The proteasome inhibitor bortezomib (also known as PS-341/Velcade) is a dipeptidyl boronic acid that has recently been approved for use in patients with multiple myeloma. Bortezomib inhibits the activity of the 26S proteasome and induces cell death in a variety of tumor cells; however, the mechanism of cytotoxicity is not well understood. In this report, oligonucleotide microarray analysis of the 8226 multiple myeloma cell line showed a predominant induction of gene products associated with the endoplasmic reticulum secretory pathway following short-term, high-dose exposure to bortezomib. Examination of mediators of endoplasmic reticulum stress–induced cell death showed specific activation of caspase 12, as well as of caspases 8, 9, 7, and 3, and cleavage of bid. Treatment of myeloma cells with bortezomib also showed disregulation of intracellular Ca2+ as a mechanism of caspase activation. Cotreatment with a panel of Ca2+-modulating agents identified the mitochondrial uniporter as a critical regulatory factor in bortezomib cytotoxicity. The uniporter inhibitors ruthenium red and Ru360 prevented caspase activation and bid cleavage, and almost entirely inhibited bortezomib-induced cell death, but had no effect on any other chemotherapeutic drug examined. Additional Ca2+-modulating agents, including 2-amino-ethoxydiphenylborate, 1,2-bis (o-aminophenoxy) ethane-tretraacetic acid (acetoxymethyl) ester, and dantrolene, did not alter bortezomib cytotoxicity. Analysis of intracellular Ca2+ showed that the ruthenium-containing compounds inhibited Ca2+ store loading and abrogated the desensitized capacitative calcium influx associated with bortezomib treatment. These data support the hypothesis that intracellular Ca2+ disregulation is a critical determinant of bortezomib cytotoxicity.

Bortezomib (PS-341/Velcade) is a dipeptide containing boronic acid, which has been recently approved for therapeutic use in refractory multiple myeloma (1, 2). Bortezomib forms a covalent bond with the active site threonine in the core of the 20S proteasome and inhibits the chymotryptic activity of the proteasome; however, its exact mechanism of cytotoxicity and selectivity for transformed cells is not known. A large number of proteins that are involved with carcinogenesis are known to be regulated by the ubiquitin-proteasome system of degradation, including transcription factors such as activator protein-1 and p53, signal transduction molecules such as Jak2 and cbl, the cell cycle regulators p21 and p27, and regulatory factors such as the nuclear factor-κB inhibitors IκBα, β, ε, and p100. Many laboratories have investigated these proteins as potential targets of bortezomib-mediated cytotoxicity, and these studies have clearly shown that bortezomib is a potent inhibitor of the 26S proteasome with effects that can be directly related to protein stabilization (35). However, the molecular switch that initiates cell death in this pathway has not yet been identified.

Over the past 10 years, two primary apoptotic pathways have been described. The “extrinsic pathway” is initiated by ligation of cell-surface death receptors such as CD95 (Fas/Apo-1), tumor necrosis factor receptor 1, and death receptors 4 and 5 (see review in ref. 6). Upon receptor ligation, the adapter protein Fas-associated death domain protein is recruited, which in turn recruits procaspase 8, resulting in the formation of the death-inducing signaling complex. Death-inducing signaling complex formation is generally thought to initiate apoptosis by induced proximity autocatalytic activation of caspase 8 and subsequent downstream effectors. In contrast, the “intrinsic pathway” is associated with various cell damaging agents such as reactive oxygen species and DNA strand breaks, and is initiated by mitochondrial release of cytochrome c, formation of the apoptosome, and activation of procaspase 9. Cross talk between these two pathways occurs through the Bcl-2 family, which includes both proapoptotic and antiapoptotic members (7). For example, bid is a proapoptotic factor that has been shown to be cleaved by caspase 8, releasing an activated fragment, which can induce mitochondrial release of cytochrome c. Bcl-xL is an antiapoptotic factor that has been shown to inhibit cell death induced by both death receptor ligation and cytotoxic drugs in some cell types (8). Thus, bid activation by caspase 8 and mitochondrial release of cytochrome c are thought to amplify extrinsic apoptotic signals by recruiting involvement of the mitochondrial pathway.

More recently, a third pathway has been identified that is initiated by the endoplasmic reticulum (ER; see review in ref. 9). At least two different mechanisms have been associated with ER stress–initiated apoptosis, the unfolded protein response and disregulation of Ca2+ homeostasis.

Disregulation of intracellular Ca2+ was among the first hallmarks of apoptosis, predating the identification of the caspase cascade as a mechanism of programmed cell death (see review in ref. 10). Early work identified a biphasic increase in cytosolic Ca2+ associated with apoptosis: first, a transient spike occurring immediately following the cellular insult; second, sustained Ca2+ influx that was considered to be the lethal event. Subsequent studies have shown that even minor disruptions in either total Ca2+ or subcellular distribution can modulate the apoptotic response to a large number of stimuli. In most cell types, the ER is the primary intracellular store of Ca2+, where it participates in the folding, modification, and sorting of newly synthesized proteins. Homeostasis between Ca2+ stores and cytosolic Ca2+ is maintained by ER-resident channels and transporters. The primary mechanism for ER Ca2+ influx is the smooth ER Ca2+-ATPase, which mediates store filling. Two primary Ca2+-sensitive channels mediate the release of Ca2+ from the ER, the inositol triphosphate receptor (IP3R) and the caffeine sensitive ryanodine receptor. Recent studies suggest that IP3R-activated depletion of ER stores is also involved in the regulation of Ca2+ influx from the extracellular environment, a phenomenon known as capacitative Ca2+ influx (CCI; ref. 11). Vazquez et al. showed that a genetic variant of DT40 B lymphocytes deficient in IP3R expression was unable to transport Ca2+ into the cell and was resistant to apoptosis following B-cell receptor ligation. Transfection of IP3R expression constructs restored this capability; however, the identity of the plasma membrane channel activated by elevated inositol triphosphate and the mechanism of IP3R regulation remain under investigation. Additional studies have also implicated IP3R-mediated CCI as a modulator of apoptosis (1214). Jayaraman et al. showed that transfection of T lymphocytes with IP3R antisense rendered the cells resistant to apoptotic cell death induced by dexamethasone, ionizing radiation, T-cell receptor ligation, and CD95 cross-linking. This resistance could be overcome by pharmacologically increasing cytosolic Ca2+, supporting a role for CCI in apoptosis.

In addition to their function in metabolism and apoptosis, mitochondria are known to participate in Ca2+ homeostasis. Excess cytosolic Ca2+ is taken up into the mitochondria through a low-affinity inner membrane Ca2+ uniporter and released back to the cytosol by three independent mechanisms: reversal of the uniporter, Na+/H+-dependent Ca2+ exchange, or the mitochondrial permeability transition pore opening. Additional studies have shown IP3R-mediated translocation of Ca2+ to mitochondria following treatment with staurosporine or ceramide, resulting in rapid permeability transition pore opening, cytochrome c release, and activation of downstream mediators of apoptosis (15). In these studies, apoptosis could be inhibited by ruthenium red, a cationic dye that blocks the mitochondrial uniporter, or by bongkreikic acid, which binds to the adenine nucleotide translocator and prevents permeability transition pore opening. More recently, several studies have shown a role for the Bcl-2 family of apoptotic regulators in the translocation of Ca2+ from the ER to the mitochondria (16, 17). Collectively, these studies show an additional mechanism of cross talk between the known apoptotic pathways and emphasize the complex coordination of signals that contribute to cell survival or death.

