During apoptosis, cytochrome c is released into the cytosol as the outer membrane of mitochondria becomes permeable, and this acts to trigger caspase activation. The consequences of this release for mitochondrial metabolism are unclear. Using single-cell analysis, we found that when caspase activity is inhibited, mitochondrial outer membrane permeabilization causes a rapid depolarization of mitochondrial transmembrane potential, which recovers to original levels over the next 30–60 min and is then maintained. After outer membrane permeabilization, mitochondria can use cytoplasmic cytochrome c to maintain mitochondrial transmembrane potential and ATP production. Furthermore, both cytochrome c release and apoptosis proceed normally in cells in which mitochondria have been uncoupled. These studies demonstrate that cytochrome c release does not affect the integrity of the mitochondrial inner membrane and that, in the absence of caspase activation, mitochondrial functions can be maintained after the release of cytochrome c.
Keywords: apoptosis, mitochondria, membrane potential, caspases, ATP
Apoptotic cell death is orchestrated by the activation of caspase proteases that cleave key substrates within the cell during the apoptotic process. A major pathway for caspase activation involves a permeabilization of the mitochondrial outer membrane, which releases several proteins, including cytochrome c [Liu et al. 1996; Kluck et al. 1997], Smac/Diablo [Du et al. 2000; Verhagen et al. 2000], and others [Kluck et al. 1999; Kohler et al. 1999]. Cytochrome c binds and activates an adapter, the apoptotic protease activating factor [Apaf]-1, to recruit and activate caspase-9 [Li et al. 1997; Srinivasula et al. 1998; Zou et al. 1999; Cain et al. 2000].
In mitochondria, cytochrome c plays an essential role in generation of mitochondrial transmembrane potential [ΔΨm]. This potential is essential for various functions including the production of ATP via oxidative phosphorylation. Outer membrane permeability resulting in cytochrome c release should therefore impact on mitochondrial function.
Proapoptotic and antiapoptotic Bcl-2 family proteins regulate the mitochondrial outer membrane permeabilization. However, the exact mechanism by which this event occurs is controversial [for reviews see Gross et al. 1999; Vander Heiden and Thompson 1999; Waterhouse and Green 1999], and the various models that have been suggested impact on mitochondrial function in different ways. Cytochrome c release may proceed through the generation of pores or channels in the outer membrane, composed all or in part of proapoptotic Bcl-2 family proteins [Shimizu et al. 2000]. Alternatively, it may occur through disruption of the outer membrane after swelling of the mitochondrial matrix due to opening of the permeability transition pore [Marzo et al. 1998a,Marzo et al. 1998b; Brenner et al. 2000] or as a consequence of a closure of the voltage-dependent anion channels in the mitochondrial outer membrane [Vander Heiden and Thompson 1999; Vander Heiden et al. 2000]. In each case, however, the behavior and function of mitochondria should be affected, because the proton gradient generated by the electron transport chain should be impaired. Disruption of the ΔΨm may even kill a cell if downstream apoptotic effects are blocked. Such Bcl-2–regulated, caspase-independent cell death has been described [McCarthy et al. 1997; Martinou et al. 1999; Deshmukh et al. 2000; Haraguchi et al. 2000].
Mitochondrial function may therefore impact on the death of a cell in several ways. Altered ΔΨm may lead to cytochrome c release and activation of caspases or conversely cytochrome c release may alter mitochondrial function, which in the absence of caspase activity may lead to the death of the cell. Here, we take advantage of single-cell analysis to follow changes in the mitochondrial transmembrane potential in relation to mitochondrial outer membrane permeabilization in cells triggered to undergo apoptosis after toxic insults. We observed that a reduction in ΔΨm followed within minutes after the release of cytochrome c. We found, however, that in the absence of caspase activity mitochondria use, cytochrome c at the concentration maintained within the cytoplasm to regenerate ΔΨm and maintain ATP generation.
HeLa cells stably expressing green fluorescent protein [GFP]–tagged cytochrome c [Cc-GFP-HeLa] [Goldstein et al. 2000], Cc-GFP-HeLa cells expressing Bcl-2 and murine embryonic fibroblasts deficient in Apaf-1 [a gift from Dr. F. Cecconi, University of Rome, Rome, Italy] were cultured in DME [GIBCO BRL], and Jurkat cells were grown in RPMI-1640 [GIBCO BRL]. All cell lines were maintained at 37°C in a humidified atmosphere of 95% air, 5% CO2, and all media were supplemented with 2 mM glutamine, 200 μg/ml penicillin, 100 μg/ml streptomycin sulphate, and 10% FBS. Adherent cells were subcultured 1:10 by incubating them in 0.25% trypsin [GIBCO BRL] when they were 70% confluent, and resuspending cells were subcultured in growth medium. Suspension cells were subcultured 1:10 when they reached 106 cells/ml.
