et al., 2008), and heat shock (Tu et al., 2006). DHEA, an inhibitor of the pentose phosphate pathway, has been shown to activate caspase-2 in Xenopus oocytes, suggesting a role in metabolic pathways (Nutt et al., 2005). In neurons, caspase-2 has been shown to be activated during β-amyloid and trophic factor withdrawal-induced apoptosis (Stefanis et al., 1998; Troy et al., 2000) although in the latter case caspase-9 can compensate for the lack of caspase-2 in knockout animals (Troy et al., 2001). Caspase-2 has also been implicated in both p53-dependent (Lassus et al., 2002; Robertson et al., 2002) and p53-independent DNA damage-induced apoptosis (Shi et al., 2009; Sidi et al., 2008). However many of these observations are still considered quite controversial (Krumschnabel et al., 2009).
Consistent with a role in DNA damage, caspase-2 has been shown to be a tumor suppressor (Ho et al., 2009). This may suggest that caspase-2 functions in the nucleus. Indeed, caspase-2 has been observed in the nucleus and it has been suggested that this is the site of its activation (Baliga et al., 2003; Colussi et al., 1998; Paroni et al., 2002). For example, the DNA-PKcs-containing PIDDosome appears to activate caspase-2 in the nucleus following DNA damage (Shi et al., 2009). However, caspase-2 has also been shown to reside in the cytoplasm and at the Golgi complex (Mancini et al., 2000) and may also be activated in these subcellular locations.
A major contributing factor to the paucity of knowledge on the physiological activation of caspase-2 is that current methods are limited in their ability to measure the activation of the initiator caspases. Caspase-2 undergoes autocatalytic processing when it is activated and is also cleaved by caspase-3 (Slee et al., 1999) downstream of MOMP. However, such cleavage does not activate it (Baliga et al., 2004). Thus, monitoring cleavage alone is not sufficient to identify caspase-2 as the apical caspase. Similarly, inhibitors and fluorogenic substrates based on the preferred caspase-2 substrate sequence, VDVAD can respectively inhibit or be cleaved as efficiently by other caspases, potentially leading to spurious results (McStay et al., 2008). Furthermore, none of these methods can elucidate the localization or kinetics of caspase-2 activation. Therefore we sought to use recruitment of caspase-2 to its activation platforms, the proximal step in the caspase-2 pathway (Baliga et al., 2004), as a readout for caspase-2 activation.
In order to measure caspase-2 recruitment to activation platforms accurately, we adapted the Bimolecular Fluorescence Complementation (BiFC) technique. BiFC uses split fluorescent proteins that alone are not fluorescent but when fused to interacting proteins associate to form the fluorescent molecule (Shyu et al., 2006). In this study we have fused caspase-2 to the N-terminal half and to the C-terminal half of Venus fluorescent protein. Following recruitment of caspase-2 to its activation platform, induced proximity of caspase-2 occurs and, consequently, association of the two halves of Venus is enforced. Induced proximity of caspase-2 is required for its dimerization and is measured as increased Venus fluorescence. Using this approach, we characterize the kinetics and localization of caspase-2 activation and reveal an unexpected role for Hsp90 in the regulation of heat stress-induced caspase-2 dimerization.
To study the recruitment of caspase-2 to its activation platforms, its subsequent induced proximity and activation during apoptosis, we used the BiFC assay described in the introduction. We fused the CARD domain of caspase-2 (C2-CARD, aa 1-122) to each of the split Venus proteins, Venus C 155-239 and Venus N 1-173. We transiently expressed the C2-CARD BiFC pair in Hela cells and investigated the ability of the proteins to undergo BiFC upon co-expression of the PIDDosome components, PIDD and RAIDD (Figure 1A, B). Upon PIDD expression, almost half of the cells became Venus-positive indicating induction of BiFC of the C2-CARD pair. Expression of RAIDD induced C2-CARD BiFC in the majority of cells, while neither the CARD containing protein, Apaf-1, nor the death domain containing protein, FADD, induced BiFC above background levels. When RAIDD expression was titrated to levels where only a small amount of BiFC was observed, PIDD was able to synergize with RAIDD to restore C2-CARD BiFC in the majority of cells (Figure 1C, 1D). Thus, we were able to induce BiFC of C2-CARD by reconstituting the PIDDosome, indicating that the BiFC observed represents caspase-2 induced proximity upon recruitment to its activation platform.