In the present study, we have examined the early events associated with bortezomib-induced cell death of multiple myeloma cells. Multiple myeloma is a malignancy of secretory plasma cells, and as such, they contain a highly developed ER, characteristic of secretory cells. We show that bortezomib treatment induces the expression of proteins associated with the ER secretory pathway, activation of caspase 12, and disregulation of Ca2+ homeostasis leading to cell death. We further show that inhibition of mitochondrial Ca2+ uptake by ruthenium compounds completely abrogates the cytotoxic activity of bortezomib, whereas a series of Ca2+-modulating agents has no significant effect. These results suggest that bortezomib initiates apoptosis in myeloma cells by a unique mechanism that is not known to be activated by any other chemotherapeutic agent.

Cells and antibodies. The 8226, H929, and U266 myeloma cell lines were originally obtained from American Type Culture Collection (Rockville, MD) and maintained in RPMI 1640 (Gibco/Invitrogen, Carlsbad, CA), supplemented with 5% or 10% heat inactivated fetal bovine serum (Hyclone, Logan, UT), 1 mmol/L l-glutamine, and 100 units/mL penicillin/streptomycin (Gibco/Invitrogen). The MM.1S cell line was kindly provided by Dr. Steven Rosen (Northwestern University, Chicago, IL; ref. 18). Caspase 3, 8, 9, and 12 antibodies were obtained from Cell Signaling (Beverly, MA).

Oligonucleotide expression analysis. 8226 myeloma cells were exposed to 50 nmol/L bortezomib for 4 hours and harvested for isolation of RNA using the Qiagen Rneasy protocol followed by Oligotex mRNA kit (Qiagen, Valencia, CA). Three independent experiments were done on consecutive days, and equal quantities of RNA from each experiment were combined for oligonucleotide microarray analysis using the Affymetrix U113A gene chip, which includes oligonucleotides representative of 22,000 gene products. Data were analyzed using GeneChip software as previously described (19). Untreated cells were designated as the baseline and comparison metrics calculated to identify differences between the baseline and the cells treated with bortezomib.

Cytotoxicity assays. Cytotoxicity analysis was done either by Annexin V-FITC/Mito Tracker red (CMXRos, Molecular Probes, Eugene, OR) staining and flow cytometry or by using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) dye reduction assay as previously described (20). For all studies using combinations of a Ca2+ inhibitor plus cytotoxic agent, the inhibitor was added to the total cell population before plating. Cytotoxic activity of the Ca2+-modulating agents alone was determined in preliminary control experiments, and all subsequent assays were done at minimally toxic doses as follows: 2-amino-ethoxydiphenylborate (2-APB), 5 μmol/L; 1,2-bis (o-aminophenoxy) ethane-tretraacetic acid (acetoxymethyl) ester (BAPTA-AM), 10 μmol/L; dantrolene, 10 μmol/L; Ru360, 1.0 μmol/L; and ruthenium red, 0.1 μmol/L. All reagents were obtained from Calbiochem (San Diego, CA) with the exception of ruthenium red, which was obtained from Sigma (St. Louis, MO). Bortezomib (PS-341/Velcade) was kindly provided by Millennium Pharmaceuticals (Cambridge, MA).

Immunoblot analysis. Immunoblotting was done as previously described (19). Briefly, after drug treatment, cells were pelleted by centrifugation, washed once with ice-cold PBS, and lysed in 30 mmol/L HEPES (pH 7.5), 10 mmol/L NaCl, 5 mmol/L MgCl2, 25 mmol/L NaF, 1 mmol/L EGTA, 1% Triton X-100, 10% glycerol with 2 mmol/L NaVO4, 10 μg/mL aprotinin, 10 μg/mL soybean trypsin inhibitor, 25 μg/mL leupeptin, and 2 mmol/L phenylmethylsulfonyl fluoride for 20 minutes on ice. Lysates were cleared by centrifugation and protein was quantitated by bicinchoninic acid assay (Pierce, Rockford, IL). Equal amounts of protein (30-50 μg) were separated by SDS-PAGE electrophoresis, transferred to polyvinylidene difluoride membrane (Bio-Rad, Hercules, CA), probed with the specified antibody, and developed using Pierce Supersignal chemiluminescence substrate.

[Ca2+]i measurements. For [Ca2+]i measurements, cells were attached to #1 round glass coverslips coated with poly-d-lysine (Sigma) or fibronectin (Roche, Indianapolis, IN) with identical results on both substrates. Individual coverslips were incubated in a buffered saline (5 mmol/L KCl, 0.3 mmol/L KH2PO4, 138 mmol/L NaCl, 0.2 mmol/L NaHCO3, 0.3 mmol/L Na2HPO4, 20 mmol/L HEPES, 1.3 mmol/L CaCl2, 0.4 mmol/L MgSO4, and 5.6 mmol/L glucose, pH 7.3, at 37°C) containing 2.5 μmol/L Fura-PE3/AM (TefLabs, Austin, TX) with 0.0025% pluronic acid for 20 minutes at 37°C. The coverslips were then rinsed in HBSS for 20 minutes at 37°C to allow full hydrolysis of the Fura-AM to free acid. The coverslip with the dye-loaded cells was then placed in a 37°C imaging chamber, which is mounted on the stage of an inverted Olympus IX70 microscope (Melville, NY). Light emitted from a 100-W Hg bulb was passed alternately through 340- and 380-nm filters using a filter wheel. The emitted fluorescence was collected using a 40× 1.35 numerical aperture UV-fluor objective then selected using a 10-nm bandpass filter centered at 510 nm and passed to a CCD camera (Photometrics CH-250, Tucson, AZ). The ratio of the light intensity monitored at 340/380 is used as an index for [Ca2+]i. Specific calibration procedures have been previously reported in detail (21). A minimum of 5 cells was analyzed in each experimental condition, and all experiments were repeated at least thrice. In the standard protocol, Ca2+ was removed from the media for 2 minutes before addition of cyclopiazoic acid, then after 7 minutes Ca2+ was added back to analyze the magnitude of CCI. The peak changes in Ca2+ following cyclopiazoic acid and Ca2+ addition were calculated as percent change from the zero Ca2+ baseline (baseline) using the formula [(Rpeak following treatment) − (R immediately before treatment)] / (Rpeak immediately before treatment), where R is the 340/380 ratio. Rate of rise was calculated as [(Rpeak) − (Rbaseline)] / (time at Rpeak − time at Rbaseline). Significance was determined using the Student's two-sided t test.

