Mitochondria and cell death : Mechanistic aspects and methodological issues

Eur. J. Biochem. 264, 687±701 (1999) q FEBS 1999

R E V I E W A R T I C L E

Mitochondria and cell deathMechanistic aspects and methodological issues

Paolo Bernardi1, Luca Scorrano1, Raffaele Colonna1, Valeria Petronilli1 and Fabio Di Lisa2

CNR Unit for the Study of Biomembranes and Departments of 1Biomedical Sciences and 2Biological Chemistry, University of Padova, Padova, Italy

Mitochondria are involved in cell death for reasons that go beyond ATP supply. A recent advance has been the

discovery that mitochondria contain and release proteins that are involved in the apoptotic cascade, like

cytochrome c and apoptosis inducing factor. The involvement of mitochondria in cell death, and its being cause

or consequence, remain issues that are extremely complex to address in situ. The response of mitochondria may

critically depend on the type of stimulus, on its intensity, and on the specific mitochondrial function that has been

primarily perturbed. On the other hand, the outcome also depends on the integration of mitochondrial responses

that cannot be dissected easily. Here, we try to identify the mechanistic aspects of mitochondrial involvement in

cell death as can be derived from our current understanding of mitochondrial physiology, with special emphasis

on the permeability transition and its consequences (like onset of swelling, cytochrome c release and respiratory

inhibition); and to critically evaluate methods that are widely used to monitor mitochondrial function in situ.

Keywords: mitochondria; cell death; apoptosis; necrosis; permeability transition; calcium; caspases; fluorescence;

probes; cytochrome c.

Mitochondria are central to the life of eukaryotic cells. In recentyears, it has become clear that mitochondria also play a key rolein the pathways to cell death [1±8]. This role of mitochondriacannot be explained by a mere `loss of function' resulting in anenergetic deficit. Rather, it is increasingly recognized as an`active' process mediated by regulated effector mechanisms ina wide variety of conditions. A large body of observations fromthe fields of immunology, toxicology, oncology, neurology andcardiology testifies about the centrality of this issue. However,the general reader is probably puzzled by the growing numberof conflicting reports, and by the diverging conclusions reachedby different groups when mechanistic issues are addressed and/or cause-relationship effects are discussed. A clear example isrepresented by the question of cytochrome c release, and ofhow this event relates to mitochondrial respiration, main-tenance of the mitochondrial membrane potential difference(Dcm) and volume homeostasis.

In our opinion, part of the problem resides in the intrinsicdifficulties one faces when measuring `mitochondrial function'in intact cells, an issue that is not always adequatelyappreciated. Mitochondrial responses in vivo may be extremely

hard to decipher, and a large discrepancy currently existsbetween our understanding of mitochondrial function in theisolated organelle and in the intact cell. However, an acceptabledescription of in vivo events must always take into accountfacts and concepts established in decades of in vitro studies.Also, it should be realized that discrepancies may arise frominapparent differences in the experimental conditions, but alsofrom artifacts or misinterpretation of the in situ experiments. Asshould become clear later in the review, particular care shouldbe exerted in studies with fluorescent probes. These are easilydetected by cytofluorimetric and imaging techniques, but oftenhave not been adequately characterized for phototoxicity andfor directs effects on mitochondrial function that may precludeor seriously limit their use with living cells.

Major goals of this review are to identify and discuss themechanistic aspects of mitochondrial involvement in cell deathas can be derived from our current understanding ofmitochondrial physiology; and to critically evaluate the keyissues emerged from recent studies, with the objective ofenucleating points of controversy and of identifying out-standing problems. This review is not intended to provide anextensive coverage of the literature, which can be found inrecent reviews [9±24].

T H E M E M B R A N E P O T E N T I A L ,M I T O C H O N D R I A L C A T I O N T R A N S P O R TA N D M A T R I X S W E L L I N G

The initial event of energy conservation is charge separation atthe inner mitochondrial membrane. Electrons deriving fromoxidation of substrates are funneled to oxygen through theredox carriers of the respiratory chain, and this process iscoupled to H+ ejection on the redox pumps at complexes I, IIIand IV. As the passive permeability to H+ and to cations andanions is generally low, H+ ejection results in the establishmentof a H+ electrochemical gradient, mÄH [25,26]. The magnitude

Correspondence to P. Bernardi/F. Di Lisa, CNR Unit for the Study of

Biomembranes, Viale Giuseppe Colombo 3, I-35121 Padova, Italy.