To induce death, Cc-GFP-HeLa cells were treated with actinomycin D [1 μM], staurosporine [1 μM], or UVC ultraviolet C [180 mJ/cm2]. For UV treatment, cells were washed in PBS and irradiated with UV light in PBS at 37°C. The PBS was then aspirated, and the media were replaced. Apaf−/− cells were treated with actinomycin [1 μM], and Jurkat cells were treated with actinomycin D [500 nM], staurosporine [500 nM], or etoposide [40 μM]. In experiments where glucose-free medium was used, the cells were washed once with serum-free, glucose-free DME and cultured for 12–15 h in DME containing 10% FBS dialyzed against PBS.
Cells having undergone specific apoptotic events were detected by flow cytometry using a FACScan® [Becton Dickinson] with a 488-nm laser line and analyzed using Cell Quest software. Phosphatidylserine exposed on the outside of cells was determined by annexin V binding. In brief, cells were pelleted and resuspended in 100 μl of Annexin V-FITC [Calbiochem] diluted 1:100 in annexin buffer [10 mM Hepes, 100 mM NaCl, 10 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2]. Cells were incubated for 5 min at 37°C, and 200 μl annexin buffer containing propidium iodide [0.5 μg/ml] was added before FACS® analysis. Annexin V-FITC fluorescence was detected in FL-1, and propidium iodide was detected in FL-2.
Cytochrome c release was detected by incubating 4 × 104 Cc-GFP-HeLa cells in 100 μl of ice-cold cell lysis and mitochondria intact [CLAMI] buffer [250 mM sucrose, 70 mM, KCl 50 μg/ml digitonin in PBS] for 5 min. Aliquots from the suspensions were stained with 0.1% Trypan blue in PBS to ensure that >95% of the cells were lysed. 200 μl of ice-cold digitonin-free CLAMI buffer was added to the cell suspension, and the cells were measured immediately by flow cytometry. Cytochrome c–GFP fluorescence was detected in FL-1. At the concentration used here, digitonin selectively permeabilizes the plasma membrane, allowing any cytochrome c that has been released from the mitochondria to exit the cells. The fluorescence of cells with intact mitochondria was approximately one-third brighter than cells in which the mitochondria had released cytochrome c.
ΔΨm was measured primarily using tetramethylrhodamine ethyl ester [TMRE]. Cells were incubated at 37°C for 20 min in media containing TMRE [50 nM]. TMRE fluorescence was detected by flow cytometry using FL-2. ΔΨm was also measured using DiOC[6]3 [40 nM], and CMTM-Ros [150 nM] also diluted in media.
To determine the extent of cytochrome c release in cells by western blotting, 1.5 × 106 Jurkat cells were incubated on ice for 5 min in 100 μl of ice-cold CLAMI buffer containing 200 μg/ml digitonin. The lysis of >95% of the cells was confirmed by Trypan blue exclusion. The cells were pelleted [1,000 g for 5 min at 4°C], and the supernatant containing cytosolic protein was stored at −80°C. The pellets were incubated at 4°C for 10 min in universal immunoprecipitation buffer [50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 2 mM EDTA, 2 mM EGTA, 0.2% Triton X-100, 0.3% NP-40 1× complete™ protease inhibitor [Roche]. The samples were centrifuged [10,000 g for 10 min at 4°C], and the supernatant containing mitochondrial protein was stored at −80°C. Protein from each sample were boiled for 5 min in 5× sample loading buffer and electrophoresed in individual lanes of 15% SDS-PAGE gels. The proteins were transferred to supported Hybond C nitrocellulose [Amersham Pharmacia Biotech] and Western blotted using anti–cytochrome c [7H8.2C12; PharMingen] and antiactin [C4; ICN Biomedicals] diluted 1:1,000. The immobilized proteins were incubated with horseradish peroxidase secondary antibody, and the signal was detected using Dura Signal chemiluminescence reagent [Pierce Chemical Co.].