Bortezomib treatment induces the expression of gene products associated with the endoplasmic reticulum secretory pathway. To investigate the mechanism of bortezomib-mediated cell death, oligonucleotide microarray analysis was used to examine the differential gene expression profile of the 8226 myeloma cell line following exposure to bortezomib. These studies were designed to examine the acute response of myeloma cells to bortezomib. Therefore, myeloma cells were treated with a short-term (4-hour) exposure using a dose corresponding to the approximate IC70 for a 24-hour exposure as determined by MTT and Annexin V-FITC. No phenotypic indications of cell death are visible at this time point. Using untreated cells as baseline, 25 gene products were identified as significantly expressed (fluorescence signal >200) and increased by >2-fold in cells maintained in suspension, whereas only 8 were decreased at the level of RNA expression. The most consistent and dramatic increase occurred in gene products associated with the ER secretory pathway (Table 1). Specifically, two isoforms of the chaperone 70 kDa heat shock protein (hsp70), 1A and 1B, were induced by 2.6- and 2.9-fold, respectively. The hsp70-interacting protein, BCL2-associated athanogene 3, was similarly induced 2.0-fold, and the luminal ER Ca2+-regulating protein, calreticulin, was induced by 2.0-fold. Protein expression of these gene products was confirmed by Western blot analysis (data not shown).

Table 1.

Oligonucleotide analysis of gene expression in myeloma cells following 4 hours of treatment with 50 nmol/L bortezomib

Fold changeGene description
0.3 butyrate response factor 2 (epidermal growth factor-response factor 2) 
0.4 homologue of murine (IFN-inducible protein p78; MX1) 
0.4 multiple membrane spanning receptor TRC8 
0.4 IFN-stimulated protein, 15 kDa (ISG15) 
0.4 IFN-stimulated transcription factor 3, γ (48 kDa; ISGF3G) 
0.5 IFN regulatory factor 7 (IRF7), transcript variant c 
0.5 chromosome 19, cosmid R32184 
0.5 nuclear receptor subfamily 4, group A, member 2 
2.0 BCL2-associated athanogene 3 (BAG3) 
2.0 calreticulin 
2.0 KIAA0648 protein 
2.0 prominin (mouse)-like 1 (PROML1) 
2.1 basic-leucine zipper nuclear factor (JEM-1) 
2.1 IFN-related developmental regulator 1 
2.3 chondroitin sulfate proteoglycan 2 (versican) 
2.3 cDNA DKFZp434M054 
2.3 cDNA FLJ11868 fis, clone HEMBA1006993. 
2.4 chromosome 16 open reading frame 7 
2.4 shc pseudogene, p66 isoform 
2.5 FLJ00032 protein, partial cds 
2.6 heat shock 70 kDa protein 1A (HSPA1A) 
2.7 hypothetical protein FLJ20063 
2.8 TATA box binding protein (TBP)-associated factor, RNA polymerase II (TAF2) 
2.9 VAMP-associated protein of 33 kDa mRNA 
2.9 heat shock 70 kDa protein 1B (HSPA1B) 
3.0 ribosomal protein L39 
3.1 ATP-binding cassette, subfamily F (GCN20), member 2 (ABCF2) 
3.2 hypothetical protein FLJ21032 (FLJ21032) 
3.2 cDNA DKFZp586H0722 
3.7 cDNA FLJ13829 fis, clone THYRO1000625 
3.7 EST pseudogene similar to UBL1 [ubiquitin-like 1 (sentrin)] 
4.8 Sec23-interacting protein p125 
5.9 EST for novel 7 transmembrane receptors 
Fold changeGene description
0.3 butyrate response factor 2 (epidermal growth factor-response factor 2) 
0.4 homologue of murine (IFN-inducible protein p78; MX1) 
0.4 multiple membrane spanning receptor TRC8 
0.4 IFN-stimulated protein, 15 kDa (ISG15) 
0.4 IFN-stimulated transcription factor 3, γ (48 kDa; ISGF3G) 
0.5 IFN regulatory factor 7 (IRF7), transcript variant c 
0.5 chromosome 19, cosmid R32184 
0.5 nuclear receptor subfamily 4, group A, member 2 
2.0 BCL2-associated athanogene 3 (BAG3) 
2.0 calreticulin 
2.0 KIAA0648 protein 
2.0 prominin (mouse)-like 1 (PROML1) 
2.1 basic-leucine zipper nuclear factor (JEM-1) 
2.1 IFN-related developmental regulator 1 
2.3 chondroitin sulfate proteoglycan 2 (versican) 
2.3 cDNA DKFZp434M054 
2.3 cDNA FLJ11868 fis, clone HEMBA1006993. 
2.4 chromosome 16 open reading frame 7 
2.4 shc pseudogene, p66 isoform 
2.5 FLJ00032 protein, partial cds 
2.6 heat shock 70 kDa protein 1A (HSPA1A) 
2.7 hypothetical protein FLJ20063 
2.8 TATA box binding protein (TBP)-associated factor, RNA polymerase II (TAF2) 
2.9 VAMP-associated protein of 33 kDa mRNA 
2.9 heat shock 70 kDa protein 1B (HSPA1B) 
3.0 ribosomal protein L39 
3.1 ATP-binding cassette, subfamily F (GCN20), member 2 (ABCF2) 
3.2 hypothetical protein FLJ21032 (FLJ21032) 
3.2 cDNA DKFZp586H0722 
3.7 cDNA FLJ13829 fis, clone THYRO1000625 
3.7 EST pseudogene similar to UBL1 [ubiquitin-like 1 (sentrin)] 
4.8 Sec23-interacting protein p125 
5.9 EST for novel 7 transmembrane receptors 

NOTE: Gene products known to be associated with the ER secretory pathway are highlighted.

Caspase 12 is activated by bortezomib treatment. One of the mechanisms proposed to contribute to ER stress–mediated death is the activation of the ER-resident caspase 12 (22). Caspase 12 is localized on the cytoplasmic side of the ER. It has been shown to be proteolytically activated by agents that induce ER stress, including tunicamycin, thapsigargin, brefeldin A (23), and m-calpain (24). Manganese(II) (25), β-amyloid protein, and disruption of Ca2+ homeostasis (24) are also reported to activate caspase 12. Examination of caspase 12 activation in 8226 myeloma cells following drug treatment with a variety of cytotoxic agents showed that bortezomib activates caspase 12 in myeloma cells, whereas other cytotoxic agents, including doxorubicin, melphalan, and CD95 ligation, did not (Fig. 1). This is not due to absence of drug-induced cytotoxicity, as analysis of apoptosis by Annexin V-FITC and flow cytometry showed comparable cell death in all drug-treated samples (data not shown). Additionally, examination of additional caspase cleavage products identified the predicted cleavage of caspase 9 in all experimental conditions, caspase 8 by CD95 cross-linking, and caspases 7 and 3 by all cytotoxic agents. Interestingly, caspase 8 was also cleaved by bortezomib treatment, demonstrating the involvement of apoptotic mediators typically associated with the extrinsic pathway.

Figure 1.

Caspase activation in 8226 myeloma cells by various cytotoxic stimuli. 8226 myeloma cells were treated with the indicated agent for 16 hours, harvested, and 30 μg of total protein were analyzed by Western blot for activated caspase cleavage products. Concurrent analysis of cell death by Annexin V-FITC staining showed 50% to 70% apoptosis in all samples (data not shown). Representative of three independent experiments.

Figure 1.

Caspase activation in 8226 myeloma cells by various cytotoxic stimuli. 8226 myeloma cells were treated with the indicated agent for 16 hours, harvested, and 30 μg of total protein were analyzed by Western blot for activated caspase cleavage products. Concurrent analysis of cell death by Annexin V-FITC staining showed 50% to 70% apoptosis in all samples (data not shown). Representative of three independent experiments.