Fax: + 39 049827 6361,

E-mail: [email protected] or [email protected]

Abbreviations: Dcm, mitochondrial membrane potential difference; mÄH,

mitochondrial proton electrochemical gradient; Dcp, plasma membrane

potential difference; MTP, mitochondrial permeability transition pore; PT,

permeability transition; Cs, cyclosporin; PN, pyridine nucleotides; ANT,

adenine nucleotide translocase; AIF, apoptosis inducing factor; [Ca2+]m,

mitochondrial matrix free Ca2+ concentration; [Ca2+]c, cytosolic free Ca2+

concentration; DiOC6(3), 3,3 0-dihexyloxacarbocyanine iodide; CMTMRos,

chloromethyltetramethyl rosamine; FCCP, carbonylcyanide-p-trifluoro-

methoxyphenyl hydrazone; MDR, multidrug resistance pump; JC-1,

5,5 0,6,6 0-tetrachloro-1,1 0,3,3 0-tetraethylbenzimidazolocarbocyanine iodide;

TMRM, tetramethylrhodamine methyl ester; AM, acetoxymethyl.

(Received 8 April 1999, accepted 19 July 1999)

688 P. Bernardi et al. (Eur. J. Biochem. 264) q FEBS 1999

of the proton electrochemical gradient is about 2220 mV, andunder physiological conditions most of the gradient is in theform of a Dcm [27].

Monovalent cations

The inside-negative Dcm represents the major driving force formitochondrial cation accumulation. As the cytosolic concen-trations of K+ and Na+ are about 150 and 5 mm, respectively,for a Dcm of 2180 mV equilibrium matrix [K+] and [Na+]should be 150 m and 5 m, respectively. This is never achievedfor two main reasons: (a) the high permeability of the innermembrane to water, which prevents the buildup of a cationconcentration gradient and therefore maintains the matrixisoosmolar relative to the intermembrane and extramito-chondrial spaces despite cation uptake, a point that is notalways appreciated [28]; because of this, any net uptake ofcation salts down the electrochemical gradient, no matter howslow, would eventually lead to swelling with inner membraneunfolding and outer membrane rupture; (b) the operation of theelectroneutral H+-Na+ and H+-K+ exchangers, which catalyzecation efflux and thus prevent thermodynamic equilibration ofthe gradients; because of this, the cation accumulation ratio iskinetically modulated by the relative rates of uptake via the K+

and Na+ channels and efflux via the H+-Na+ and H+-K+

exchangers [29,30]. These fluxes are tightly regulated toachieve volume homeostasis without major energy drain. Adetailed coverage of these transport pathways, which is beyondthe scope of this review, can be found in [31].

Calcium

As cytosolic free [Ca2+] ([Ca2+]c) oscillates between about 0.1and 1 mm, for a Dcm of 2180 mV equilibrium matrix free[Ca2+] ([Ca2+]m) should be 0.1±1.0 m, i.e. at least 100 000 to1 000 000-fold higher than the actual values measured inisolated mitochondria and in intact cells [32]. Also in this casedisplacement from thermodynamic equilibrium is due to thefact that Ca2+ distribution represents a steady state whereelectrophoretic Ca2+ uptake via the Ca2+ uniporter and/or therapid uptake mode [33] is matched by Ca2+ efflux on twoseparate pathways, the `Na+-dependent Ca2+ efflux pathway'and the `Na+-independent Ca2+ efflux pathway' [34,35]. As themaximal rate of Ca2+ efflux via these pathways is much lowerthan that of Ca2+ uptake, mitochondria are exposed to thehazards of Ca2+ overload when [Ca2+]c rises steadily, or whenthe frequency of its oscillations increases. Due to the low[Ca2+]c and [Ca2+]m, and to the presence of Pi and otherCa2+-complexing anions and proteins in the matrix, Ca2+

uptake is unlikely to be a direct cause of mitochondrialswelling. In vitro at least, Ca2+-dependent swelling is rather dueto opening of a high-conductance channel, the mitochondrialpermeability transition pore (MTP) which is briefly coveredbelow. At variance from swelling that depends on K+ uptake,swelling caused by MTP opening occurs in fully depolarizedmitochondria, and it demands a concentration gradient ofsolute(s) between the intermembrane and matrix spaces.

T H E P E R M E A B I L I T Y T R A N S I T I O N

The permeability transition (PT) is a sudden increase of theinner membrane permeability to solutes with molecular massbelow approximately 1500 Da [36±38]. This phenomenon ismost easily observed after the matrix accumulation of Ca2+, andit is widely believed to be caused by opening of a regulated

channel. This channel, MTP, can be defined as a voltage-dependent, cyclosporin (Cs) A-sensitive, high-conductanceinner membrane channel. In the fully open state, the apparentpore diameter is about 3 nm [39] and the pore open±closedtransitions are highly regulated by multiple effectors at sitesthat are summarized in Table 1.

Many of these sites can be affected by conditions andmediators implied in a variety of models of cell death, andevidence exists to suggest that the PT may be an early event incommittment to apoptosis, particularly in the immune system[50]. Regulation of the MTP has recently been covered in detailelsewhere [31]. Here, we will rather focus on its consequencesbecause these are central to understanding its potential role incell death.