We have previously shown that heat shock activates caspase-2 (Tu et al., 2006) and we therefore determined if heat shock can similarly induce BiFC of caspase-2. Following heat shock, cells expressing the C2-CARD BiFC pair became strongly Venus-positive (Figure 2A, 2B). This effect was dose-dependent with respect to the amount of the BiFC pair expressed, while at the maximum amount of plasmid the level of spontaneous Venus re-association remained low.
To determine if the CARD of caspase-2 accurately recapitulated the behavior of the intact protein, we generated Venus C (VC) and Venus N (VN) epitope-tagged full-length caspase-2 (C2-FL, Figure 2C). The catalytic cysteine (C303) in caspase-2 was mutated to an alanine to prevent apoptosis upon expression of the caspase, to facilitate analysis. Caspase-2 contains a classical nuclear localization sequence (NLS) in its prodomain (aa 131-138) that is not present in C2-CARD (Baliga et al., 2003). Thus, we also generated Venus C and Venus N-tagged versions of a fragment of caspase-2 containing the prodomain including the NLS (C2-Pro, aa 1-147). When we expressed each of these BiFC pairs in Hela cells they all induced BiFC following heat shock (Figure 2D and Supplemental Figure S1A) suggesting that the NLS is not required for caspase-2 induced proximity.
We next explored if caspase-2 induced proximity was induced by other apoptotic stimuli, specifically those reported to activate caspase-2. As before, we transiently expressed the C2-CARD BiFC pair in Hela cells and observed induction of BiFC under different conditions (Figure 2E, 2F, Supplemental Figure S2). Treatment with TNF or etoposide, resulted in an increase in the number of Venus-positive cells over background although the cells showed marginal brightness (Figure 2E). The tubulin disruptors, taxol, vincristine and colchicine, also caused an increase in the number of positive cells. These stimuli induced similar levels of BiFC when the C2-Pro or C2-FL pair was expressed under similar conditions (Supplemental Figure S1B, S1C, S1D). In contrast, the actin disruptor cytochalasin B had little effect, while cytochalasin D showed an increase over background (Figure 2F, Supplemental Figure S2). The pentose phosphate pathway inhibitor, DHEA, also induced considerable C2-CARD BiFC (Supplemental Figure S2). Treatment with anti-Fas, which induces substantial apoptosis in Hela cells (data not shown), did not induce BiFC (Supplemental Figure S2), indicating that the induced proximity observed only occurs in response to certain stresses reported to activate caspase-2, including DNA damage, cytoskeletal disruption and metabolic stress (Ho et al., 2008; Nutt et al., 2005; Robertson et al., 2002). Of the conditions analyzed, heat shock resulted in the most robust BiFC response and thus we proceeded to rigorously characterize the regulation of caspase-2 activation induced by heat stress.
Using time-lapse confocal microscopy, we analyzed the kinetics of caspase-2 recruitment to activation platforms in Hela cells expressing the C2-CARD Venus pair during heat shock-induced apoptosis. C2-CARD complexes were detected as fluorescent punctate spots in the cytoplasm as early as 5 hr after heat shock that increased in intensity over time (Figure 3A, Supplemental Movie S1). Similar kinetics were observed when the C2-Pro or C2-FL pair were expressed (Supplemental Figure S3). To clarify the origin and destination of these punctate spots within the cell we increased the resolution of the time-lapse imaging by taking a number of confocal sections through the z-plane of the cell. Analysis of BiFC in the 3D time-lapse showed that the punctate spots originated at the periphery of the cell and translocated to accumulate in a region adjacent to the nucleus (Figure 3B, Supplemental Movie S2). The average intensity of Venus in each cell was measured at each time point, and showed a steady increase in the fluorescence starting at approximately 5 hr after heat shock (Figure 3C). In untreated cells no increase in Venus intensity was observed (Figure 3C). Similarly, cells exposed to a non-lethal heat shock of 42°C failed to induce an appreciable amount of bi-fluorescence. Thus this effect was specific to heat shock at 45°C, the temperature required to induce apoptosis in Hela cells, demonstrating that the kinetic events we observed represent heat shock-induced capase-2 induced proximity in real time.