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Caspase 8 modulates the activation of caspase 12. Caspase activation has been shown to occur in a relatively linear sequence, and specific tetrapeptide inhibitors can be used to determine hierarchical ordering. To identify the apical caspase in bortezomib-mediated cytotoxicity, 8226 myeloma cells were treated with 50 nmol/L bortezomib in the presence and absence of the caspase 3 inhibitor DEVD, the caspase 8 inhibitor IETD, or the pan caspase inhibitor Z-VAD. All reagents were FMK conjugated for cell permeability, and Z-FA-FMK was used as a negative control. As shown in Fig. 2, 50 nmol/L bortezomib-induced activation of caspases 12, 3, 8, and 9, and was not inhibited by FA-FMK. The caspase 3 inhibitor DEVD partially prevented the cleavage of caspase 3, but had no effect on caspase 12, 8, or 9, and only slightly inhibited cell death. In contrast, the caspase 8 inhibitor IETD reduced the cleavage of caspases 12, 3, and to a lesser extent, caspase 9, and reduced Annexin V-FITC staining by nearly 50%, suggesting that the activation of caspase 8 is at least one of the primary initiating caspases in bortezomib-mediated cell death.

Figure 2.

Inhibition of bortezomib-mediated caspase cleavage by tetrapeptide inhibitors. 8226 myeloma cells were incubated with 50 nmol/L of the indicated tetrapeptide inhibitor for 30 minutes before 16-hour treatment with 50 nmol/L bortezomib. Cells were harvested and 30 μg of total protein were analyzed by Western blot for caspase cleavage products. Viability was determined by Annexin V-FITC staining and flow cytometric analysis of cell aliquot before lysis. β-Actin is used to show equal protein loading. Representative of three independent experiments.

Figure 2.

Inhibition of bortezomib-mediated caspase cleavage by tetrapeptide inhibitors. 8226 myeloma cells were incubated with 50 nmol/L of the indicated tetrapeptide inhibitor for 30 minutes before 16-hour treatment with 50 nmol/L bortezomib. Cells were harvested and 30 μg of total protein were analyzed by Western blot for caspase cleavage products. Viability was determined by Annexin V-FITC staining and flow cytometric analysis of cell aliquot before lysis. β-Actin is used to show equal protein loading. Representative of three independent experiments.

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Inhibition of mitochondrial Ca2+ uptake prevents bortezomib-induced cytotoxicity. Intracellular Ca2+ is a second messenger that has been associated with caspase 12 activation and regulates a large number of processes, including apoptosis (see review in ref. 10). To determine if disregulation of intracellular Ca2+ contributes to bortezomib-mediated cytotoxicity, drug activity was examined in the presence of a series of Ca2+-interacting agents. These studies were based on the hypothesis that if disregulation of intracellular Ca2+ is required for bortezomib-mediated cell death, then inhibition of intracellular Ca2+ homeostasis using a panel of pharmacologic inhibitors would alter the response of myeloma cells to bortezomib-induced cell death. Calcium release from the ER is regulated by two primary receptors, the IP3R and the ryanodine receptor. The ryanodine receptor can be inhibited by dantrolene, which is used clinically as a muscle relaxant to block Ca2+ release from the sarcoendoplasmic reticulum. 2-APB was originally described as an IP3R inhibitor (26), although more recent data have shown that 2-APB is also a potent blocker of capacitative Ca2+ entry channels independent of its inhibition of the IP3R (27, 28). Free cytosolic Ca2+ can be lowered by incubation with the Ca2+ chelating agent, BAPTA-AM. Finally, mitochondrial uptake of Ca2+ via the mitochondrial uniporter can be inhibited by ruthenium red (29) or its dinuclear analog, Ru360 (30). Preliminary experiments showed that neither ruthenium red nor Ru360 induced significant cell death at concentrations up to 100 μmol/L (data not shown). All other inhibitors were used at doses that resulted in less than 15% cell death in control experiments. 8226 myeloma cells were simultaneously exposed to bortezomib in the presence or absence of minimally toxic doses of each Ca2+-interacting agent, and cytotoxicity was analyzed by MTT dye reduction. As shown in Fig. 3A, cotreatment of 8226 cells with the mitochondrial Ca2+ uptake inhibitor ruthenium red (0.1 μmol/L) or Ru360 (1.0 μmol/L) almost entirely abrogated bortezomib-induced cytotoxicity. Identical results were obtained for additional cell lines, including the myeloma cells H929, U266, and MM.1S, diffuse large cell lymphoma cells DB and Raji, and the mantle cell lymphoma cell line Granta-1 (data not shown). No significant inhibition of bortezomib-mediated cytotoxicity was seen in cells cotreated with agents that inhibit Ca2+ efflux from the ER or with the Ca2+ chelator BAPTA-AM (Fig. 3B). Increasing concentrations of 2-APB up to 100 μmol/L did not produce any additional activity, nor did dantrolene at concentrations up to 50 μmol/L (data not shown). However, preliminary control experiments showed that BAPTA-AM is significantly toxic to myeloma cells, with 30% apoptosis at 5 μmol/L, and it is possible that the concentration used for these experiments may not have been sufficient to entirely deplete cytosolic Ca2+. To rule out the possibility that ruthenium red and Ru360 were affecting the activity of mitochondrial succinate dehydrogenase, which is required for MTT dye reduction, apoptosis was verified by Annexin V-FITC staining and flow cytometry analysis. Identical results were obtained (data not shown). Additionally, 4-day MTT analysis shows that myeloma cells treated with bortezomib in the presence of ruthenium-containing agents continue to proliferate, suggesting that inhibition of the mitochondrial uniporter does not only delay apoptosis but prevents cell death.

Figure 3.

Effects of Ca2+-modulating agents on bortezomib cytotoxicity. A, 8226 myeloma cells were exposed to the indicated concentration of bortezomib in the presence or absence of 0.1 μmol/L ruthenium red or 1.0 μmol/L Ru360 for 24 hours, and cell viability analyzed by MTT dye reduction assay. Data are expressed as (mean absorbance treated / mean absorbance control) × 100, where control is ruthenium red, Ru360, or untreated. B, 24-hour MTT dye reduction analysis of 8226 myeloma cells treated with bortezomib in the presence or absence of 5 μmol/L 2-APB, 10 μmol/L dantrolene, or 5 μmol/L BAPTA-AM. Preliminary experiments showed less than 10% cytotoxicity with 5 μmol/L 2-APB and 10 μmol/L dantrolene. BAPTA-AM induces 30% cytotoxicity at 5 μmol/L. Data are expressed as (mean absorbance treated / mean absorbance control) × 100, where control is 2-APB, dantrolene, BAPTA-AM, or untreated. The effects of ruthenium red (RR) on thapsigargin cytotoxicity were determined exactly as in A. Columns, mean of at least three independent experiments. *, P < 0.05.

Figure 3.