Confocal microscopy was performed using a Nikon Eclipse TE 300 microscope and a MRC 1024 confocal microscope [Bio-Rad Laboratories] using an Ar/Kr laser. For immunocytochemistry, Cc-GFP-HeLa cells were grown on LabTek four-well chamber slides [Nalge Nunc International]. Cells were fixed with 4% paraformaldehyde in PBS for 15 min. The cells were washed in blocking buffer [0.05% saponin, 3% BSA, in PBS] and incubated overnight at 4°C with anti-Bax antibody [PharMingen] diluted 1:200 in blocking buffer. The cells were washed in blocking buffer and incubated for 1 h at room temperature with rabbit Ig conjugated to Texas red [Amersham Pharmacia Biotech] diluted 1:200 in blocking buffer. The fluorescence of GFP and Texas red were detected by confocal microscopy using excitation wavelengths of 488 or 522 nm and detection wavelengths of 568 or 605 nm, respectively. Images were Kalman averaged.
For time-lapse analysis, Cc-GFP-HeLa cells were grown on glass-bottom microwell dishes [MatTek]. Cells were treated with apoptotic stimuli in phenol red–free DME, supplemented with 10% FBS, 20 mM Hepes, pH 7.2, 2 mM l-glutamine, 200 μg/ml penicillin, 100 μg/ml streptomycin sulphate, and TMRE [50 nM], and returned to an incubator for 2–12 h. The media were overlaid with mineral oil [Sav-On], and the dish was placed on the confocal microscope. The temperature was maintained at 37°C using an MS-C Temp Controller [Narishige]. Cells were excited using a 488-nm laser line attenuated at 96%. Cytochrome c–GFP and TMRE fluorescence were detected using 568 or 605 nm, respectively. Images were Kalman averaged three times each at 2-min intervals. Untreated cells followed under these conditions for 400 frames were undamaged, to the extent that mitosis was observed in many cells during this period.
Images were analyzed with Metamorph v4.0 [Universal Imaging Corp.] by drawing regions around individual cells and then computing standard deviation [punctate/diffuse] and integrated brightness [total brightness]. For ΔΨm, the total brightness of TMRE was divided by the total brightness of the cytochrome c–GFP to account for any movement of the cell, except in Apaf-deficient murine embryonic fibroblasts, which contained no cytochrome c–GFP. The punctate–diffuse index and the relative TMRE fluorescence index were calculated by dividing each value by the average of the first six values. In Fig. 3 C, the relative TMRE fluorescence of each cell was calculated by dividing each value by the average of the first 119 values. Quicktime movies were processed using NIH Image J software [National Institutes of Health].
Time course analysis of ΔΨm during apoptosis. Cc-GFP-HeLa cells stained with TMRE and treated with actinomycin D [1 μM] in the presence [+zVAD] or absence [−zVAD] of 100 μM zVADfmk as indicated, and were followed by time-lapse confocal microscopy. Images were taken every 2 min and the relative brightness of TMRE fluorescence and punctate–diffuse index of cytochrome c–GFP was calculated and plotted over time. [A] Two cells, which commenced cytochrome c–GFP release at 534 and 474 min show a subsequent drop in ΔΨm, commencing at 536 and 478 min, respectively. [B] Similar cells in the absence or presence of zVADfmk [100 μM] were followed by time-lapse confocal microscopy, and the relative brightness of TMRE fluorescence was calculated and plotted over time. For comparative purposes, cytochrome c–GFP release [indicated by arrow] was set at 4 h in each case. [C] The mean brightness of TMRE in individual cells aligned for cytochrome c release at 4 h. n represents the number of cells averaged. Error bars indicate SD.
ATP assays were performed using the ATP bioluminescence assay kit HSII [Roche] following the manufacturer's instructions. In brief, 105 cells were resuspended in 100 μl lysis buffer and stored at −80°C. Aliquots from each sample were diluted to 100 μl in dilution buffer. 100 μl of luciferase reagent was added to each sample and after a delay of 1 s, the luminescence was integrated over 10 s using a monolight 2010 luminescence recorder [Analytical Luminescence Laboratory]. The ATP concentration of the samples was determined by comparing values to a standard curve for ATP performed at the same time. The ATP concentration was standardized using the total cellular protein estimated by the micro BCA assay [Pierce Chemical Co.].
