Bax Inhibitor-1-Mediated Inhibition of Mitochondrial Ca2+ Intake Regulates Mitochondrial Permeability Transition Pore Opening and Cell Death

Lee, Geum-Hwa; Lee, Hwa-Young; Li, Bo; Kim, Hyung-Ryong; Chae, Han-Jung

doi:10.1038/srep05194

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Published: 05 June 2014

Bax Inhibitor-1-Mediated Inhibition of Mitochondrial Ca²⁺ Intake Regulates Mitochondrial Permeability Transition Pore Opening and Cell Death

Geum-Hwa Lee¹,
Hwa-Young Lee¹,
Bo Li¹,
Hyung-Ryong Kim² &
…
Han-Jung Chae¹

Scientific Reports volume 4, Article number: 5194 (2014) Cite this article

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Abstract

A recently studied endoplasmic reticulum (ER) stress regulator, Bax inhibitor-1 (BI-1) plays a regulatory role in mitochondrial Ca²⁺ levels. In this study, we identified ER-resident and mitochondria-associated ER membrane (MAM)-resident populations of BI-1. ER stress increased mitochondrial Ca²⁺ to a lesser extent in BI-1–overexpressing cells (HT1080/BI-1) than in control cells, most likely as a result of impaired mitochondrial Ca²⁺ intake ability and lower basal levels of intra-ER Ca²⁺. Moreover, opening of the Ca²⁺-induced mitochondrial permeability transition pore (PTP) and cytochrome c release were regulated by BI-1. In HT1080/BI-1, the basal mitochondrial membrane potential was low and also resistant to Ca²⁺ compared with control cells. The activity of the mitochondrial membrane potential-dependent mitochondrial Ca²⁺ intake pore, the Ca²⁺ uniporter, was reduced in the presence of BI-1. This study also showed that instead of Ca²⁺, other cations including K⁺ enter the mitochondria of HT1080/BI-1 through mitochondrial Ca²⁺-dependent ion channels, providing a possible mechanism by which mitochondrial Ca²⁺ intake is reduced, leading to cell protection. We propose a model in which BI-1–mediated sequential regulation of the mitochondrial Ca²⁺ uniporter and Ca²⁺-dependent K⁺ channel opening inhibits mitochondrial Ca²⁺ intake, thereby inhibiting PTP function and leading to cell protection.

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Introduction

The endoplasmic reticulum (ER) contains a high concentration of Ca²⁺ in the millimolar range^1,2; other organelles, such as mitochondria, also contain Ca²⁺, but only in the micromolar range³. In normal or adaptive conditions, Ca²⁺ is released from the ER and transferred to the mitochondria, in which the fine modulation of Ca²⁺ homeostasis plays a fundamental role in normal mitochondria physiology^4,5. However, abnormal Ca²⁺ efflux from the ER and Ca²⁺ accumulation in the mitochondria are linked to the effects of apoptotic stimuli, including ER stress⁶. In addition, mitochondrial Ca²⁺ levels and opening of the mitochondrial membrane permeability transition pore (PTP) have recently been proposed to play a role in ER stress-induced cell death⁷.

Bax inhibitor-1 (BI-1) performs a protective role against ER stress-induced cell death^8,9,10. Embryonic fibroblasts from BI-1^-/- mice are hypersensitive to ER stress-induced apoptosis, a finding that has been attributed to increased release of Ca²⁺ from the internal stores⁸. Recently, BI-1-mediated protection against ER stress was proposed to be involved in Ca²⁺ regulation¹¹, implying that BI-1 may possess a pH-sensitive motif for sensing cellular pH. The low levels of [Ca²⁺]_ER observed upon overexpression of BI-1 are related to a low mitochondrial Ca²⁺ concentration ([Ca²⁺]_mito) in BI-1-overexpressing cells¹². Considering that the mechanism of ER stress-induced cell death involves both the ER and mitochondria¹³, low [Ca²⁺]_ER in HT1080/BI-1 may affect [Ca²⁺]_mito, which likely plays a role in cell protection. However, [Ca²⁺]_mito can be also affected by mitochondrial physiological functions¹⁴. For example, the mitochondrial membrane potential (Δψm) is both directly and indirectly related to mitochondrial Ca²⁺ channel-like proteins, such as the Ca²⁺ uniporter¹⁵ and the Ca²⁺-dependent mitochondrial K⁺ channel¹⁶.

The role of BI-1 has been also studied in the context of mitochondrial physiology. A recent study using a yeast system to investigate the effects of Arabidopsis BI-1 (AtBI-1) concluded that mitochondrial electron transport chain proteins are required for BI-1-mediated protection against Bax¹⁷. Overexpression of BI-1 has also been shown to alter mitochondrial function through a mechanism proposed to involve reduced mitochondrial glucose metabolism and O₂ consumption¹⁸. Although the extent to which BI-1 affects various parameters of mitochondrial function, such as Δψm, has not been fully clarified, we hypothesize that BI-1 affects [Ca²⁺]_mito, thereby altering mitochondrial function. In this study, we focused on elucidating the mechanism by which BI-1 reduces [Ca²⁺]_mito by examining the opening of the mitochondrial permeability transition pore and the subsequent release of cytochrome c. We also investigated the relationship between [Ca²⁺]_mito and the regulation of cell death by BI-1.

Results

BI-1 is localized to mitochondria-associated membranes and the ER

To gain insight into the role of BI-1 in mitochondrial function, we first determined its subcellular localization. Initially, we performed biochemical fractionation of HT1080 fibrosarcoma cells stably transfected with either a construct conferring neomycin resistance (HT1080/Neo), or a construct driving the overexpression of BI-1 (HT1080/BI-1). Western blot analysis of the fractions revealed that some of the BI-1 was fractionated with the mitochondria-associated membrane (MAM), although the majority was fractionated with the ER (Fig. 1A). Expression of BI-1 was also observed in crude mitochondria fractions but not in pure mitochondria fractions, showing the absence of BI-1 in the mitochondria organelle.