Effects of Ca2+-modulating agents on bortezomib cytotoxicity. A, 8226 myeloma cells were exposed to the indicated concentration of bortezomib in the presence or absence of 0.1 μmol/L ruthenium red or 1.0 μmol/L Ru360 for 24 hours, and cell viability analyzed by MTT dye reduction assay. Data are expressed as (mean absorbance treated / mean absorbance control) × 100, where control is ruthenium red, Ru360, or untreated. B, 24-hour MTT dye reduction analysis of 8226 myeloma cells treated with bortezomib in the presence or absence of 5 μmol/L 2-APB, 10 μmol/L dantrolene, or 5 μmol/L BAPTA-AM. Preliminary experiments showed less than 10% cytotoxicity with 5 μmol/L 2-APB and 10 μmol/L dantrolene. BAPTA-AM induces 30% cytotoxicity at 5 μmol/L. Data are expressed as (mean absorbance treated / mean absorbance control) × 100, where control is 2-APB, dantrolene, BAPTA-AM, or untreated. The effects of ruthenium red (RR) on thapsigargin cytotoxicity were determined exactly as in A. Columns, mean of at least three independent experiments. *, P < 0.05.

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To determine if the mitochondrial uniporter is involved in cell death induced by standard chemotherapeutic drugs, myeloma cells were treated with a variety of cytotoxic agents representing diverse mechanisms of activity in the presence and absence of 0.1 μmol/L ruthenium red. Cotreatment with ruthenium red has no effect on the cytotoxic activity of the topoisomerase II inhibitors doxorubicin and mitoxantrone, alkylating agents melphalan and cis-platinum, or the mitochondrial targeting agents imexon (31) and dexamethasone (data not shown). Additionally, the cytotoxic effect of thapsigargin, which is an irreversible inhibitor of the smooth ER Ca2+-ATPase and induces Ca2+ store depletion, is not inhibited by ruthenium red (Fig. 3B). These data show the dramatic specificity of the inhibitory effects of ruthenium-containing agents on bortezomib-induced death and suggest that the critical determinant of bortezomib cytotoxicity is the mitochondrial-dependent disregulation of Ca2+.

Mitochondrial Ca2+ disregulation occurs upstream of caspase activation. To determine if mitochondrial Ca2+ disregulation was required to initiate caspase activation or, conversely, if caspase activity was required for Ca2+ disregulation, caspase activity was examined in the presence of the Ca2+-modulating agents. 8226 myeloma cells were cotreated with 50 nmol/L PS-341 in the presence or absence of 2-APB, BAPTA-AM, or ruthenium red and examined for caspase activation. Similar to Fig. 1, bortezomib alone induces significant caspase activation (Fig. 4). Inhibition of the mitochondrial Ca2+ uniporter with 0.1 μmol/L ruthenium red entirely prevented activation of caspases 12, 8, and 3 and cleavage of bid. Cotreatment with 1.0 μmol/L Ru360 gave identical results (data not shown). In contrast, 2-APB and BAPTA-AM had no effect. These data further support the hypothesis that mitochondrial Ca2+ is the critical determinant of bortezomib-mediated cytotoxicity. Furthermore, mitochondrial Ca2+ disregulation occurs before caspase activation, including caspase 8, which is typically considered upstream of mitochondrial permeability transition.

Figure 4.

Effects of Ca2+-modulating agents on caspase activation. 8226 myeloma cells were treated for 16 hours with 50 nmol/L bortezomib in the presence or absence of 5 μmol/L 2-APB, 5 μmol/L BAPTA-AM, or 0.1 μmol/L ruthenium red, harvested, and 30 μg of total protein were analyzed by Western blot for caspase cleavage products. Viability was determined by Annexin V-FITC staining and flow cytometric analysis of cell aliquot before lysis. Representative of three independent experiments.

Figure 4.

Effects of Ca2+-modulating agents on caspase activation. 8226 myeloma cells were treated for 16 hours with 50 nmol/L bortezomib in the presence or absence of 5 μmol/L 2-APB, 5 μmol/L BAPTA-AM, or 0.1 μmol/L ruthenium red, harvested, and 30 μg of total protein were analyzed by Western blot for caspase cleavage products. Viability was determined by Annexin V-FITC staining and flow cytometric analysis of cell aliquot before lysis. Representative of three independent experiments.

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Ca2+ homeostasis is disregulated following bortezomib treatment. To identify the effectors of Ca2+ homeostasis that may be contributing to bortezomib-mediated cytotoxicity, imaging of cytosolic Ca2+ was done using the ratiometric dye Fura-AM. Initial studies examining the acute affects of bortezomib exposure showed that bortezomib induces a rapid and transient increase in cytosolic Ca2+, which quickly recovers to basal levels (Fig. 5A). Cytosolic Ca2+ peaked within 5 to 8 minutes of treatment, with complete recovery to baseline occurring by 15 minutes in all cells analyzed. This transient increase in [Ca2+]i following bortezomib treatment may be due to release of Ca2+ from intracellular stores and/or influx from the extracellular space. Subsequent addition of cyclopiazoic acid, to induce store unloading, elicited another large transient increase in [Ca2+]i, indicating that significant levels of Ca2+ remained within the stores following bortezomib exposure (Fig. 5A).

Figure 5.

Effects of bortezomib on Ca2+ homeostasis. A, acute effects of bortezomib on cytosolic Ca2+. 8226 myeloma cells were immobilized on coverslips and loaded with Fura-2 ratiometric dye before treatment with 50 or 500 nmol/L bortezomib. Following recovery from the bortezomib response, 5 μmol/L ionomycin was added to equilibrate Ca2+ across the plasma membrane and to obtain a measure of Rmax. Representative time course of response of a single cell treated with 500 nmol/L bortezomib. The experiment was repeated thrice, with a minimum of 5 cells analyzed in each experiment. Similar data were obtained with 50 nmol/L bortezomib. B, effects of 2-hour bortezomib treatment on Ca2+ homeostasis. In the absence of external Ca2+ and the presence of Ca2+ efflux inhibitors, 30 μmol/L cyclopiazoic acid was added to assess the level of stored Ca2+. As shown in B, the extracellular media was exchanged for Ca2+- and Na+-free HBSS supplemented with 1 μmol/L vanadate to block Ca2+ influx and efflux pathways. After 2 minutes following the reduction in cytosolic Ca2+, 30 μmol/L cyclopiazoic acid (CPA) was added to release Ca2+ from internal stores. Following recovery from the cyclopiazoic acid–induced increase in [Ca2+]i, an equal volume of HBSS containing 2.6 mmol/L Ca2+ was added to the chamber, resulting in a physiologic extracellular Ca2+ concentration of 1.3 mmol/L, and the CCI was recorded. At the conclusion of the experiment, 5 μmol/L ionomycin was added to determine the Rmax. Points, mean of at least 5 cells analyzed in three independent experiments; bars, SE. No differences were seen in cells that adhered to fibronectin compared with those that adhered to poly-d-lysine. Control: dashed line; Bortezomib: solid line.

Figure 5.