The only primary consequence of MTP opening in vitro ismembrane depolarization, and common PT assays in intact cellsare indeed based on the fluorescence changes of probes that areaccumulated by mitochondria in reponse to the Dcm. Thetheoretical and practical aspects of the use of probes, and theirlimits, will be discussed in some detail below. What needs to bestressed here is that although MTP opening is always followedby depolarization, depolarization is not always caused by MTPopening. It is unfortunate that many otherwise excellent studiesconsider depolarization in situ as an unequivocal evidence thata PT has occurred, which can be an obvious source ofmisunderstandings when the experimental results are inter-preted in mechanistic terms.

Pore opening can have consequences on respiration thatdepend on the substrates being oxidized. With pyridine nucleo-tide (PN)-linked substrates MTP opening is followed byrespiratory inhibition because matrix PN are lost, an experi-mental finding that preceded definition of the Ca2+-dependentpermeability change as a PT [51]. With complex II-linkedsubstrates the PT is rather followed by uncoupling. Theconsequences of a PT on respiration in vivo (and on the relatedissue of production of reactive oxygen species) thereforedepend on whether, and to what extent, PN depletion andsubsequent hydrolysis by outer membrane glycohydrolase takeplace [52]. Irrespective of whether respiration is inhibited orstimulated, collapse of the mÄH caused by the PT should curtailATP synthesis as long as the pore is open. Together withincreased hydrolysis by the mitochondrial ATPase, this shouldcontribute to cellular ATP depletion (reviewed in [53]).

Pore opening is followed by equilibration of ionic gradientsand of species that have a molecular mass lower than about

Table 1. Modulators of the mitochondrial permeability transition.

Open probability

Control point Ref. Increase Decrease

Voltage [40] Depolarization Hyperpolarization

Matrix pH [41] Alkalinization Acidification

Surface potential [42] More negative More positive

Matrix Me2+ site [37] Ca2+ Mg2+, Sr2+, Mn2+

External Me2+ site [43] ± Ca2+, Mg2+

ANT [36] `c' conformation `m' conformation

Cyclophilin D [44,45] ± CsA

Dithiols/glutathione [46,47] Oxidation Reduction

Pyridine nucleotides [46,47] Oxidation Reduction

Quinones [48,49] ± Ubiquinone 0

Decylubiquinone

q FEBS 1999 Mitochondria and cell death (Eur. J. Biochem. 264) 689

1500 Da, and this represents the basis for the common swellingassays of MTP opening in vitro. It must be stressed, however,that swelling can be easily prevented by low concentrations ofpore-impermeant solutes (such as 25 mm poly(ethylene glycol)3400) [54]; and that for short open times, solute equilibration(and therefore swelling) may not occur at all [55±57]. Thus,whether swelling, outer membrane rupture and cytochrome crelease necessarily follow a PT in vivo remains difficult topredict.

T W O M O D E S O F M I T O C H O N D R I A LS W E L L I N G

In summary, we can safely state that mitochondrial swelling isalways an osmotic process that results from net solute andwater diffusion towards the matrix. This may occur accordingto two basic mechanisms:

(a) Electrophoretic (energy-dependent) uptake of monovalentcations, K+ in particular. This type of swelling occurs withoutloss of energy other than that spent for the process of cationaccumulation. The mÄH is largely regenerated by respiration, theinner membrane permeability remains low, and the swollenmitochondria retain a high Dcm and a high level of coupling.Indeed, following energy-dependent swelling due to accumu-lation of K+ in the presence of valinomycin, mitochondria canutilize the K+ gradient to synthesize ATP [58]. This type ofswelling coincides with the `high energy' swelling defined byAzzone and Azzi in 1965 [59].

(b) Passive diffusion of species down their concentrationgradients following a decrease of the permeability barrier. Thistype of swelling can be caused by MTP opening, occurs indeenergized mitochondria (`low energy' swelling [59]), and isfollowed by repolarization upon pore closure and reener-gization with ATP or respiration. Shrinkage and volumerecovery are only possible when the species responsible forswelling can be extruded by endogenous transport pathways.This is the case for K+ and Na+ but not for sugars like sucrose,which is the standard osmotic support for studies with isolatedmitochondria, a choice that contributed to convey the mis-leading impression that pore-mediated swelling is an irrever-sible process.

Whether, and when, mitochondrial swelling occurs in theprogression towards cell death, whether swelling occurs with orwithout energy conservation and whether outer membranerupture is an essential requisite for release of intermembraneproteins involved in caspase and nuclease activation appear tobe central issues that need clarification in order to assess therole of mitochondria in cell death.

M I T O C H O N D R I A L U LT R A S T R U C T U R EA N D R E L E A S E O F I N T E R M E M B R A N EP R O T E I N S

A series of observations on thick sections (200±2000 nm orgreater) of mitochondria subjected to tomographic recon-struction after high-voltage electron microscopy has revealedan ultrastructure that differs significantly from textbookpictures based on interpretation of 2D images from thinsections (50±80 nm) of fixed and stained samples [60,61]. Themajor point is that the intercristal compartments are pleio-morphic structures that communicate with the peripheral (inter-membrane) space, and sometimes between themselves, throughnarrow tubular regions [62]. These findings suggest that theintercristal and intermembrane spaces could be functionally

distinct. This is consistent with several earlier observationsconcerning the status of mitochondrial cytochrome c.