We also confirmed the integrity of each fraction by immunoblotting for subcellular organelle marker proteins: calnexin for ER and MAM, Tom20 for mitochondria and voltage-dependent anion channel (VDAC) which enrich in pure mitochondria and also present in MAM^3,19,20,21 (Fig. 1A). In addition, we performed ultrastructural studies to analyze the localization of BI-1. Electron microscopy (EM) images of HA-immunogold staining revealed that the majority of BI-1 was ER-localized and a detectable proportion was also present on the MAM (Fig. 1B). Similarly, in a fluorescence study, BI-1 expression was co-localized with calnexin and VDAC (Fig. 1C). Figure 1D shows quantitative analysis of the overlap between BI-1 and calnexin (about 55%) or between BI-1 and VDAC (about 20%). Consistent with previous findings, these data indicated that BI-1 protein was clearly observed in MAM, as well as the ER.

BI-1 regulates ER stress-induced mitochondrial Ca²⁺ accumulation

The protective role of BI-1 against ER stress-induced cell death has been well characterized^8,10,22. Because the accumulation of mitochondrial Ca²⁺ plays a role in ER stress-induced cell death²³, we investigated the effect of BI-1 on [Ca²⁺]_mito. We measured thapsigargin-stimulated [Ca²⁺]_mito using the fluorescent dye Rhod II and observed that HT1080/BI-1 exhibited lower levels of fluorescence than parental HT1080 cells or control, HT1080/Neo (Fig. 2A). Next, we analyzed thapsigargin-induced [Ca²⁺]_ER changes in the absence of extracellular Ca²⁺ using an ER-targeted version of aequorin (erAEQ). Even in the absence of ER stress, HT1080/BI-1 showed lower levels of erAEQ luminescence than parental HT1080 or HT1080/Neo cells (Fig. 2A, lower), indicating that HT1080/BI-1 cells have lower [Ca²⁺]_ER. We next exposed cells to classic ER stress-inducing agents, such as tunicamycin, an inhibitor of N-acetylglycosylation and brefeldin A, a protein transport inhibitor (Fig. 2B). Both agents markedly increased [Ca²⁺]_mito; however, HT1080/BI-1 exhibited lower [Ca²⁺]_mito than HT1080/Neo. The basal [Ca²⁺]_mito was also consistently lower in HT1080/BI-1 than in HT1080/Neo. To further clarify the protective role of BI-1 against mitochondrial Ca²⁺, we developed cell lines with different levels of BI-1 expression. Thapsigargin-induced mitochondrial Ca²⁺ was regulated in a BI-1 expression-dependent manner (Fig. 2C). Furthermore, the decrease in [Ca²⁺]_ER and the increase in [Ca²⁺]_mito were consistently of a higher magnitude in BI-1 knockout mouse embryonic fibroblasts (BI-1^-/- MEF cells) compared with BI-1 wild-type cells (BI-1^+/+) (Fig. 2D).

Mitochondrial Ca²⁺ intake is reduced in HT1080/BI-1 cells

To confirm the low mitochondrial Ca²⁺ in the HT1080/BI-1, we indirectly measured [Ca²⁺]_mito in Fura-2AM-loaded HT1080/Neo and HT1080/BI-1. As expected, the increase in intracellular Ca²⁺ ([Ca²⁺]_i) was significantly lower in HT1080/BI-1, compared with HT1080/Neo, upon treatment with a mitochondrial uncoupler, carbonylcyanide m-chlorophenylhydrazone (CCCP) (Fig. 3A). This suggests that the [Ca²⁺]_mito was lower in the BI-1 overexpressing system compared with the control. Analysis of Rhod II fluorescence revealed that HT1080/BI-1 accumulated less mitochondrial Ca²⁺ than HT1080/Neo (Fig. 3B), consistent with the results shown in Figure 3A. We next examined ⁴⁵Ca²⁺ intake by mitochondrial membrane fractions and found that ⁴⁵Ca²⁺ intake by mitochondria from HT1080/BI-1 was significantly lower than that by mitochondria from HT1080/Neo (Fig. 3C). Direct exposure of mitochondria to Ca²⁺ also revealed lower levels of mitochondrial Ca²⁺ intake in HT1080/BI-1 compared with HT1080/Neo (Fig. 3D).

The Ca²⁺ uniporter is involved in mitochondrial Ca²⁺ intake in HT1080/BI-1 cells

To further investigate the mechanism underlying the reduced [Ca²⁺]_mito intake in HT1080/BI-1, we examined whether the Ca²⁺ uniporter is involved. This channel regulates both [Ca²⁺]_mito intake and ER stress-induced cell death²³. Using the Rhod II fluorescence assay, we observed that RU360-mediated inhibition of the Ca²⁺ uniporter decreased [Ca²⁺]_mito intake in HT1080/Neo, but had a smaller effect in HT1080/BI-1 (Fig. 4A). The values of [Ca²⁺]_ER and [Ca²⁺]_mito with or without RU360 treatment were shown in Fig. 4B. In our erAEQ-transfected cell system, we did not observe any effects of RU360 on [Ca²⁺]_ER; therefore, we conclude that RU360 specifically affects [Ca²⁺]_mito. Using the Ca²⁺ intake assay, we observed that inhibition of the Ca²⁺ uniporter decreased [Ca²⁺]_mito intake in HT1080/Neo, but had no effect on [Ca²⁺]_mito intake in HT1080/BI-1 (Fig. 4C). We also tested the effects of ER stress in this system and observed that RU360 treatment regulated thapsigargin-induced cell death without affecting the protective effect of overexpression of BI-1 (Fig. 4D). These data suggest that the mitochondrial Ca²⁺ uniporter contributes to mitochondrial Ca²⁺ accumulation upon exposure to ER stress and that BI-1 regulates the Ca²⁺ uniporter, leading to modulation of mitochondrial Ca²⁺ intake.