Cc-GFP-HeLa cells [2 × 106] were incubated for 20 min at 37°C in media containing TMRE [50 nM]. Buffers in all subsequent steps contained TMRE [50 nM]. The cells were trypsinized and incubated for 5 min on ice in 1 ml of ice-cold mitochondria isolation buffer [MIB] [200 mM mannitol, 50 mM sucrose, 10 mM Hepes, 10 mM succinate, 70 mM KCl, 1 mM DTT] containing 50 μg/ml digitonin. When >95% of cells were permeable to Trypan blue, the cells were washed twice in ice-cold MIB containing 0.1% BSA. For Fig. 2 A, the cells were incubated for 30 min at 37°C in 1 ml of MIB containing truncated Bid [tBid] [20 μg/ml] and diluted in 1:10 in MIB containing BSA [0.1%] ATP [1 mM], creatin-phosphate [5 mM], creatin kinase [0.1 mg/ml], oligomycin [10 μg/ml], and the concentrations of cytochrome c indicated. Carbonyl cyanide p-[trifluoromethoxy] phenylhydrazone [FCCP] [10 μM] and KCN [1 mM] were added as indicated. The cells were analyzed by flow cytometry after 20 min, measuring cytochrome c–GFP fluorescence in FL-1 and TMRE fluorescence in FL-2. Analysis confirmed that >95% of cells treated with tBid had released cytochrome c–GFP. For Fig. 2 B, the permeabilized apoptotic cells were directly resuspended in the MIB containing cytochrome c, and the cells were analyzed by flow cytometry after 20 min.
Cytochrome c concentration limits respiration in mitochondria that have undergone outer membrane permeabilization. [A] Permeabilized cells stained with TMRE [50 nM], either untreated or treated with tBid, were incubated at 37°C for 20 min with the concentrations of horse heart cytochrome c indicated. ΔΨm was measured by flow cytometry. FCCP [10 μM] was used as a control for depolarized mitochondria. KCN [1 mM] was used to block the involvement of cytochrome c in the electron transport chain. [B] Cc-GFP-HeLa cells were treated for 12 h with actinomycin D [1 μM] and stained with TMRE [50 nM]. The cells were permeabilized with digitonin and incubated for 20 min with horse heart cytochrome c. ΔΨm and cytochrome c–GFP were measured by flow cytometry. The cells were gated for cytochrome c–GFP release, and the relative fluorescence of TMRE was compared with that of cells that had not released cytochrome c–GFP. Error bars indicate SD.
Supplemental video of Fig. 4 shows loss and regeneration of ΔΨm after cytochrome c release. Cc-GFP-HeLa cells were treated with actinomycin D [1 μM] in the presence of N-benzoylcarbonyl-Val-Ala-Asp-fluoromethylketone [zVADfmk] [100 μM], and confocal images were taken every 2 min. The cytochrome c–GFP [green, left] shows the coordinate release of cytochrome c in the individual cells [the staining goes from punctate to diffuse upon release]. TMRE fluorescence in the same cells [red, right] shows the loss and recovery of ΔΨm. The red and green images are of the same cells taken at the same time. The frames are separate rather than overlaid for clarity, and a mathematical representation of loss and regeneration of ΔΨm in a similarly treated cell is shown in Fig. 4 A. Video is available at //www.jcb.org/cgi/content/full/153/2/319/DC1.
Dissipation and regeneration of ΔΨm after cytochrome c release in the absence of caspases. [A] Time lapse of the relative brightness of TMRE and the punctate–diffuse index of cytochrome c–GFP in one Cc-GFP-HeLa cell treated with actinomycin D [1 μM] in the presence of zVADfmk showing a drop in ΔΨm, followed by a slow regeneration of ΔΨm after cytochrome c–GFP release. Cytochrome c–GFP release was set at 120 min. [B] Pictographic representation of ΔΨm and cytochrome c–GFP in the cell depicted in A shows that ΔΨm is regenerated, whereas cytochrome c–GFP remains diffuse throughout the cell. More cells can be seen in Quicktime movie format at //www.jcb.org/cgi/content/full/153/2/319/DC1. In the movie, green [left] indicates cytochrome c–GFP, whereas red [right] indicates TMRE staining. [C and D] Time-lapse of the relative brightness of TMRE fluorescence of one Apaf-1–deficient murine embryonic fibroblast [apaf-deficient MEF] treated with 1 μM actinomycin D [C], one Cc-GFP-HeLa cell treated with 1 μM staurosporine in the presence of 100 μM zVADfmk [Di], or one Cc-GFP-HeLa cell treated with 1 μM staurosporine in the presence of 100 μM zVADfmk and 10 μg/ml of oligomycin [Dii]. Bars, 10 μm.
To examine the relationship between ΔΨm and the permeabilization of the outer mitochondrial membrane, we took advantage of a recently described HeLa cell line [Cc-GFP-HeLa] stably expressing cytochrome c–GFP in the mitochondrial intermembrane space [Goldstein et al. 2000]. In these cells, the release of cytochrome c–GFP faithfully mirrors the release of cytochrome c, and this occurs in a kinetically rapid all-or-nothing manner during apoptosis. We monitored ΔΨm using TMRE, which incorporates into the mitochondria in a nernstian manner in relation to ΔΨm [Farkas et al. 1989; Fink et al. 1998].