The mitochondrial outer membrane possesses a rotenone-insensitive NADH cytochrome b5 reductase that is able toreduce cytochrome c in isolated mitochondria [63]. Only asmall fraction of cytochrome c can be reduced, which is thenreoxidized by cytochrome oxidase and thus catalyzes a shuttleof electrons between the outer and inner mitochondrialmembranes [64]. The fraction of cytochrome c that can bereduced by NADH via the outer membrane pathway (i.e. in thepresence of rotenone and antimycin A) varies from a minimumof 3% in low ionic strength media to a maximum of 10±15%above 80 mm KCl [64]. These findings define two pools ofcytochrome c: (a) a small pool of intermembrane cytochrome c,which can be reduced both by the bc1 complex at the innermembrane and by the NADH-cytochrome b5 reductase at theouter membrane; and (b) a larger pool of cytochrome c, whichcan only be reduced by the bc1 complex and would be localizedinside the intercristal compartments [64]. In this scenariopermeabilization of the outer membrane per se can onlyaccount for the release of about 10±15% of the totalcytochrome c, which could have a minimal impact onrespiration and therefore on maintenance of the Dcm. Swellingof the matrix would be required to make more cytochrome cavailable for release into the cytosol, with concomitantinhibition of respiration and depolarization (Fig. 1). Swellingshould not be necessarily intended as a massive, irreversibleand synchronous process, as swelling can reversibly occur in avariable fraction of mitochondria. Because apoptosis inducingfactor (AIF, see below) and caspases activated by cytochrome ccan feed back into the mitochondria to amplify the process, AIFand cytochrome c release could result into a self-amplifying (orself-limiting) process depending on the state of the cell deathsignalling machinery at the moment of their release. A rigorousassessment of the mechanism(s) of cytochrome c and AIFrelease, and of its relationship with mitochondrial depolar-ization, swelling and outer membrane rupture or permeabil-ization thus demands a careful measurement of severalvariables. We shall return to this problem after a discussionof the methods that are most widely used to assessmitochondrial function in the living cell.

A N A L Y S I S O F M I T O C H O N D R I A LF U N C T I O N I N S I T U

Monitoring mitochondrial function in situ is today performedessentially by means of fluorescence techniques. So far, mosttechniques that have been developed are for the study of Dcm

and intramitochondrial [Ca2+] ([Ca2+]m), two parameters whichare crucial in defining the role of mitochondria in cell death.These powerful methodologies are generating results whichcould not have been predicted on the basis of preexistingconcepts. These advances, which are largely related to the roleof mitochondria in apoptosis, have considerably expanded thescope of mitochondrial research thus contributing to animproved understanding of mitochondrial function in situ. Onthe other hand, a number of reports have been published thateither openly contrast with established concepts of mito-chondrial physiology, or reach conclusions that are underminedby a lack of appreciation of the limits of currently availablemethods. Far from presenting a probe directory, the followingparagraphs are intended to review the properties of fluorescentprobes with particular emphasis on potential sources of artifactsthat may prevent progress in this complex field of research.


M E A S U R E M E N T S O F T H EM I T O C H O N D R I A L M E M B R A N EP O T E N T I A L

At the single cell level, Dcm can be measured by using cationicfluorescent probes in conjunction with microscopy techniques.In the epifluorescence mode the emission is detected by aphotomultiplier tube as the total signal from the entire field ofobservation. In more complex setups the fluorescence image ofthe cell is acquired and digitized by means of video camerasand dedicated softwares that allow analysis of differences evenamong the mitochondria of a single cell. Although at highmagnification (of the order of 1000�) it is possible to analysesingle mitochondria [65], the prolonged exposure time requiredto obtain such a resolution dramatically affects both cell

viability and mitochondrial function. This problem is worsenedin the case of confocal microscopy, where cells are exposed tothe enormous power of laser beams.