BI-1 regulates opening of the mitochondrial membrane permeability transition pore

As a messenger of cell death, mitochondrial Ca²⁺ opens the mitochondrial membrane permeability transition pore (PTP), thereby releasing cytochrome c into the cytoplasm²⁴. PTP opening has been shown to be involved in ER stress-associated cell death^25,26. We therefore examined whether the regulation of PTP is related to the cell protective effect in BI-1–overexpressing cells through trypan blue exclusion analysis. In HT1080/Neo, thapsigargin reduced cell viability to 27%; however, the PTP inhibitor cyclosporine A (CsA) increased cell viability. As expected, HT1080/BI-1 cells were resistant to thapsigargin. The protective effect of BI-1 in HT1080/BI-1 cells was greater than that of CsA in thapsigargin-treated HT1080/Neo cells, implying another protective mechanism in addition to the PTP regulation seen in HT1080/BI-1. The presence of CsA had little effect on the viability of thapsigargin-treated HT1080/BI-1 (Fig. 5A), suggesting that PTP regulation is at least one mechanism by which BI-1 protects cells against thapsigargin. Annexin V/propidium iodide analysis, another assay for cell death, showed similar protection against thapsigargin in HT1080/BI-1 (Fig. 5B). CsA also protected HT1080/Neo against the ER-stress agents, tunicamycin and brefeldin A (Fig. 5C). Consistent with these findings, HT1080/BI-1 cells were protected against the ER-stress agents and were minimally affected by CsA. To confirm the mechanistic link between Ca²⁺ and the resultant PTP opening, [Ca²⁺]_ER and [Ca²⁺]_mito were analyzed in the presence of ER stress with or without the PTP inhibitor, CsA. CsA treatment did not affect either [Ca²⁺]_ER or [Ca²⁺]_mito in ER stress-exposed HT1080/Neo and HT1080/BI-1 (Fig. 5D), suggesting that PTP opening is not a cause of the Ca²⁺ disturbance, but a consequence of [Ca²⁺]_mito accumulation. For direct PTP measurement, mitochondria from HT1080/Neo and HT1080/BI-1 were exposed to the same amount of Ca²⁺ and mitochondrial swelling was measured. Immunoblotting with anti-Hsp60, anti-caspase-3, or anti-calreticulin antibody confirmed the appropriate mitochondrial fractionation of HT1080/Neo and HT1080/BI-1 (Fig. 6A, upper). To eliminate interference by the small amount of Ca²⁺ released spontaneously from the mitochondria, we routinely included 25 μM EGTA. Under these conditions, mitochondria from HT1080/Neo and HT1080/BI-1 were exogenously exposed to 100 μM Ca²⁺. Swelling of mitochondria, which is considered a marker for PTP opening^27,28, was clearly observed in the Ca²⁺-exposed mitochondria from HT1080/Neo by monitoring optical density at 540 nm (Fig. 6A). Mitochondrial PTP opening was significantly suppressed by CsA. PTP opening/mitochondria swelling was also observed in Ca²⁺-exposed mitochondria from HT1080/BI-1, but to a lesser extent than in HT1080/Neo. In the BI-1-overexpression system, treatment with CsA had no additional regulatory effect on PTP opening, suggesting that the presence of BI-1 itself induces the regulation of PTP opening. Accumulation of Ca²⁺ in the mitochondrial matrix induces mitochondrial PTP opening and loss of mitochondrial membrane potential, Δψm²⁶. This study also showed a clear loss of Δψm after Ca²⁺ exposure in HT1080/Neo compared with HT1080/BI-1 (Fig. 6B). The consequential release of cytochrome c was confirmed with purified mitochondria from HT1080/Neo and HT1080/BI-1. Exposure of mitochondria to Ca²⁺ increased the amount of extra-mitochondrial/supernatant cytochrome in the HT1080/Neo group, but this was suppressed by inhibition of PTP opening by CsA (Fig. 6C). However, the release of cytochorome c was not observed in Ca²⁺-exposed mitochondria from HT1080/BI-1. We next investigated the effects of Ca²⁺ on the induction of cytochrome c release via PTP opening in BI-1^+/+ and BI-1^-/- MEF cells. Immunoblotting with antibodies against Hsp60, caspase-3, or calreticulin confirmed appropriate mitochondrial fractionation from these MEF cells (Fig. 6D, upper). Ca²⁺-induced PTP opening was greater in BI-1^-/- MEF cells; however, CsA was able to effectively block PTP opening in both cell types (Fig. 6D). Although the initial Δψm was similar between BI-1^+/+ and BI-1^-/- cells, the BI-1^-/- cells were more susceptible to Ca²⁺-induced disruption of Δψm than BI-1^+/+ cells (Fig. 6E). Consistent with these findings, cytochrome c release into the extra-mitochondrial environment was higher in BI-1^-/- cells than in BI-1^+/+ cells (Fig. 6F). Mitochondria from BI-1^-/- livers also showed higher sensitivity to Ca²⁺-induced PTP opening and cytochrome c release than mitochondria from BI-1^+/+ livers (Fig. S1A, B).

The mitochondrial Ca²⁺-dependent K⁺ channel (mitoK_Ca) is activated in HT1080/BI-1

Under continuous leakage of Ca²⁺ from the ER, other ions such as K⁺ enter the mitochondria instead of Ca²⁺ to maintain mitochondrial homeostasis^29,30. K⁺-related channel opening is also involved in the maintenance of mitochondrial membrane potential^31,32. To understand the role of channel proteins in the Δψm, Ca²⁺-exposed mitochondria from HT1080/Neo and HT1080/BI-1 cells were co-treated with NS1619 (a cell-permeable mitoK_Ca channel opener³²), paxilline (a mitoK_Ca channel inhibitor³³), diazoxide (a mitoK_ATP channel opener), or 5-hydroxydecanoate (5-HD; a mitoK_ATP channel blocker³⁴) and the Δψm was measured. Consistent with the data in Fig. 6B, basal Δψm was lower in HT1080/BI-1 than in HT1080/Neo. Under Ca²⁺ exposure, mitochondria from HT1080/BI-1 showed resistance to the Ca²⁺, revealing a relatively stable Δψm compared with those from HT1080/Neo (Fig. 7A). Among the inhibitors, the mitoK_Ca channel inhibitor paxilline had the greatest effect on Δψm in Ca²⁺-exposed mitochondria from HT1080/BI-1. Compared with HT1080/BI-1, paxilline had less effect on the Δψm in Ca²⁺-exposed HT1080/Neo, suggesting that BI-1 is involved in opening of the mitochondrial Ca²⁺-dependent K⁺ channel and providing a possible mechanism for the reduced, but stably maintained, Δψm in HT1080/BI-1.