The results in Figures 3 and S3 suggest that caspase-2 activation platforms occur in the cytosol after heat shock. However, numerous reports suggest that caspase-2 is activated in the nucleus (Baliga et al., 2003; Paroni et al., 2002). Using immunofluorescence, we found that, consistent with previous reports, endogenous caspase-2 is localized to the Golgi apparatus, the nucleus and the cytosol ((Mancini et al., 2000); Supplemental Figure S4). After heat shock, caspase-2 remained associated with the Golgi apparatus, although the Golgi itself became dismantled. To more accurately determine the subcellular compartment where induced proximity of caspase-2 occurs, we transiently expressed each of the BiFC pairs in Hela cells and investigated the subcellular localization of caspase-2 BiFC. Following heat shock the C2-FL, C2-Pro and C2-CARD BiFC pairs each appeared as a series of punctate spots in the cytoplasm (Figure 4A and Supplemental Movies S3-6). The 3D images of the cells clearly show that caspase-2 induced proximity did not occur in the nucleus and was distinctly cytoplasmic for C2-CARD, C2-FL and C2Pro. We observed similar patterns of caspase-2 induced proximity in response to treatment with vincristine, taxol and colchicine (Figure 4B and Supplemental Figure S5) indicating that cytoskeletal disruption also activates caspase-2 in the cytosol. Finally, treatment with etoposide also resulted in a cytosolic rather than nuclear localization of caspase-2 induced proximity (Supplemental Figure S5). These results indicate that caspase-2 activation generally occurs in the cytoplasm rather than the nucleus as detected by this approach.
Given that we observed efficient caspase-2 BiFC in response to heat shock and that we have previously shown that heat shock-induced activation of caspase-2 is RAIDD dependent (Tu et al., 2006), we investigated the dependency of casapse-2 induced proximity on RAIDD. The introduction of two point mutations, D83A and E87A into the CARD domain of caspase-2 has been reported to disrupt the interaction between caspase-2 and RAIDD (Duan and Dixit, 1997). Co-immunoprecipitation experiments showed that these mutations attenuated the binding of caspase-2 to RAIDD rather than completely disrupting it (Figure 5A). Consistent with the weaker binding of C2-CARD to RAIDD, the level of BiFC after heat shock was greatly reduced when the C2-CARD mutant was expressed compared to that of the wild type C2-CARD (Figure 5B, 5C). This result suggested that caspase-2 induced proximity requires its binding to RAIDD. This result also indicates that the BiFC observed is due to a RAIDD-CARD/C2-CARD interaction and not a C2-CARD/C2-CARD interaction.
To further investigate the dependence of caspase-2 activation on its interaction with RAIDD during heat shock, we expressed the C2-CARD, C2-Pro and C2-FL pairs in RAIDD-deficient mouse embryonic fibroblasts (MEF). Similar to our observations in Hela cells, when wild type MEF were subjected to heat shock, the number of Venus-positive cells increased in a dose-dependent manner with respect to the amount of the caspase-2 pair expressed. However, in the absence of RAIDD, the percentage of Venus-positive cells was greatly reduced in each case (Figure 5D, Supplemental Figure S6). When RAIDD was reintroduced into RAIDD-deficient cells by transient transfection, its expression restored the C2-CARD BiFC induced by heat shock to wild type levels (Figure 5E, 5F). Together, these results demonstrate that caspase-2 requires RAIDD for induced proximity. Thus we conclude that the activation platform that recruits caspase-2 after heat shock includes the adaptor protein RAIDD.
The anti-apoptotic proteins Bcl-2 and Bcl-xL are potent inhibitors of heat shock induced apoptosis (Cuende et al., 1993), suggesting that MOMP is required in this pathway. As expected, Hela cells expressing Bcl-xL were protected from heat shock-induced apoptosis (Figure 6A). However, when the C2-CARD pair was expressed in Hela cells expressing Bcl-xL the increase in BiFC intensity over time after heat shock was identical to that observed for Hela cells (Figure 6B). The inability of Bcl-xL to block C2-CARD induced proximity is consistent with previous reports that caspase-2 activation occurs upstream of MOMP (Bonzon et al., 2006; Tu et al., 2006). To formally test this, we fused the catalytic domains (aa 110-436) of caspase-2 to FV (F36V), a modified FK506 binding protein (FKBP). Addition of FKBP to a protein enables conditional dimerization by addition of a ligand containing two FK506 moieties (AP20187) to the medium. Enforced dimerization of caspase-2 induced apoptosis in these cells and this death was completely inhibited by Bcl-xL (Figure 6C). Therefore activation of caspase-2 occurs upon dimerization of the caspase, inducing apoptosis in a Bcl-xL inhibitable manner. Thus, caspase-2 activation is upstream of MOMP and apoptosis.