Single cell studies are unique in that they allow thecontinuous monitoring of the relationships between variousparameters, such as, for example, Dcm and [Ca2+]c [1,65±71].These procedures, however, are limited by definition to thenumber of cells that can be analyzed at a time. When Dcm hasto be assessed in a large cell population, electrodes orradioactive tracers can be utilized to determine the distributionof lipophilic cations between cells and suspending buffers.Radioactive cations, such as[3H]triphenylmethylphosphonium,have also been used to measure Dcm in tissues [72,73].However, as these distribution analyses require several millioncells, flow cytometry is increasingly used to detect the

Fig. 1. Mitochondrial ultrastructure and release of cytochrome c. The scheme in panel A depicts an intramitochondrial compartment created by infoldings

of the inner membrane (i.m.). This intercristal space communicates with the intermembrane space through a narrow tubular region, as can be deduced by

tomographic reconstructions of thick sections after high-voltage electron microscopy [60±62]. Only a small fraction of cytochrome c is located in the

intermembrane space, where it can accept electrons from either the outer membrane (o.m.) rotenone-insensitive NADH-cytochrome b5 reductase or from the

inner membrane complex III. Proton pumping creates a mÄH, which is represented here in the form of a Dcm and denoted by signs. Note that H+ pumping also

occurs towards the compartments where most of cytochrome c is located. Panel B depicts the consequences of MTP opening, based on the assumption that a

solute gradient exists between the intermembrane and matrix spaces. The membrane potential collapses, and net solute influx is accompanied by water

resulting in the buildup of hydrostatic pressure pushing the matrix towards the intermembrane space. This may result in more cytochrome c gaining access to

the intermembrane space, possibly through widening of the tubular connecting regions (which remains entirely hypothetical). The increased pressure

eventually causes rupture of the outer membrane, with release of cytochrome c and other intermembrane soluble proteins (including AIF which is not

depicted here) in the extramitochondrial space (panel C).


fluorescence changes of a relatively small number of cells(103±105) loaded with the same probes utilized in fluorescencemicroscopy studies.

The ease with which one can get impressive images ofmitochondria in a living cell tends to dim the attention forpotential sources of artifacts, which always have to be takeninto account when fluorescence emissions are related tomitochondrial function. The following list highlights commonproblems which are frequently overlooked in studies ofmitochondrial function in situ.

Cellular uptake and extrusion

Once inside the cell, lipophilic cations are electrophoresed intomitochondria but their rate of uptake from the extracellularmedium depends on the plasma membrane potential (Dcp).Therefore, the intracellular distribution of the probe depends onboth Dcm and Dcp, and on the time of incubation [74]. It hasbeen demonstrated that depending on Dcp values, time ofincubation, absolute dye concentrations and cell/dye ratios thefluorescence emitted by the cyanine derivative 3,3 0-dihexyloxa-carbocyanine iodide [DiOC6(3)] can be significantly affected

by both Dcm and Dcp, the maximal response of fluorescencechanges to Dcm changes being observed at the lowest dye/cellratio and for probe concentrations lower than 40 nm [74]. Athigher concentrations the probe may be more responsive tochanges of Dcp than of Dcm, and it must be noted that arelevant part of the studies where changes of Dcm have beenrelated to apoptosis has been performed by flow cytometryusing DiOC6(3) at concentrations higher than 100 nm. Unfortu-nately, the possibility that apoptogenic substances or conditionscan affect Dcp has not been investigated in the same models,although one major hallmark of apoptosis is the flipping ofphosphatidylserine from the inner to the outer leaflet of theplasma membrane [75], which might in turn affect its electricalproperties. A further problem is that this cyanine derivativebinds to the endoplasmic reticulum as well [76], a property thatis indeed commercially advertised for the staining of thisorganelle. The signal coming from mitochondria and theendoplasmic reticulum may therefore be difficult to sort unlesscells are imaged.

The most important problem with the vast majority offluorescent probes for Dcm, however, is that their cellularaccumulation can be drastically reduced because of efficient

Fig. 2. Inhibition of the MDR pump improves mitochondrial loading with TMRM. Fluorescence images of MH1C1 cells loaded with 100 nm

tetramethylrhodamine methyl ester. The low level of mitochondrial fluorescence observed in control cells (panel A) is dramatically increased by treatment

with CsH (panel B), verapamil (panel C) or CsA (panel D). Bar, 20 mm.


extrusion by the multidrug-resistance pump (MDR). Expressionof this glycoprotein is indeed routinely investigated bymonitoring the decreased uptake of Rhodamine derivatives[77]; and the increased mitochondrial signal observed with Dcm

probes after the addition of CsA could be largely due to aneffect on the MDR pump. This is illustrated in Fig. 2, whichshows that the efficiency of loading of MH1C1 cells withTMRM is dramatically increased by verapamil, CsH or CsA,which all inhibit the MDR [78,79], while only CsA inhibits theMTP. A lack of effect of verapamil or CsH on mitochondrialfluorescence is therefore required before one can suspect, butnot prove, that the effect of CsA is on MTP, an essential controlthat is almost invariably missing.

Binding and quenching

The extent of accumulation of lipophilic cations withinmitochondria depends on both their initial extramitochondrialconcentration and the magnitude of Dcm. Very high intra-mitochondrial concentrations can in turn result in extensiveself-quenching of the dye fluorescence. Indeed, all the methodsfor Dcm estimation based on the fluorescence changes ofcationic probes in mitochondrial suspensions are based on themagnitude of quenching of the total fluorescence. Conversely,in both cell imaging and flow cytometry Dcm is estimated fromthe intensity of the cell fluorescence and not from thequenching of the total suspension fluorescence. Therefore, tomaximize the response of the cell fluorescence to themagnitude of Dcm it is necessary to minimize the quenchingof the dye associated with mitochondria. It has also to bepointed out that profound differences exist among the variousfluorescent probes concerning their ability to bind to mito-chondrial and other cellular membranes independent of Dcm.The fraction of the dye which actually responds to Dcm changesshould always be calibrated based on the fluorescence changesinduced by uncouplers.