Entry of K⁺ through mitochondrial K⁺ channels leads to net oxidation of mitochondria. Unique fluorescence assays can measure this oxidation^35,36 and have been used to examine the mitochondrial entry of K⁺. Therefore, we performed a flavoprotein fluorescence assay to examine the effects of BI-1 on mitoK_Ca channel opening. Figure 7B shows a representative time course of the changes in flavoprotein fluorescence in HT1080/Neo and HT1080/BI-1 cells exposed to NS1619 and/or paxilline. In these experiments, NS1619 reversibly oxidized flavoprotein, with subsequent exposure to 2,4-dinitrophenol (DNP) leading to full flavoprotein oxidation. In the presence of paxilline, NS1619 failed to oxidize flavoprotein, indicating complete inhibition of NS1619-induced oxidation by paxilline. In general, flavoprotein oxidation was higher in HT1080/BI-1 than in HT1080/Neo. We also examined the modulation of mitoK_Ca channels in ER stress-exposed cells by examining the effects of thapsigargin on flavoprotein oxidation. Figure 7C shows a representative experiment investigating the effects of thapsigargin on NS1619-induced flavoprotein oxidation. Thapsigargin enhanced the oxidative effect of NS1619, when added after the NS1619 effect had reached a steady state. The thapsigargin-induced flavoprotein oxidation was significantly higher in HT1080/BI-1 than in HT1080/Neo. These results also indicate that mitoK_Ca channels are more highly activated in HT1080/BI-1 than in HT1080/Neo.

Analysis of mitochondrial Ca²⁺ revealed that NS1619 treatment markedly inhibited thapsigargin-increased mitochondrial Ca²⁺ concentrations in HT1080/Neo, but only slightly affected mitochondrial Ca²⁺ in HT1080/BI-1 (Fig. 8A). The area under the curve of mitochondrial Ca²⁺ fluorescence was quantified in Figure 8B and also showed the regulatory effect of NS1619 on mitochondrial Ca²⁺ in HT1080/Neo, but not in HT1080/BI-1. These data suggest that BI-1 is probably involved in the open state of the Ca²⁺-dependent K⁺ channel, leading to the reduced mitochondrial Ca²⁺. Next, cell viability assays were performed with K⁺-modulating agents. No significant effect on the viability of HT1080/Neo or HT1080/BI-1 was seen with either the mitoK_ATP channel activator diazoxide or the mitoK_ATP channel inhibitor 5-HD. However, the mitoK_Ca activator NS1619 significantly protected against thapsigargin or tunicamycin-induced cell death, especially in HT1080/Neo (Fig. 8C, D). Furthermore, the mitoK_Ca inhibitor, paxilline, inhibited the BI-1–mediated protection from ER stress, implying that the mitoK_Ca channel is activated and functional in HT1080/BI-1. The BI-1–induced regulation of mitochondrial Ca²⁺ and mitoK_Ca channel opening was also confirmed in other cells, including BI-1–overexpressing A549 lung carcinoma cells (A549/BI-1) and tetracycline-inducible BI-1–expressing Hela cells (Fig. S2, S3). Taken together, these data indicate that BI-1 activates mitoK_Ca channels and that this activation is related to low mitochondrial Ca²⁺ and protection against ER stress.

Discussion

Results of this study show that BI-1 protects against ER stress-induced cell death by reducing [Ca²⁺]_mito intake, leading to the regulation of mitochondrial PTP opening and cytochrome c release. This study also suggests that BI-1–associated mitochondrial Ca²⁺ regulation is related to alteration of mitochondrial Ca²⁺ intake capacitance as well as the resistance against ER stress-induced cell death. We propose two mechanisms to explain the reduced amount of mitochondrial Ca²⁺ observed in HT1080/BI-1 cells: (1) ER-localized and MAM-localized BI-1 induces Ca²⁺ leakage from the ER, thereby generating low basal levels of intra-ER Ca²⁺. Low concentrations of intra-ER Ca²⁺ consequently lead to low concentrations of mitochondrial Ca²⁺ in HT1080/BI-1; (2) Mitochondrial Ca²⁺ intake is altered in HT1080/BI-1. BI-1-mediated regulation of [Ca²⁺]_mito is related to the mitochondrial membrane Ca²⁺ uniporter and mitoK_Ca channel opening.

The main theme of this study is why and how mitochondrial Ca²⁺ is maintained at a relatively low level in HT1080/BI-1 cells, compared with HT1080/Neo. When exposed to ER stress, reduced Ca²⁺ transfer from the ER to mitochondria in HT1080/BI-1 could explain the reduced [Ca²⁺]_mito compared with HT1080/Neo (Fig. 2A, B). Even in the absence of stress, HT1080/BI-1 exhibited lower basal [Ca²⁺]_mito, probably through ER- or MAM-localized BI-1 Ca²⁺ channel-like activity. The lower [Ca²⁺]_mito in HT1080/BI-1 seems to be related to Ca²⁺ transfer between the ER, with low levels of Ca²⁺ and the physically juxtaposed mitochondria. However, mitochondria that are not associated with the ER have different mitochondrial Ca²⁺ intake capacitance in HT1080/BI-1 from those in HT1080/Neo (Fig. 3C, 3D). In addition to Ca²⁺ transfer from the ER to physically close mitochondria, the Ca²⁺ intake characteristics of mitochondria may also explain the reduced mitochondrial Ca²⁺ in HT1080/BI-1. To understand this issue, we studied differences in physiological mitochondria functions including mitochondrial membrane potential (Δψm). The basal Δψm, a mitochondrial characteristic used to reflect the Ca²⁺ intake capacity^5,30, was lower in HT1080/BI-1 than in HT1080/Neo (Fig. S4). This observation provides insight into the mechanism of reduced mitochondrial Ca²⁺ intake in HT1080/BI-1 since mitochondrial Ca²⁺ intake depends on Δψm¹⁵. The opening of the Ca²⁺ uniporter is also dependent upon the Δψm¹⁵. In HT1080/BI-1, the reduced Δψm may be one of the mechanisms responsible for reduced Ca²⁺ intake, likely due to reduced Ca²⁺ uniporter activity (Fig. 4A, B).