Effects of bortezomib on Ca2+ homeostasis. A, acute effects of bortezomib on cytosolic Ca2+. 8226 myeloma cells were immobilized on coverslips and loaded with Fura-2 ratiometric dye before treatment with 50 or 500 nmol/L bortezomib. Following recovery from the bortezomib response, 5 μmol/L ionomycin was added to equilibrate Ca2+ across the plasma membrane and to obtain a measure of Rmax. Representative time course of response of a single cell treated with 500 nmol/L bortezomib. The experiment was repeated thrice, with a minimum of 5 cells analyzed in each experiment. Similar data were obtained with 50 nmol/L bortezomib. B, effects of 2-hour bortezomib treatment on Ca2+ homeostasis. In the absence of external Ca2+ and the presence of Ca2+ efflux inhibitors, 30 μmol/L cyclopiazoic acid was added to assess the level of stored Ca2+. As shown in B, the extracellular media was exchanged for Ca2+- and Na+-free HBSS supplemented with 1 μmol/L vanadate to block Ca2+ influx and efflux pathways. After 2 minutes following the reduction in cytosolic Ca2+, 30 μmol/L cyclopiazoic acid (CPA) was added to release Ca2+ from internal stores. Following recovery from the cyclopiazoic acid–induced increase in [Ca2+]i, an equal volume of HBSS containing 2.6 mmol/L Ca2+ was added to the chamber, resulting in a physiologic extracellular Ca2+ concentration of 1.3 mmol/L, and the CCI was recorded. At the conclusion of the experiment, 5 μmol/L ionomycin was added to determine the Rmax. Points, mean of at least 5 cells analyzed in three independent experiments; bars, SE. No differences were seen in cells that adhered to fibronectin compared with those that adhered to poly-d-lysine. Control: dashed line; Bortezomib: solid line.

Close modal

To investigate the potential mechanism for the bortezomib effects on [Ca2+]i, 8226 cells were incubated with 500 nmol/L PS-341 for 2 hours before analysis of [Ca2+]i. At this time following bortezomib treatment, mitochondrial membrane potential remains intact, as determined by control experiments using the mitochondrial dye CMXRos as well as by Western blotting for cytochrome c release (data not shown). A standard cyclopiazoic acid response protocol was used to determine if store Ca2+ was altered following bortezomib treatment. Plasma membrane influx and efflux of Ca2+ must be eliminated to effectively compare stored Ca2+ in control and treated cells. To do this, cells were perfused in Ca2+- and Na+-free HBSS with 1 μmol/L vanadate to block Ca2+ influx and the activities of the Na+-Ca2+ exchanger and plasma membrane Ca2+-ATPase, respectively. Treatment of 8226 cells with bortezomib for 2 hours had no significant effect on the level of stored Ca2+ (Figs. 5B and 6A), again indicating that if bortezomib acutely altered store Ca2+, this was a transient effect.

Figure 6.

Analysis of the regulation of [Ca2+]i following 2 hours of bortezomib treatment. Myeloma cells were treated with 500 nmol/L bortezomib in the presence or absence of 1.0 μmol/L Ru360 for 2 hours before loading with Fura-2 dye and fluorescence analysis (see Materials and Methods). A, change in Ca2+ recorded after addition of cyclopiazoic acid. B, peak [Ca2+]i attained following elevation of extracellular Ca2+ to 1.6 mmol/L (CCI). Intracellular Ca2+ concentration is calculated based on a standard curve as previously described. C, rate of rise, CCI response. Data are calculated as [(Rpeak) − (Rbaseline)] / (time at Rpeak − time at Rbaseline).

Figure 6.

Analysis of the regulation of [Ca2+]i following 2 hours of bortezomib treatment. Myeloma cells were treated with 500 nmol/L bortezomib in the presence or absence of 1.0 μmol/L Ru360 for 2 hours before loading with Fura-2 dye and fluorescence analysis (see Materials and Methods). A, change in Ca2+ recorded after addition of cyclopiazoic acid. B, peak [Ca2+]i attained following elevation of extracellular Ca2+ to 1.6 mmol/L (CCI). Intracellular Ca2+ concentration is calculated based on a standard curve as previously described. C, rate of rise, CCI response. Data are calculated as [(Rpeak) − (Rbaseline)] / (time at Rpeak − time at Rbaseline).

Close modal

Depletion of Ca2+ stores with agents such as thapsigargin or cyclopiazoic acid activates store-operated channels, and thereby CCI. To examine the extent of CCI following treatment with cyclopiazoic acid, the media Ca2+ was increased to 1.3 mmol/L in the presence of Ca2+ efflux inhibitors and [Ca2+]i was analyzed. As shown in Fig. 5B (t = 11-14 min) and Fig. 6B and C, both the magnitude and rate of CCI increased in bortezomib-treated cells compared with controls (Fig. 6C), suggesting that bortezomib alters either the number of store-operated Ca2+ channels or the length of time such channels remain open following store depletion.

The effect of inhibiting the mitochondrial Ca2+ uniporter on the [Ca2+]i responses to cyclopiazoic acid and the subsequent CCI were evaluated by incubating cells with ruthenium-containing compounds before analysis of [Ca2+]i responses. Incubation of cells with Ru360 in the presence of Ca2+ efflux inhibitors significantly blunted the cyclopiazoic acid response in both control and bortezomib-treated cells, suggesting a role for mitochondria in regulating the efflux from or loading of Ca2+ into the ER (Fig. 6). Ru360 also significantly and substantially decreased the rate and magnitude of CCI in both untreated controls and bortezomib-treated cells (Fig. 6). Moreover, the peak [Ca2+]i attained in cells treated with bortezomib in the presence of Ru360 was significantly less than that seen with bortezomib alone (Fig. 6B). The measured [Ca2+]i following CCI in myeloma cells exposed to 500 nmol/L bortezomib attained mean levels of 0.73 μmol/L (SD 0.089) compared with 0.56 μmol/L (SD 0.043) in control cells (P = 0.05). Concurrent treatment with Ru360 abrogated the [Ca2+]i to 0.49 μmol/L (SD 0.053) with bortezomib exposure or 0.44 μmol/L (SD 0.032) in control cells, suggesting that inhibition of the mitochondrial uniporter normalized the effect of bortezomib on the overall Ca2+ fluxes. These data are in accordance with previous studies demonstrating that mitochondria are an active participant in the maintenance of cellular Ca2+ homeostasis and in the activation and extent of CCI (10).

The boronic acid dipeptide bortezomib is a small molecule inhibitor of the 20S proteasome that has shown antitumor activity in a number of cell types, and is particularly effective in the treatment of patients with multiple myeloma. The findings presented here show a unique apoptotic pathway initiated by bortezomib that uses Ca2+ to activate both intrinsic and extrinsic mediators.

The ubiquitin-proteasome system of protein degradation plays an essential role in quality control of newly synthesized proteins. The data presented here show that bortezomib induces the expression of proteins associated with the ER secretory pathway, indicating a potential ER stress–initiated pathway to apoptosis. While these studies were in progress, two additional studies reported gene expression profiling in cells treated with proteasome inhibitors. Fleming et al. (32) examined the profile of Saccharomyces cerevisiae treated with bortezomib or β-lactone, whereas Mitsiades et al. (33) reported the profile of the human multiple myeloma cell line MM.1S following exposure to 100 nmol/L bortezomib. These data are highly compatible; as in all three studies, the most significant and dramatic induction is that of proteins associated with the ER secretory pathway, suggesting that the mechanism of bortezomib cytotoxicity involves ER stress.