During apoptosis, the mitochondrial intermembrane space proteins cytochrome c, Omi and Smac are released simultaneously (Munoz-Pinedo et al., 2006). Therefore, to further characterize the temporal relationship between caspase-2 activation and MOMP, we expressed the C2-CARD pair in Hela cells expressing Omi fused to the fluorescent protein mCherry. Using time-lapse microscopy, we compared the onset of C2-CARD BiFC to the release of OmimCherry from the mitochondria. As shown in Figure 6D and 6E, induction of BiFC starts approximately 4 hours after heat shock while the onset of MOMP occurs 6-10 hr later (Supplemental Movie S7). It is of note that not all cells undergo MOMP during the timeframe of the time-lapse experiments although the BiFC is induced in these cells at similar times (in the example shown in Figure 6E, right, MOMP did not occur, despite caspase-2 dimerization). Thus recruitment of caspase-2 to its activation platform alone may not be sufficient to commit the cell to apoptosis, indicating regulatory steps upstream of MOMP.
To identify some of these regulatory elements upstream of MOMP, we further explored the relationship between heat shock-induced stress and caspase-2 activation. To determine if caspase-2 induced proximity was regulated by heat-induced transcription of pro-apoptotic factors, we heat shocked cells expressing the C2-CARD BiFC pair in the presence of the protein synthesis inhibitor cycloheximide. Rather than inhibiting heat shock-induced C2-CARD BiFC, we observed that the same proportion of cells became Venus-positive when cells were heated in the presence and absence of cycloheximide. (Figure 7A). This indicated that caspase-2 induced proximity does not require heat shock-induced transcriptional upregulation of pro-apoptotic proteins to proceed. This suggests that assembly of caspase-2 activation platforms such as the PIDDosome is not due to heat shock-induced expression of its components, such as PIDD or RAIDD.
Nevertheless, it is well established that heat regulates the transcription of the heat shock proteins (Hsps) that can promote cell survival (Li and Werb, 1982; Strasser and Anderson, 1995; Subjeck et al., 1982). Stress-induced expression of Hsps is mediated primarily by the transcription factor heat shock factor-1 (HSF-1), which is protective against lethal heat stress (McMillan et al., 1998). As predicted, HSF-1−/− MEF were much more sensitive to heat shock-induced apoptosis compared with wild type MEF (Figure 7B). We expressed the C2-CARD BiFC pair in HSF-1+/+ and HSF-1−/− cells and subjected them to heat shock. At temperatures that are non-lethal in wild type cells (43°C), a greater proportion of HSF-1-deficient cells became Venus-positive compared to wild type cells (Figure 7C). We observed similar results in cells expressing C2-Pro BiFC or C2-FL BiFC (Supplemental Figure S7A and S7B). This result indicated that HSF-1 inhibits heat shock-induced caspase-2 activation.
One potential means for this inhibition is through HSF-1-mediated expression of a heat shock protein, such as Hsp90α (the inducible form of Hsp90 (Xiao et al., 1999)). To investigate this, we blocked Hsp90 function by adding a pharmalogic inhibitor, 17-(dimethylaminoethylamino)-17-demethoxygeldanamycin (17-DMAG; (Sharp and Workman, 2006)). Treatment of HSF-1+/+ cells expressing the C2-CARD, C2-Pro or C2-FL BiFC pair with 17-DMAG did not lead to a significant increase in Venus-positive cells (Figure 7D, 7E, Supplemental Figure S7A, S7B). However, when we mildly heat stressed HSF-1+/+ cells in the presence of 17-DMAG, the proportion of cells displaying BiFC was comparable to that observed in HSF-1−/− cells subjected to heat shock (Figure 7D, 7E, Supplemental Figure S7A, S7B). In Hela cells we observed that treatment with 17-DMAG alone led to a modest amount of C2-CARD induced proximity (Supplemental Figure S7C). When we combined 17-DMAG with mild heat stress at 43°C, that does not normally induce C2-CARD dimerization, we restored the level of BiFC to that observed when the cells were heat shocked at 45°C. This strongly suggests that activation of HSF-1 by heat induces the expression of Hsp90 that, in turn, can inhibit induced proximity of caspase-2.