Based on the reduced mitochondrial Ca²⁺ intake capacitance, we hypothesized that opening of the mitochondrial permeability transition pore associated with mitochondrial Ca²⁺ overload might be attenuated in HT1080/BI-1. For clarification of mitochondrial functional differences between HT1080/Neo and HT1080/BI-1, mitochondria were prepared from both cell lines and exposed to Ca²⁺. The Ca²⁺-induced permeability transition pore opening and related cytochrome c release were delayed or inhibited in the presence of BI-1 (Fig. 6A, C, D, F). Even in the absence of ER membrane-localized BI-1, mitochondria membranes from HT1080/BI-1 showed resistance against the loaded Ca²⁺, suggesting that mitochondrial characteristics might be changed as a result of MAM-localized BI-1. Recently, Sano et al reported a role for BI-1 as a regulator of Ca²⁺ transfer from the ER to mitochondria, thereby changing mitochondrial bioenergetics³⁷. We have also reported that BI-1 modulates ATP levels, lactate production and mitochondrial O₂ consumption¹⁸. It has been suggested that the continuous Ca²⁺ leak contributes to the opening of Ca²⁺-dependent K⁺ channels³². Due to the K⁺ channel opening, aquaporin channel gets to be opened, thereby the intake of water is increased, resulting in swelling of mitochondria^14,38. In the HT1080/BI-1, the mitochondrial swelling has been also reported¹⁸. The continuous Ca²⁺ leak might also affect mitochondrial physiological characteristics including electron transfer activity, mitochondrial membrane lipid composition, or mitochondrial membrane potential^39,40,41,42. In HT1080/BI-1, numerous mitochondrial enzymes such as pyruvate dehydrogenase, α-ketoglutarate dehydrogenase and enzymes for oxidative phosphorylation may be affected by the reduced mitochondrial Ca²⁺⁴³. Similarly, lipid synthesis/metabolism enzymes on the MAM might be affected⁴⁴. In addition, MAM-localized lipid handling has a profound impact on mitochondrial metabolism such as respiration and membrane dynamics^45,46. These changes in mitochondrial function may affect Ca²⁺ homeostasis in mitochondria, leading to the resistance against Ca²⁺.

Mitochondrial membrane potential, another factor that is affected by the leakage of Ca²⁺, was resistant to Ca²⁺ and stably maintained in HT1080/BI-1, compared with HT1080/Neo (Fig. 6A, B). To understand how Δψm is regulated in the BI-1 system, mitochondrial K⁺ channel inhibitors were applied to the Ca²⁺-induced Δψm collapse model. The mitoK_Ca channel inhibitor paxilline induced the greatest collapse of the stably maintained membrane potential against Ca²⁺ in the BI-1 system (Fig. 7A), indicating that mitoK_Ca channel activity contributes to the BI-1-associated stably maintained Δψm. Similarly, Δψm was previously reported to be regulated by Ca²⁺-dependent mitochondrial K⁺ channel opening/closing^47,48. Instead of Ca²⁺ continuously entering the mitochondria, cationic ions such as K⁺ tend to enter, affecting Δψm^32,48. It has been suggested that the highly activated K⁺ channel is a protective mechanism against ischemia/hypoxic stress-induced intracellular Ca²⁺ overloading^49,50,51. Activation of other K⁺ channels also contributes to the stably maintained and resistant Δψm against cell-death stimuli^52,53.

We propose that regulation of mitochondrial Ca²⁺ could explain the protective role of BI-1 against ER stress-induced cell death. Additionally, reactive oxygen species (ROS) are affected by BI-1 and may also contribute to the BI-1 protective mechanism. Both ER stress-mediated production of ROS and subsequent cell death are regulated in BI-1-overexpressing cells¹⁰. We previously reported that basal ROS levels were significantly lower in HT1080/BI-1 than in HT1080/Neo¹⁰. Mitochondria are well established as a primary source of ROS^54,55; activation of the mitochondrial electron transport chain (ETC) leads to electron uncoupling, thereby producing toxic ROS⁵⁶. Since ETC protein complexes are involved in pumping protons (H⁺) into the intermembrane space, thereby generating Δψm, the reduced Δψm observed in HT1080/BI-1 may be linked to reduced ETC activity and the resultant reduction in mitochondrial ROS. We recently observed that mitochondrial ETC activity, in addition to other mitochondrial functions, was stably reduced in HT1080/BI-1, compared with control cells (data not shown). To fully understand the relationship between ROS and BI-1, further experiments are required.

In previous studies on BI-1, Ca²⁺ regulation has been consistently suggested as a protective mechanism against ER stress^8,57,58 The intra-ER leaky Ca²⁺ channel effect seems to be linked to the change in mitochondrial membrane potential and its associated mitochondrial Ca²⁺ intake alteration, as well as the recently reported enhanced lysosomal functional effect⁵⁹.

In summary, reduced mitochondrial Ca²⁺ intake is linked to the control of mitochondrial PTP opening and cell death. Our data support a model in which the mitochondrial Ca²⁺ uniporter and the mitoK_Ca channel are involved in maintaining low [Ca²⁺]_mito in BI-1-overexpressing cells as part of the mechanism by which BI-1 protects cells from ER stress.