As with many cellular stress responses, if the level of stress exceeds the adaptive capacity of the cell, an apoptotic pathway is initiated. ER stress–mediated apoptosis has been associated with the activation of caspase 12 (23, 34, 35), although the existence of caspase 12 in humans is somewhat controversial. Caspase 12 was originally identified in a murine system, and Fischer et. al. (36) reported a human caspase 12 sequence that is predicted to encode multiple splice variants and stop codons. Antibodies directed against a conserved region of the mouse protein identify a human cellular protein that is of similar relative molecular mass with appropriately sized cleavage fragments (23, 34). In our system, we cannot rule out the possibility that this anti–caspase 12 antibody may cross-react with another as yet unidentified caspase, however, the activation of this caspase is clearly restricted to ER stress–initiated apoptosis. Several mechanisms of caspase 12 activation have been identified, and would seem to be cell type specific (37). The present study shows that bortezomib-mediated activation of caspase 12 may be modulated by caspase 8, but is dependent on mitochondrial Ca2+ disregulation. Our findings also show mitochondrial Ca2+ influx as an event upstream of caspase activation and bid cleavage. Inhibition of the mitochondrial uniporter with ruthenium-containing compounds entirely abrogated bortezomib-induced caspase activation. In contrast, Darios et al. (38) showed that mitochondrial Ca2+ uptake is a consequence of these events in a model system of ceramide-mediated apoptosis in neuronally differentiated PC12 cells. Additionally, in this study, they were unable to inhibit ceramide-induced cell death with ruthenium-containing compounds, suggesting that the mechanism of Ca2+ disregulation incurred by ceramide is distinct from the cellular response to proteasome inhibition.

The regulation of Ca2+ signaling and Ca2+-dependent proteins has long been associated with apoptosis, but only recently have their roles been defined and the regulatory mechanisms begun to be identified. The regulation of apoptosis by Ca2+ signaling was first shown with the identification of a Ca2+/Mg2+-dependent endonuclease responsible for the DNA fragmentation, which is often considered the hallmark of apoptosis (39). Increased cytosolic Ca2+ concentration has been found to subsequently activate Ca2+-dependent enzymes that modulate cell death, including μ-calpain and calcineurin (40). Furthermore, Ca2+ influx into mitochondria, if substantial, can activate cytochrome c release, which subsequently activates caspase activity (15). The lumen of the ER is the primary storage location for Ca2+, where it is either free or bound to luminal proteins such as calreticulin, an unfolded protein-responsive chaperone that we found to be induced by 2-fold following bortezomib treatment. In this respect, Arnaudeau et al. (41) showed that increased expression of calreticulin resulted in an increased rate of agonist-induced Ca2+ release and reduced mitochondrial Ca2+ retention. Although a functional role for calreticulin in bortezomib-mediated cell death has not yet been defined, the observed elevation of mRNA and protein levels suggests that calreticulin expression may be induced as a cellular response to ER stress, and therefore alterations in ER Ca2+ storage or release may play a role in the observed disregulation of Ca2+ homeostasis. However, the data shown in Figs. 5 and 6 clearly show that releasable store Ca2+ levels are not substantially different in cells treated with bortezomib. The primary difference in Ca2+ homeostasis observed in bortezomib-treated cells was a significant elevation in cyclopiazoic acid–activated CCI (Figs. 5 and 6), indicating an increased potential for Ca2+ influx following store release. Interestingly, inhibition of mitochondrial Ca2+ uptake by ruthenium red significantly reduced ER Ca2+ loading and the bortezomib enhancement of store-activated influx. The mechanism by which depletion of ER Ca2+ activates plasma membrane channels is unclear; however, one model suggests that mitochondria play an important role in regulating both the loading state of Ca2+ stores (42) and the coupling of store Ca2+ released for channel activation (43). We propose a model where bortezomib evokes a transient release of Ca2+ stores leading to mitochondrial Ca2+ influx. Mitochondrial Ca2+ sensors associated with the uniporter initiate CCI, which is enhanced in bortezomib-treated cells (Fig. 6), leading to caspase activation. Ruthenium compounds would then be protective by blocking mitochondrial loading and CCI activation and the Ca2+-dependent signal transduction pathways that initiate cell death (44). The identity of the signal pathways involved in this regulation remains under intense investigation. Further definition of the mediators in this pathway will promote the design of strategies to enhance drug activity, reduce toxicity, and overcome drug resistance.

Grant support: National Cancer Institute grant CA17094 (R.T. Dorr), Arizona Cancer Center Core Grant CA23074, American Cancer Society grant IRG7400128 (T.H. Landowski), and Howard Hughes Medical Institute grant 52003749 to the University of Arizona (C. Megli).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank the H. Lee Moffitt Microarray Core Facility (CA76292).