Previously, we have found that in CEM and HeLa cells treated with UV radiation in the presence of the caspase inhibitor zVADfmk, cytochrome c can be released without significantly affecting ΔΨm [Bossy-Wetzel et al. 1998]. To confirm this finding in the cell lines used in the this study, we treated Jurkat or Cc-GFP-HeLa cells with actinomycin D, staurosporine, UV, or etoposide and examined ΔΨm by TMRE staining. As shown in Fig. 1, loss of ΔΨm in Jurkat cells treated with etoposide or Cc-GFP-HeLa cells treated with actinomycin D was prevented by addition of the caspase inhibitor. Similar results were obtained in both cell lines treated with staurosporine and in Cc-GFP-HeLa cells treated with UV [data not shown], and when other dyes including CMTM-ROS and DiOC[6]3 were used to measure ΔΨm [not shown]. In every case, zVADfmk did not affect cytochrome c release in Cc-GFP-HeLa cells [Goldstein et al. 2000].
Loss of ΔΨm is caspase dependent during apoptosis. [A] Jurkat cells treated with etoposide [40 μM] or Cc-GFP-HeLa cells were treated with actinomycin D [1 μM] in the presence or absence of zVADfmk [100 μM], harvested at the times indicated, stained with TMRE [50 nM], and analyzed by flow cytometry. Low fluorescence indicates a loss of ΔΨm. [B] A representation of A, in which untreated cells [thin lines] are overlaid directly on cells treated with the apoptosis inducer in the presence of zVADfmk [zVAD] [100 μM]. In the Cc-GFP-Hela cells, 57% of cells had released cytochrome c–GFP by this point.
The maintenance of ΔΨm may be through either the electron transport chain or through ATP-dependent reversal of ATP synthase [Simbula et al. 1997]. We have previously shown that oligomycin, an ATP synthase inhibitor, does not dissipate the ΔΨm maintained after cytochrome c release, whereas inhibitors of complex III [which donates electrons to cytochrome c] and complex IV [which accepts electrons from cytochrome c] do [Goldstein et al. 2000]. This suggested that cytochrome c that is still present in the cell after its release from mitochondria is sufficient to maintain electron transport and ΔΨm.
To test this idea, we induced the release of intermembrane proteins from mitochondria in digitonin-permeabilized cells, using recombinant tBid to mimic the apoptotic process [von Ahsen et al. 2000]. As shown in Fig. 2 A, treatment with tBid caused a rapid loss of ΔΨm, to the same extent produced by the protonophore FCCP. Upon addition of cytochrome c, however, ΔΨm was maintained, despite the presence of oligomycin to block the generation of ΔΨm by hydrolysis of ATP. ΔΨm was not maintained in the presence of the complex IV inhibitor KCN. Thus, the loss of ΔΨm was due to the release of cytochrome c and not other mitochondrial changes.
Then, we examined the role of cytochrome c in maintaining ΔΨm in cells induced to undergo apoptosis in the presence of caspase inhibitors. Cc-GFP-HeLa cells were treated with actinomycin D plus zVADfmk, and cytochrome c–GFP release was monitored [not shown]. At a time point when the majority of cells exhibited cytochrome c–GFP release, the cells were treated with digitonin to permeabilize the plasma membrane. This resulted in a complete loss of ΔΨm [Fig. 2 B] as all available cytochrome c was washed free. Addition of exogenous cytochrome c significantly restored ΔΨm in these cells. Therefore, the outer membranes of mitochondria in permeabilized tBid-treated or apoptotic cells are permeable to cytochrome c, which can restore electron transport in these mitochondria. It follows that the maintenance of ΔΨm after mitochondrial outer membrane permeabilization is through the use of the low levels of cytosolic cytochrome c. By comparing the total amount of cytochrome c in these cells with standard concentrations of horse heart cytochrome c in immunoblots, we estimated that if cytochrome c were evenly dispersed throughout the cell, it would be ∼10 μM [not shown]. Addition of 10 μM cytochrome c in our permeabilized cell assays was sufficient to maintain the ΔΨm [Fig. 2 A].
Competing models that explain the permeabilization of the mitochondrial outer membrane make different predictions regarding changes in ΔΨm before the release of cytochrome c. Outer membrane rupture after permeability transition [Marzo et al. 1998b] or closure of the voltage-dependent anion channels [Vander Heiden and Thompson 1999] involve dissipation or increase in ΔΨm, respectively.
In time-lapse confocal microscopy experiments, we consistently observed that, upon addition of the protonophore FCCP, TMRE staining was lost in