Methods

Materials and cell culture

Unless otherwise noted, reagents, including thapsigargin, tunicamycin, brefeldin A, KCN and CsA were obtained from Sigma Chemical Company (St. Louis, MO, USA). RU360 was obtained from Calbiochem (San Diego, CA, USA). Fluorescent probes, including Fura2-AM and Rhod II AM, were obtained from Molecular Probes (Eugene, OR, USA). The human HT1080 fibrosarcoma cells were cultured in Dulbecco's modified Eagle's Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 20 mM Hepes, 100 μg/ml streptomycin and 100 units/ml penicillin. HT1080 cells were stably transfected with the pcDNA3 (Neo), or pcDNA3-BI-1-HA plasmids using the SuperFect transfection reagent (Qiagen, Valencia, CA). The transfected cells were cultured for 3 weeks in 1 mg/ml G418 (Invitrogen, Carlsbad, CA). Cells were maintained in DMEM containing 10% fetal bovine serum, 1 mM L-glutamine and penicillin (Invitrogen). The stably transfected cells were named as HT1080/Neo or HT1080/BI-1. Mouse embryonic fibroblasts (MEFs) obtained from BI-1^+/+ and BI-1^-/- mice were cultured in DMEM supplemented as above.

Subcellular fractionation

Cells were resuspended in buffer A [250 mM sucrose, 20 mM HEPES (pH 7.5), 10 mM KCl, 1.5 mM MgCl₂, 1 mM EDTA, 1 mM EGTA and 1× protease inhibitor cocktail] on ice for 30 min prior to lysis with a Dounce homogenizer. Samples were centrifuged at 750 × g for 10 min at 4°C to remove unlysed cells and nuclei and the resultant supernatants were centrifuged at 10,000 × g for 20 min at 4°C. The pellets (heavy membrane fraction, HM) were used for mitochondrial Ca²⁺ intake experiments, mitochondrial PTP opening assays and immunoblotting analysis with anti-Hsp60, anti-caspase-3, or anti-cytochrome c antibodies. The supernatants were centrifuged at 100,000 × g for 1 hr at 4°C and subsequently loaded onto a three-layered sucrose gradient. Purified ER membranes were then isolated by centrifugation at 152,000 × g. Further purification of mitochondria and the MAM fraction was performed by centrifugation (9,500 × g, 30 min, 4°C) on a 30% Percoll gradient as reported previously¹⁹. Mitochondria, MAM and ER fractions were lysed in RIPA buffer (1% sodium deoxycholate, 0.1% SDS, 1% Triton X-100, 10 mM Tris (pH 8.0), 0.14 M NaCl) for immunoblotting with antibodies against calnexin, Tom20, VDAC, HA, or BI-1.

Immunogold electron microscopy

Immunogold experiments were carried out using a previously described method^60,61 with modifications. HT1080 cells were fixed by immersion in Karnovsky solution (2% glutaraldehyde and 2% formaldehyde in 0.05 M sodium cacodylate buffer, pH 7.2) for 4 hrs at 4°C. Cells were then dehydrated in a graded series of ethanol and embedded in Spurr's resin. Thin sections (approximately 80 nm thick) were cut for transmission electron microscopy with a NOVA ultramicrotome (LKB) and picked up on 100-mesh gold grids (Electron Microscopy Sciences). Sections on grids were blocked for 1 hr by floating on drops of PBS containing 3% BSA and 0.05% Triton X-100 (T-PBS), followed by washing on drops of T-PBS (five changes, 3 min each). Sections were then incubated for 4 hrs at 4°C with anti-rat HA (Roche, Mannheim, Germany), or anti-mouse HA antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA) diluted 1:100 in T-PBS containing 0.1% BSA and 0.05% Triton X-100. The grids were washed by floating on drops of T-PBS. Bound antibodies were visualized by incubating the sections with gold-conjugated secondary antibodies (10-nm particle size; Sigma-Aldrich) diluted 1:200 in PBS containing 0.1% BSA and 0.05% Triton X-100 for 1 hr. Finally, grids were washed on drops of water (five changes, 5 min each) before being stained with 2% aqueous uranyl acetate for 30 sec. All incubations, except for the primary antibody step, were carried out at room temperature and all buffers used were 0.22-mm filtrates. The specimens were observed with a transmission electron microscope (H7650, accelerating voltage 100 kV, Hitachi) at the Center for University-Wide Research Facilities (CBNU).

Immunofluorescence microscopy

Immunofluorescence microscopy images were prepared as previously described⁶². HT1080/BI-1 cells were grown on a coverglass bottom dish for 24 hrs. Cells were washed with PBS and fixed with methanol for 5 min. Cells were then incubated overnight with the first primary antibody (1:100 dilution; rat anti-HA monoclonal; Roche). The first secondary antibody was FITC-conjugated anti-rat (1:200; Invitrogen). After three washes using PBS, cells were incubated with a second primary antibody (1:100 dilution for anti-calnexin polyclonal or anti-VDAC polyclonal; Santa Cruz Biotechnology) for 1 hr. The second secondary antibody was TRITC-conjugated anti-rabbit (1:200; Invitrogen). Finally, the cells were mounted in VECTASHIELD mounting medium with DAPI. Signals were detected using the FITC, TRITC and DAPI filter with a Delta Vision Deconvolution Microcopy System (Applied Precision, Issaquah, WA, USA). Colocalization was visualized and Pearson's correlation coefficient of the entire cell was determined using soft image analysis software (Applied Precision).

Measurement of PTP opening and cytochrome c release

Large amplitude swelling was measured as described previously²⁴ by spectrophotometric measurement of absorbance at 540 nm. Isolated mitochondria (1 mg/ml) were incubated in suspension with various agents in buffer A [200 mM sucrose, 10 mM Tris-MOPS (pH 7.4), 5 mM succinate, 1 mM potassium phosphate, 2 μM rotenone, 1 μg/ml oligomycin and 25 μM EGTA] at 30°C for 100 min. To monitor mitochondrial swelling, absorbance at A₅₄₀ was measured at designated time points throughout the incubation using a spectrophotometer (Amersham Pharmacia Biotech, Ultrospec Plus). To measure cytochrome c release, 50 μg of mitochondria was incubated at 30°C in 30 μl of Buffer A for 1 hr with various reagents. Supernatants were separated from mitochondria by centrifugation at 10,000 × g for 10 min at 4°C. Supernatants and pellets were resolved by SDS-PAGE on 8-14% gels and immunoblotted with anti-cytochrome c antibodies.