Toxicity and bleaching

Owing to the combined driving forces of Dcm and Dcp,lipophilic cations are accumulated within the mitochondrialmatrix at concentrations exceeding those present in theextracellular milieu by approximately four orders of magnitude.Thus, phototoxic effects are likely to be elicited from thefluorescent molecules, causing in turn deleterious con-sequences. The imidazolic ring of histidyl residues of theadenine nucleotide translocase (ANT) is highly susceptible tophotodamage, particularly by singlet oxygen [80,81]. Also thesensitivity of MTP to various control factors, including Ca2+, isaltered by singlet oxygen produced by irradiation of hemato-porphyrin-loaded mitochondria [82]. In addition, and probablyindependent of photodynamic effects, several mitochondrialfunctions are inhibited by high concentrations of any potentio-metric fluorescent probe available. Among the mitochondrialtargets, complex I shows a high susceptibility to inhibition [74].Because of their (photo)toxic effects, potentiometric fluores-cent probes are in fact under investigation as potentialtherapeutic agents against various types of tumors [83±85],which appear to accumulate these molecules more efficientlythan normal, non transformed cells. In this context it is worthmentioning that merocyanine, one of the first Dcm probes inisolated mitochondria, is a powerful inducer of apoptosis[86,87], a finding that does support a role for mitochondria inthe execution of the cell death program. Thus, if dyeconcentrations and exposure times are not thoroughly tested,

the fluorescent probes could contribute to cause or potentiaterather than simply measure the changes of Dcm. The ensuingalterations of mitochondrial function and structure could resultin an increased production of oxyradicals [12]. It would thenbecome difficult to ascertain whether a decrease in cellularfluorescence depends on a fall of Dcm or rather on thebleaching caused by oxyradicals [65], an issue that is rarelygiven due attention.

The problem of `fixable' probes

Studies of mitochondrial involvement in cell death oftendemand the assessment of a series of parameters that mayrequire cell fixation, which is obviously not compatible with themaintenance of Dcm and leads to immediate release of the Dcm

probes previously accumulated by energized mitochondria. Tocircumvent this problem, Kroemer and coworkers introducedthe use of chloromethyl derivatives of the fluorescent cationicrosamine probes, which are marketed by Molecular Probes(Eugene, OR, USA) under the trademark of `mitotrackers'. Therationale of this approach is that the positively charged probewill be accumulated by energized mitochondria in response toDcm, followed by binding to mitochondrial SH groups viathe probe chloromethyl moiety. At this point, the covalentlybound probe should not be released despite deenergization, andtherefore stably `mark' the Dcm existing prior to disruption ofmembrane integrity. Macho et al. indeed showed that accumu-lation of chloromethyltetramethyl rosamine (CMTMRos) bythymocytes can be partly prevented by the uncouplercarbonylcyanide-p-trifluoromethoxyphenyl hydrazone (FCCP).After cell fixation, the ratio of CMTMRos fluorescence ofuntreated versus FCCP-pretreated cells under optimal con-ditions was measurable but extremely small [88]. It is obvious,however, that the probe cannot reliably measure a decrease ofDcm once it has bound matrix and membrane SH groups. Thiseasily explains why release of cytochrome c, caspase activationand poly(ADP-ribose) polymerase cleavage occurred prior toany detectable change of cellular CMTMRos fluorescence instaurosporine-induced apoptosis, a finding that has been takento mean that mitochondria maintained a high Dcm despiterelease of substantial amounts of cytochrome c [89]. To makematters worse, our recent studies demonstrate that CMTMRosis a powerful inducer of the mitochondrial PT and an inhibitorof respiratory complex I [90]. Thus, swelling due to a PTremains a plausible mechanism for cytochrome c release instaurosporine-induced apoptosis because CMTMRos maycause a PT-dependent depolarization that cannot be detectedby its own fluorescence changes.

Signal calibration

At variance from the case of isolated mitochondria, calibrationof the fluorescence signal with the magnitude of Dcm cannot beachieved in situ [74]. A further problem posed by the availablepotentiometric fluorescent dyes is that they belong to the singleexcitation-single emission type of molecules. Thus, a decreaseof the signal due to mitochondrial depolarization is indistin-guishable from a Dcm-independent leak of the dye into theextracellular medium, especially when prolonged incubationsare required. These problems preclude a meaningful com-parison between the fluorescent intensities observed in differentcells.