To determine directly if Hsp90 can inhibit caspase-2 activation, we investigated the ability of Hsp90 to inhibit processing of caspase-2 in vitro. We heated Jurkat lysates to 37°C to induce spontaneous PIDDosome assembly (Tinel and Tschopp, 2004) in the presence or absence of Hsp90α recombinant protein. Hsp90α completely blocked caspase-2 processing after 30 min and, after 1 hr; only the p31 intermediate fragment was detected while the p19 and p12 fragments that represent complete cleavage were not produced (Supplemental Figure S6D). Therefore, we concluded that Hsp90α is an efficient inhibitor of caspase-2 activation in vitro. As we have noted, cleavage of caspase-2 is not a demonstration that it has been activated. However, these results are consistent with our findings in cells that HSP90α interferes, directly or indirectly, with the activation of caspase-2.
To further investigate if caspase-2 activity is inhibited by Hsp90α, we used RNAi to specifically knockdown Hsp90α. We achieved approximately 50% knockdown of Hsp90α without any effect on Hsp90β (Figure 7F). Under these conditions we found that siRNA mediated knockdown of Hsp90α had a similar effect to 17-DMAG treatment, such that it sensitized cells to formation of the caspase-2 activation platform at 43°C as detected by BiFC (Figure 7G). Together these results strongly suggest that Hsp90α negatively regulates caspase-2 activation.
Detecting initiator caspase activation is both difficult and frequently inaccurate using existing methods. In this study, we took advantage of the fact that initiator caspase activation is driven by induced proximity (Boatright et al., 2003; Read et al., 2002; Salvesen and Dixit, 1999) to develop a real time technique to monitor initiator caspase activation in single cells. Using this approach, we visualized caspase-2 induced proximity in cells in response to heat shock and a number of other stimuli reported to activate caspase-2 (Figure 2).
Of the initiator caspases, the regulation of caspase-2 activation is the least well characterized. However, similar to caspase-8 and caspase-9, it has been shown that cleavage of the caspase is not required for enzymatic activity (Baliga et al., 2004). Since caspase-2 can be cleaved by active executioner caspases (Slee et al., 1999), monitoring the kinetics of cleavage of caspase-2 does not truly reflect its activation state. Therefore we have developed a method that directly measures engagement of caspase-2 by its activation platform and consequently is a specific and accurate method of measuring initiator caspase activation.
The refolding of split Venus fragments and its associated fluorescence is bright, highly photostable (Shyu et al., 2006) and rapid (Hu et al., 2002; Schmidt et al., 2003). This allowed us to accurately pinpoint the onset of caspase-2 induced proximity as well as the subcellular distribution of activated caspase-2. We detected caspase-2 induced proximity as early as five hours after heat shock (Figure 3C). The delay between caspase-2 activation and onset of MOMP varied from 4-10 hours (Figure 6D, 6E). Despite caspase-2 BiFC being detected in the majority of cells, not all of these cells underwent MOMP. This could be due to inefficient cleavage of Bid by caspase-2 compared to caspase-8 (Bonzon et al., 2006). Alternatively, this could result from negative regulation of caspase-2 by the components of the assay system.
Caspase-2 BiFC appeared as fluorescent dots in the cytoplasm that are not associated with mitochondria (Figure 3). These fluorescent structures likely represent caspase-2 activation platforms and are similar to the aggregates observed in the nucleus when caspase-2-GFP is overexpressed, (Baliga et al., 2003). It has been proposed that the latter aggregates may represent caspase-2 recruitment to PML bodies in the nucleus (Sanchez-Pulido et al., 2007; Tang et al., 2005). Alternatively, these aggregates may represent sites of the nuclear PIDDosome that contains DNA-PKcs, which seems to function primarily in the maintenance of a cell cycle checkpoint rather than apoptosis (Shi et al., 2009). If caspase-2 is recruited to such structures, our results show that it is not brought into sufficient proximity to allow BiFC. Furthermore, this complex reportedly does not require RAIDD, while our results clearly show that the recruitment of caspase-2 to activation platforms in the cytoplasm induced by heat shock requires RAIDD (Figure 5).