Immunoblotting

Equivalent protein amounts were resolved by SDS-PAGE on 12% acrylamide-0.8% bisacrylamide minigels, electroblotted onto PVDF membranes (Bio-Rad, CA, USA) and sequentially immunoblotted with antibodies against cytochrome c (PharMingen, San Diego, CA, USA), Hsp60 (Stressgen, CA, USA), calreticulin (Santa Cruz Biotechnology), caspase-3 (PharMingen), BI-1 (Abcam, Cambridge, UK) or HA (Roche).

Time-sequential analysis of mitochondrial and ER Ca²⁺ levels

Analysis of mitochondrial Ca²⁺ levels was performed as previously described^11,24. Briefly, cells were loaded with 1 μM Rhod II AM (Molecular Probes) and 0.02% Pluronic F-127 (Molecular Probes) in Krebs-Ringer bicarbonate buffer [25 mM HEPES (pH 7.2), 111 mM NaCl, 25 mM NaHCO₃, 5 mM KCl, 2 mM CaCl₂ and 1 mM MgCl₂] containing 5 mM glucose for 30 min. Cells were washed once with recording medium containing 129 mM NaCl, 4 mM KCl, 1 mM MgCl₂, 2 mM CaCl₂, 4.2 mM glucose and 10 mM HEPES (pH 7.4), followed by incubation at room temperature for 60 min in recording medium supplemented with 0.02% Pluronic F-127 and 3 μM Rhod II AM. Culture medium was changed to recording medium and the cells were exposed to either thapsigargin or tunicamycin. The fluorescence intensity was determined every minute. Images were obtained using a confocal laser-scanning microscope equipped with a helium-neon laser (LSM510, Carl Zeiss, Jena, Germany), using an objective lens with numeral apertures of 0.5 for 20-fold magnification. Fluorescence images labeled with Rhod II were collected using an excitation wavelength of 543 nm. Rhod II fluorescence was detected using single excitation and emission filters and plotted as percentage change from the initial value. Parameters of illumination and detection were controlled digitally for consistent settings throughout the experiments. Two successive digital images were collected, usually at 5126512 pixels in the same visual field. To confirm the dye localization, cells were exposed to Rhod II AM for 1 hr, followed by staining with 100 nM MitoTracker Green FM for 15 min at 37°C. Cells were washed three times and observed under a fluorescence microscope to visualize mitochondria. Drugs were prepared in recording medium immediately before each use.

In separate experiments, measurements of [Ca²⁺]_ER were performed in HT1080/Neo and HT1080/BI-1. Briefly, 24 hrs after transfection with ER-localized aequorin, cells were incubated in saline buffer [135 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 1 mM MgSO₄, 0.4 mM KH₂PO₄, 5.5 mM glucose and 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, pH 7.4] supplemented with 5 mM n-coelenterazine, 4 mM ionomycin and 600 mM ethyleneglycol tetra-acetic acid (EGTA) at 37°C for 40 min. Cells were transferred to a perfusion chamber within a purpose-built luminometer, washed extensively with saline buffer containing 100 mM EGTA and 2% bovine serum albumin and exposed to thapsigargin. The fluorescence intensity was determined every minute. Light emission was measured in a luminescence counter (MicroBeta, LumiJET Microplate Counter PerkinElmer) at 469 nm every 0.1 sec. Finally, to release the remaining aequorin pool and to calibrate the aequorin signal, the cells were lysed with 100 mM digitonin in a hypotonic Ca²⁺-rich solution (10 mM CaCl₂ in H₂O). Aequorin luminescence signals were collected and calibrated to [Ca²⁺] values as described previously⁶³.

Mitochondrial Ca²⁺ analysis using the mitochondrial uncoupler CCCP

To compare mitochondrial Ca²⁺ between HT1080/Neo and HT1080/BI-1, cells were plated onto glass-bottomed perfusion chambers. The chambers were mounted on the stage of an inverted microscope (Nikon Eclipse TE2000) and incubated with Fura-2 AM (2 μM, 1-[2-(5-carboxyoxazol-2-yl)-6-aminobenzoFURAn-5-oxy]-2-(2-amino-5-methylphenoxy)-ethane-N,N,N′N′-tetraacetic acid pentaacetoxymethyl ester, Molecular Probes) for 30 min at room temperature in Hanks' balanced salt solution (HBSS). After loading, cells were washed three times in isotonic buffer without Ca²⁺ (KH buffer: 132 mM NaCl, 5 mM KCl, 10 mM dextrose, 10 mM HEPES, 1.05 mM MgCl₂). Cells were then promptly treated with 1 μM CCCP (Carbonyl cyanide m-chlorophenyl hydrazone), a mitochondrial uncoupler, to compare the amount of mitochondrial Ca²⁺ between HT1080/Neo and HT1080/BI-1⁶⁴. As an indirect marker for [Ca²⁺]_mt, changes in [Ca²⁺]_i as a result of treatment with CCCP were determined as a ratio of 340 nm/380 nm excitation for 512 nm emission using an integrated spectrofluorometer (Photon Technology International, Birmingham, NJ). Ca²⁺ concentrations were calculated using the equation [Ca²⁺]_i = K_d (F₃₈₀max/F₃₈₀min) (R − R_min)/(R_max − R); a K_d value of 229 nM was assumed for the binding of calcium to Fura-2AM. R_max and R_min were determined in each experimental group by the consecutive addition of 30 μM digitonin (R_max) and 50 mM EGTA (R_min). Using CCCP, mitochondrial Ca²⁺ was also measured in a Rhod II AM dye-loaded condition, similar to the time-sequential analysis of mitochondrial Ca²⁺ described above.