The cyanine derivative, 5,5 0,6,6 0-tetrachloro-1,1 0,3,3 0-tetra-ethylbenzimidazolo carbocyanine iodide (JC-1), is frequently(but quite improperly) referred to as a `ratiometric' probe for


Dcm [68,91,92]. JC-1 displays two major emission peaks (597and 539 nm with excitation at 490 nm), which correspond tothe multimeric and monomeric forms of the dye, respectively[91]. The red fluorescence of the multimer is observed inaqueous solution at high ionic strengths, whereas the greenfluorescence of the monomer is present when the probe islocated in hydrophobic environments [93]. In isolated mito-chondria both the monomer and the multimer emissions can beobserved. The green monomer emission was shown to beresponsive to values of Dcm below 2140 mV, a range ofmembrane potentials that hardly modified the multimeremission [93]. Conversely, the red multimer emission wasshown to respond to higher (more negative) values of Dcm. Inenergized cells loaded with JC-1 mitochondria display a brightred fluorescence, which is decreased by deenergization. At thesame time, deenergization causes an increase of the greenfluorescence reflecting an increase of the monomer concen-tration, but this is not limited to the mitochondrial membranes.Owing to the hydrophobic interactions of the monomer releasedfrom the matrix, the green fluorescence spreads all over cellularstructures (Fig. 3). Thus, only the multimer form measures theDcm-dependent mitochondrial accumulation, whereas the greenfluorescence depends on passive binding of JC-1 to any cellularmembrane; and comparing the ratios of the two emissions ofJC-1 is a questionable practice that relates phenomenaoccurring in different cellular regions.

Interactions between fluorescent molecules

When simultaneous imaging of two mitochondrial probes isrequired, the possible interference between probes shouldalways be checked in solution, although the results in situstill remain difficult to predict. For instance, we showed thatintramitochondrial calcein emission is quenched by exposinghepatocytes and hepatoma cells to high concentrations oftetramethylrhodamine methyl ester (TMRM) [57].

From the above points it emerges that the ideal Dcm

probe: (a) should accumulate within mitochondria only inresponse to Dcm; (b) should not be a substrate of the MDRpump; (c) should not be toxic; (d) should not bind passively tomitochondria or to other intracellular organelles; and (e) shouldnot cause photodynamic effects or other forms of photodamage.Although such an ideal probe is not available, reliable resultscan already be achieved by the correct choice of probe; bycarefully adjusting the dye/cell ratio to maximize the Dcm overDcp response; by checking the contribution of MDR activity toprobe loading with verapamil; by minimizing the illuminationintensity; and by performing proper controls for phototoxicityand other basic effects on mitochondrial function (such asrespiratory activity and maintenance of the membrane poten-tial). Among the many commercial molecules our preferencegoes to the rhodamine group, in particular TMRM, which in thelow nanomolar range exclusively stains the mitochondria and isnot retained by the cell upon Dcm collapse.

M E A S U R E M E N T S O F M I T O C H O N D R I A LF R E E [ C a 2 + ]

The first technique for monitoring [Ca2+]m in intact cellscapitalized on the fact that cell loading with fluorescent [Ca2+]indicators also caused probe localization within mitochondria[94]. In the so called Mn2+-quenching technique the incubationof rat cardiomyocytes with Mn2+ results in the disappearance ofthe cytosolic signal, leaving the mitochondrial fluorescence ofIndo-1 almost unaffected [94]. By developing this procedure

Stern, Silverman and Coworkers could study the relationshipbetween the value of [Ca2+]m attained during anoxia and thedeath or survival of individual cardiomyocytes upon reoxy-genation [95]. This allowed the definition of a threshold of250 nm [Ca2+]m as the `point of no return' for cardiomyocytesurvival. The reliability of the quenching technique requires thetotal disappearance of the cytosolic signal, which is demon-strated by the absence of Ca2+ transients in contractingcardiomyocytes [94]. However, it might be more difficult torule out a residual cytosolic contribution in other cell types and,in any case, the cation used for quenching can interfere withmitochondrial Ca2+ homeostasis.

Fig. 3. Effect of mitochondrial uncoupling on cellular JC-1 fluores-

cence. NRK cells were loaded with JC-1 as previously described [92], and

imaged by confocal microscopy with excitation wavelenght at 488 nm.

Emissions at 525 nm (green) and 585 nm (red) were collected simul-

taneously by using two separate channels on the detector assembly. The red

fluorescence reflects the multimer form of the dye and is localized within

mitochondria under control conditions (A). The addition of FCCP (B)

induced a prompt disappearance of the red fluorescence along with the

diffusion of the green fluorescence corresponding to the monomer form of

JC-1. Bar, 10 mm. For further explanation see text.