Many of the aforementioned studies use immunofluorescence methods or overexpression of GFP-tagged caspase-2 to detect the localization of caspase-2. In contrast to the BiFC approach, these techniques cannot distinguish between active and inactive caspase-2. Our results indicated that caspase-2 is not dimerized in the nucleus and that the pattern of caspase-2 BiFC is not dependent on the NLS (Figure 4). We do not rule out, however, that there is an alternative function for caspase-2 in the nucleus that is independent of its recruitment to activation platforms or activation. The aggregates of caspase-2 we observed in the cytosol are likely to be composed of the PIDDosome, providing an activation platform for caspase-2. Supporting this hypothesis, caspase-1 has been shown to co-localize with its adaptor protein ASC to similar cytosolic structures (Stehlik et al., 2003).
Our results indicate that Hsp90 can negatively regulate caspase-2 dimerization in response to heat shock (Figure 7). Heat shock proteins have been implicated as regulators of other initiator caspases. Hsp70 and Hsp90 inhibit apoptosome assembly by binding to Apaf-1 (Beere et al., 2000; Pandey et al., 2000; Saleh et al., 2000). Similarly, Hsp90 binds and inhibits NALP3, a component of the inflammasome that regulates caspase-1 (Mayor et al., 2007). It is not clear how Hsp90 inhibits caspase-2 activation, but based on its proposed mechanism of inhibition of the other complexes, Hsp90 may operate in an analogous way by preventing PIDDosome assembly through binding one of the components of the complex. We detected binding of Hsp90α to PIDD upon exogenous expression (data not shown) but this binding is not direct evidence of a mechanism for Hsp90α-mediated inhibition of caspase-2. It is equally possible that Hsp90α inhibits caspase-2 induced proximity by some other means. A number of Hsp90 client proteins, such as p53 and ChkI (Arlander et al., 2003; Blagosklonny et al., 1996), have been implicated in the regulation of caspase-2 activation (Baptiste-Okoh et al., 2008; Sidi et al., 2008) and thus Hsp90α may inhibit caspase-2 through the regulation of these or other client proteins. Ongoing studies will help to resolve the mechanisms whereby this chaperone influences the activation of caspase-2.
The Hsp90 inhibitor, 17-DMAG, sensitized cells to heat shock-induced caspase-2 activation (Figure 7D, 7E). 17-DMAG binds to and blocks the nucleotide-binding pocket of Hsp90, causing dissociation from and destabilization of its client proteins. 17-DMAG is currently in clinical trials as a potential anti-tumor therapy (Sharp and Workman, 2006). Sensitization to caspase-2-induced apoptosis may contribute to the anti-tumor effects of 17-DMAG. Further characterization of the PIDDosome is required to fully elucidate the role of Hsp90 in the regulation of caspase-2 activation.
These results demonstrate the ability of BiFC to accurately and specifically measure recruitment of initiator caspases to their respective activation platforms. This powerful approach can be used to further dissect the more elusive aspects of caspase-2 regulation during apoptosis. Clearly this approach can also be extended to other CARD containing proteins that may be activated by induced proximity as well as other protein-protein interactions.
Human Embryonic Kidney (HEK) 293T and Hela cells were grown in Dulbecco's Modified Essential Medium (DMEM, GIBCO BRL) and mouse embryonic fibroblasts (MEF) were grown DMEM supplemented with non-essential amino acids, sodium pyruvate and 2-mercaptoethanol (55μM). All media were supplemented with 2mM glutamine, antibiotics and 10% fetal bovine serum (FBS).
To induce apoptosis by heat shock, media on the cells was exchanged for media warmed to the heat shock temperature and cells were placed in an incubator set at the same temperature for one hour. Cells were returned to 37°C for the times indicated. qVD-OPH (20μM, MP Biomedicals) was included to inhibit caspases.
Cells were imaged using a spinning disk confocal microscope (Zeiss). Hela cells were plated on dishes containing coverslips (Mattek) 24 hr prior to treatment. For time-lapse experiments media on the cells was supplemented with Hepes (20mM) and 2-mercaptoethanol (55μM). Cells were allowed to equilibrate to 37°C in 5% CO2 prior to focusing on the cells. Each experiment included control images of untreated cells to ensure that the imaging conditions did not induce phototoxicity and cell division proceeded.
We thank M. Parsons and M. Pinkoski for careful reading of the manuscript and F. Llambi for invaluable discussion, Dr. C-D Hu (Purdue University, Indiana) for BiFC plasmids, Dr. T. Mak (U. Toronto) for RAIDD-deficient mice and Dr I. Benjamin (University of Utah) for HSF-1-deficient MEF. This work was supported by NIH grant AI47891 (DRG).
Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.