Mitochondrial Ca²⁺ intake analysis

Mitochondrial Ca²⁺ intake was measured using ⁴⁵Ca²⁺. Mitochondria vesicles were suspended (1 mg/mL final concentration) in a buffer solution containing 100 mM KCl and 20 mM MOPS-Tris (pH 7.2). Free Ca²⁺ concentration in the incubation buffer was fixed at the desired values using different amounts of CaCl₂ (containing 1 μCi ⁴⁵Ca²⁺ mmol⁻¹ CaCl₂, Amersham, Buckinghamshire, UK) and EGTA. Vesicles were incubated in the buffer at 25°C for 1 min and transport was initiated at time zero. Ca²⁺ intake buffer (129 mM NaCl, 4 mM MgCl₂, 0.1 mM CaCl₂, 4.2 mM glucose, 2 mM succinate and 10 mM HEPES, pH 7.4) was mixed with 1 μCi ⁴⁵Ca²⁺ and 2 mM ATP at 37°C. The reaction was started by addition of 100 μL of protein (250 μg, total volume 500 μL). At defined time intervals 100-μL aliquots were removed, filtered on pre-wetted nitrocellulose filters (0.45-μm pore size, type WCN, Whatman, Clifton, NJ) and rapidly washed twice with 5 mL of ice-cold wash medium (140 mM KCl, 10 mM NaCl, 2.5 mM MgCl₂, 10 mM Hepes-KOH, pH 7.0,). Filtration and rinsing were completed within 30 sec. Samples of 250 μL were placed in a scintillation vial, dissolved in 1 ml scintillation grade 2-methoxyethanol and counted in a liquid scintillation analyzer (Packard Instruments Co., Downers Grove, IL). All experiments were performed using triplicates and each experiment was performed at least twice. Separately, mitochondrial Ca²⁺ intake from the medium was measured with Rhod II AM. Briefly, isolated mitochondria (0.5 mg protein) were suspended in 2 mL high KCl buffer (110 mM KCL, 0.5 mM KH₂PO₄, 1 mM MgCl₂, 20 mM HEPES, 0.01 mM EGTA, 5 mM succinate at pH 7.2) and fluorescence was recorded at room temperature at 550 nm excitation and 580 emission (Photon Technology International, Birmingham, NJ). F_min and F_max were established by addition of 1 mM EGTA and 10 mM Ca²⁺.

JC-1 assay for mitochondrial membrane potential

This assay is based on the changes in electrochemical gradient across the mitochondrial membrane measured by membrane potential (Δψm)⁶⁵. Loss of Δψm is detected by a fluorescent cationic dye JC-1 (5,59,6,69-tetra chloro-1,19,3,39-tetraethylbenzamidazolo-carbocyanin iodide (Cell Technology, Mountain View, CA). Briefly, freshly isolated mitochondria were labeled with JC-1 for 15 min at 37°C in a 5% CO₂ incubator. After two washes with assay buffer, JC-1 labeled and unlabeled mitochondria suspensions were transferred to 96-well black-walled plates. Fluorescence intensity was read at excitation 550 nm, emission 600 nm for red fluorescence and at excitation 485 nm, emission 535 nm for green fluorescence using a SpectraMax fluorescence plate reader (Molecular Devices, Sunnyvale, CA). The ratio of red to green determined the mitochondria membrane potential. A decrease in the ratio indicated collapse of Δψm.

Assessment of cell death

Cell-surface exposure of phosphatidylserine was assessed by Annexin V/PI fluorescence (MACS, Miltenyi Biotech) according to the manufacturer's protocol. Briefly, 2 × 10⁵ cells were collected, washed in PBS, pelleted and resuspended in incubation buffer [10 mM HEPES/NaOH (pH 7.4), 140 mM NaCl and 5 mM CaCl₂] containing 1% Annexin V. Samples were incubated in the dark for 15 min prior to addition of another 400 μl of incubation buffer and subsequently analyzed with a fluorescence-activated cell sorter (Partec, Germany). To measure cell viability, a trypan blue exclusion assay was performed as described previously⁶⁶. Briefly, treated or non-treated cells were assessed microscopically for dead cells by trypan blue exclusion. Cell viability was calculated by dividing the non-stained (viable) cell count by the total cell count. The number of cells was determined by averaging the number of cells in four squares and multiplying this average by a dilution factor.

Flavoprotein fluorescence

Autofluorescence of mitochondrial flavoprotein was measured using a modified version of a previously reported method^11,48. Briefly, cells were superfused with glucose-free Tyrode's solution containing 140 mmol/L NaCl, 5.4 mmol/L KCl, 1.8 mmol/L CaCl₂, 0.33 mmol/L NaH₂PO₄, 0.5 mmol/L MgCl₂ and 5 mmol/L HEPES (pH 7.4) at room temperature (22°C). Flavoprotein fluorescence was measured with excitation at 480 nm and emission at 520 nm. Relative fluorescence was calibrated according to signals after application of the mitochondrial uncoupler 2,4-dinitrophenol (DNP) (100 μmol/L).

Statistical analysis

Results are presented as means ± S.E. of n cells. Paired and unpaired Student's t-tests were used to determine statistically significant differences between test and control conditions as appropriate. MicroCal Origin software (Northampton, MA, USA) was used for statistical calculations.

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Acknowledgements

This work was supported by a Korea Science and Engineering Foundation Grant (2012R1A2A1A03001907).

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Department of Pharmacology and Cardiovascular Research Institute, Medical School, Chonbuk National University, Jeonju, 561-181, Republic of Korea
Geum-Hwa Lee, Hwa-Young Lee, Bo Li & Han-Jung Chae
Department of Dental Pharmacology and Wonkwang Dental Research Institute, School of Dentistry, Wonkwang University, Iksan, 570-749, Republic of Korea
Hyung-Ryong Kim

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All authors contributed to this work. G.H.L. and H.J.C. conceived and designed the experiments; G.H.L. and B.L. performed the experiments; H.R.K. contributed reagents/materials/analysis tools; G.H.L., H.Y.L. and H.J.C. wrote the paper.

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Lee, GH., Lee, HY., Li, B. et al. Bax Inhibitor-1-Mediated Inhibition of Mitochondrial Ca²⁺ Intake Regulates Mitochondrial Permeability Transition Pore Opening and Cell Death. Sci Rep 4, 5194 (2014). https://doi.org/10.1038/srep05194

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Received: 05 November 2013
Accepted: 20 May 2014
Published: 05 June 2014
DOI: https://doi.org/10.1038/srep05194

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