1
Adams J, Palombella VJ, Sausville EA, et al. Proteasome inhibitors: a novel class of potent and effective antitumor agents.
Cancer Res
1999
;
59
:
2615
–22.
2
Richardson P. Clinical update: proteasome inhibitors in hematologic malignancies.
Cancer Treat Rev
2003
;
29
Suppl 1:
33
–9.
3
An WG, Hwang SG, Trepel JB, Blagosklonny MV. Protease inhibitor-induced apoptosis: accumulation of wt p53, p21WAF1/CIP1, and induction of apoptosis are independent markers of proteasome inhibition.
Leukemia
2000
;
14
:
1276
–83.
4
Hideshima T, Mitsiades C, Akiyama M, et al. Molecular mechanisms mediating antimyeloma activity of proteasome inhibitor PS-341.
Blood
2003
;
101
:
1530
–4.
5
Ling YH, Liebes L, Jiang JD, et al. Mechanisms of proteasome inhibitor PS-341-induced G(2)-M-phase arrest and apoptosis in human non-small cell lung cancer cell lines.
Clin Cancer Res
2003
;
9
:
1145
–54.
6
Thorburn A. Death receptor-induced cell killing.
Cell Signal
2004
;
16
:
139
–44.
7
Kuwana T, Newmeyer DD. Bcl-2 family proteins and the role of mitochondria in apoptosis.
Curr Opin Cell Biol
2003
;
15
:
691
–9.
8
Oshiro MM, Landowski TH, Catlett-Falcone R, et al. Inhibition of JAK kinase activity enhances Fas-mediated apoptosis but reduces cytotoxic activity of topoisomerase II inhibitors in U266 myeloma cells.
Clin Cancer Res
2001
;
7
:
4262
–71.
9
Ferri KF, Kroemer G. Organelle-specific initiation of cell death pathways.
Nat Cell Biol
2001
;
3
:
E255
–63.
10
Orrenius S, Zhivotovsky B, Nicotera P. Regulation of cell death: the calcium-apoptosis link.
Nat Rev Mol Cell Biol
2003
;
4
:
552
–65.
11
Vazquez G, Wedel BJ, Bird GS, Joseph SK, Putney JW. An inositol 1,4,5-trisphosphate receptor-dependent cation entry pathway in DT40 B lymphocytes.
EMBO J
2002
;
21
:
4531
–8.
12
Hajnoczky G, Csordas G, Madesh M, Pacher P. Control of apoptosis by IP(3) and ryanodine receptor driven calcium signals.
Cell Calcium
2000
;
28
:
349
–63.
13
Boehning D, Patterson RL, Sedaghat L, Glebova NO, Kurosaki T, Snyder SH. Cytochrome c binds to inositol (1,4,5) trisphosphate receptors, amplifying calcium-dependent apoptosis.
Nat Cell Biol
2003
;
5
:
1051
–61.
14
Jayaraman T, Marks AR. T cells deficient in inositol 1,4,5-trisphosphate receptor are resistant to apoptosis.
Mol Cell Biol
1997
;
17
:
3005
–12.
15
Szalai G, Krishnamurthy R, Hajnoczky G. Apoptosis driven by IP(3)-linked mitochondrial calcium signals.
EMBO J
1999
;
18
:
6349
–61.
16
Scorrano L, Oakes SA, Opferman JT, et al. BAX and BAK regulation of endoplasmic reticulum Ca2+: a control point for apoptosis.
Science
2003
;
300
:
135
–9.
17
Zong WX, Li C, Hatzivassiliou G, et al. Bax and Bak can localize to the endoplasmic reticulum to initiate apoptosis.
J Cell Biol
2003
;
162
:
59
–69.
18
Moalli PA, Pillay S, Krett NL, Rosen ST. Alternatively spliced glucocorticoid receptor messenger RNAs in glucocorticoid-resistant human multiple myeloma cells.
Cancer Res
1993
;
53
:
3877
–9.
19
Landowski TH, Olashaw NE, Agrawal D, Dalton WS. Cell adhesion-mediated drug resistance (CAM-DR) is associated with activation of NF-κB (RelB/p50) in myeloma cells.
Oncogene
2003
;
22
:
2417
–21.
20
Landowski TH, Shain KH, Oshiro MM, Buyuksal I, Painter JS, Dalton WS. Myeloma cells selected for resistance to CD95-mediated apoptosis are not cross-resistant to cytotoxic drugs: Evidence for independent mechanisms of caspase activation.
Blood
1999
;
94
:
265
–74.
21
Martinez-Zaguilan R, Tompkins LS, Gillies RJ, Lynch RM. Simultaneous analysis of intracellular pH and Ca2+ from cell populations.
Methods Mol Biol
1999
;
114
:
287
–306.
22
Rao RV, Ellerby HM, Bredesen DE. Coupling endoplasmic reticulum stress to the cell death program.
Cell Death Differ
2004
;
11
:
372
–80.
23
Rao RV, Castro-Obregon S, Frankowski H, et al. Coupling endoplasmic reticulum stress to the cell death program: An Apaf-1-independent intrinsic pathway.
J Biol Chem
2002
;
277
:
21836
–42.
24
Nakagawa T, Zhu H, Morishima N, et al. Caspase-12 mediates endoplasmic-reticulum-specific apoptosis and cytotoxicity by amyloid-β.
Nature
2000
;
403
:
98
–103.
25
Oubrahim H, Chock PB, Stadtman ER. Manganese(II) induces apoptotic cell death in NIH3T3 cells via a caspase-12-dependent pathway.
J Biol Chem
2002
;
277
:
20135
–8.
26
Sugawara H, Kurosaki M, Takata M, Kurosaki T. Genetic evidence for involvement of type 1, type 2 and type 3 inositol 1,4,5-trisphosphate receptors in signal transduction through the B-cell antigen receptor.
EMBO J
1997
;
16
:
3078
–88.
27
Peppiatt CM, Collins TJ, Mackenzie L, et al. 2-Aminoethoxydiphenyl borate (2-APB) antagonises inositol 1,4,5-trisphosphate-induced calcium release, inhibits calcium pumps and has a use-dependent and slowly reversible action on store-operated calcium entry channels.
Cell Calcium
2003
;
34
:
97
–108.
28
Bootman MD, Collins TJ, Mackenzie L, Roderick HL, Berridge MJ, Peppiatt CM. 2-Aminoethoxydiphenyl borate (2-APB) is a reliable blocker of store-operated Ca2+ entry but an inconsistent inhibitor of InsP3-induced Ca2+ release.
FASEB J
2002
;
16
:
1145
–50.
29
Bae JH, Park JW, Kwon TK. Ruthenium red, inhibitor of mitochondrial Ca2+ uniporter, inhibits curcumin-induced apoptosis via the prevention of intracellular Ca2+ depletion and cytochrome c release.
Biochem Biophys Res Commun
2003
;
303
:
1073
–9.
30
Matlib MA, Zhou Z, Knight S, et al. Oxygen-bridged dinuclear ruthenium amine complex specifically inhibits Ca2+ uptake into mitochondria in vitro and in situ in single cardiac myocytes.
J Biol Chem
1998
;
273
:
10223
–31.
31
Dvorakova K, Payne CM, Landowski TH, Tome ME, Halperin DS, Dorr RT. Imexon activates an intrinsic apoptosis pathway in RPMI8226 myeloma cells.
Anticancer Drugs
2002
;
13
:
1031
–42.
32
Fleming JA, Lightcap ES, Sadis S, Thoroddsen V, Bulawa CE, Blackman RK. Complementary whole-genome technologies reveal the cellular response to proteasome inhibition by PS-341.
Proc Natl Acad Sci U S A
2002
;
99
:
1461
–6.
33
Mitsiades N, Mitsiades CS, Poulaki V, et al. Molecular sequelae of proteasome inhibition in human multiple myeloma cells.
Proc Natl Acad Sci U S A
2002
;
99
:
14374
–9.
34
Yoneda T, Imaizumi K, Oono K, et al. Activation of caspase-12, an endoplasmic reticulum (ER) resident caspase, through tumor necrosis factor receptor-associated factor 2-dependent mechanism in response to the ER stress.
J Biol Chem
2001
;
276
:
13935
–40.
35
Nakagawa T, Yuan J. Cross-talk between two cysteine protease families. Activation of caspase-12 by calpain in apoptosis.
J Cell Biol
2000
;
150
:
887
–94.
36
Fischer H, Koenig U, Eckhart L, Tschachler E. Human caspase 12 has acquired deleterious mutations.
Biochem Biophys Res Commun
2002
;
293
:
722
–6.
37
Lamkanfi M, Kalai M, Vandenabeele P. Caspase-12: an overview.
Cell Death Differ
2004
;
11
:
365
–8.
38
Darios F, Lambeng N, Troadec JD, Michel PP, Ruberg M. Ceramide increases mitochondrial free calcium levels via caspase 8 and Bid: role in initiation of cell death.
J Neurochem
2003
;
84
:
643
–54.
39
Wyllie AH. Glucocorticoid-induced thymocyte apoptosis is associated with endogenous endonuclease activation.
Nature
1980
;
284
:
555
–6.
40
Wang HG, Pathan N, Ethell IM, et al. Ca2+-induced apoptosis through calcineurin dephosphorylation of BAD.
Science
1999
;
284
:
339
–43.
41
Arnaudeau S, Frieden M, Nakamura K, Castelbou C, Michalak M, Demaurex N. Calreticulin differentially modulates calcium uptake and release in the endoplasmic reticulum and mitochondria.
J Biol Chem
2002
;
277
:
46696
–705.
42
Arnaudeau S, Kelley WL, Walsh JV Jr, Demaurex N. Mitochondria recycle Ca2+ to the endoplasmic reticulum and prevent the depletion of neighboring endoplasmic reticulum regions.
J Biol Chem
2001
;
276
:
29430
–39.
43
Malli R, Frieden M, Osibow K, et al. Sustained Ca2+ transfer across mitochondria is essential for mitochondrial Ca2+ buffering, store-operated Ca2+ entry, and Ca2+ store refilling.
J Biol Chem
2003
;
278
:
44769
–79.
44
Glitsch MD, Bakowski D, Parekh AB. Store-operated Ca2+ entry depends on mitochondrial Ca2+ uptake.
EMBO J
2002
;
21
:
6744
–54.