A different procedure to monitor [Ca2+]m in intact cells inculture was developed by targeting aequorin within mito-chondria [96]. The chemiluminescent signal of aequorin, aphotoprotein which emits light upon Ca2+ binding, is generallytoo low to be recorded at the single cell level, although thislimitation was recently overridden by using improved detectionmethods [97]. Nevertheless, the application of this technique tocell suspensions allowed the demonstration of the functionalcoupling between endoplasmic reticulum and mitochondria[98].

A third approach towards monitoring of [Ca2+]m was madepossible by the introduction of a rhodamine derivative, Rhod 2[99]. Its acetoxymethyl (AM) ester is the only cell-permeantCa2+ indicator that has a net positive charge, a property thatpromotes its sequestration into mitochondria. The utilization ofdihydro-Rhod 2-AM was reported to increase the selectivity ofmitochondrial loading [100]. In this case, the indicator becomesfluorescent only upon its oxidation, which should occurpreferentially within the mitochondria. The ease of loadingand the lack of interference with mitochondrial Ca2+ move-ments rapidly made Rhod 2 the most utilized probe for[Ca2+]m. However, the following points should be emphasized:(a) unexplained differences in the indicator batches affect thereproducibilty of the observations, as noted in severallaboratories including ours; (b) Rhod 2 is not a ratiometricprobe, and fluorescence changes could be produced by factorsother than variations in [Ca2+]m; (c) a variable contribution bythe cytosolic compartment can affect the mitochondrial signal;as an example, in a recent and elegant study pictures showingan exclusive mitochondrial localization of Rhod 2 coexistedwith images showing a diffuse pattern of fluorescence inuntreated cells [101]; and (d) the intracellular distribution canbe affected by MTP opening; indeed, Rhod 2 is releasedthrough the MTP in isolated mitochondria (V. Petronilli,unpublished observation) and CsA blocks the disappearanceof mitochondrial fluorescence in Rhod 2 loaded cardio-myocytes [102]; this hampers the use of Rhod 2 for investigat-ing the relationships between [Ca2+]m changes and MTPopening.

Despite these limitations, relevant results have been obtainedwith these methods. In particular, it was shown that matrix Ca2+

can undergo rapid changes [96], and that mitochondria canmodulate the frequency and the amplitude of cytosolic Ca2+

oscillations in living cells [103±105]. These findings revivedinterest in the role of mitochondria in cellular Ca2+ homeo-stasis, which had been neglected for almost 20 years (reviewedin [31]). One relevant achievement of these in situ measure-ments has been the demonstration that under resting conditions,[Ca2+]m in cardiomyocytes is lower than [Ca2+]c [95,106], afinding that rules out the possibility that mitochondria maycause cell damage by releasing Ca2+.

A link between Ca2+ overload and mitochondrial dysfunctionhas long been proposed, but its relevance in vivo remains hardto predict. Mitochondria can tolerate transient increases of[Ca2+]m exceeding 10 mm [97], while increases of [Ca2+]c in themicromolar range resulted in only slight variations of Dcm,which were perfectly reversible [65]. On the other hand, it mustbe pointed out that also a decrease in Ca2+ availability could beharmful for mitochondria, as mitochondrial Ca2+ uptake in situis accompanied by stimulation of ATP synthesis [107±109].Indeed, high-frequency shortening of cardiomyocytes isaccompanied by an increase of [Ca2+]m that stimulates energyproduction, and the lack of this [Ca2+]m response plays a pivotalrole in the evolution towards cardiomyocyte failure and death inthe syrian cardiomyopathic hamster [110].

Taken together, these observations indicate that valuableinformation on the involvement of mitochondria in cell deathmay be obtained from measurements of [Ca2+]m, and that therelationship between onset of apoptosis and changes of [Ca2+]m

should be investigated more thoroughly.

M E A S U R E M E N T S O F T H E P E R M E A B I L I T YT R A N S I T I O N

The role of MTP in cell physiology and pathology is still amatter of debate also because adequate methods to probe MTPdirectly in intact cells are lacking. The evidence for itsoccurrence in vivo is largely based on either indirect methods,such as measurements of mitochondrial depolarization, or onpharmacological tools, such as the effects of in vitro poreinducers or inhibitors (such as CsA). The shortcomings areobvious, as depolarization can be caused by a variety of events(most notably, increased ATP demand) while CsA also inter-feres with calcineurin-dependent signaling, which like MTP hasa prominent Ca2+-dependence [111]. Even MeVal-4-Cs, a CsAderivative which inhibits the pore but not calcineurin [45], stillinhibits all cellular cyclophilins and therefore is not selectivefor the MTP. In addition, the indirect approaches used so far donot allow to address the cellular modulation of MTP, leavingcrucial issues unsolved, such as the cause-effect relationshipbetween pore opening and Dcm decrease [112]. Theseconsiderations explain the intrinsic interest in developingunequivocal tools to measure MTP opening in living cells.

In principle, MTP can be probed directly by a moleculewhich is excluded (or retained) by mitochondria when the poreis closed, and is taken up by (or released from) mitochondriawhen the pore opens. Even with such a probe, however,selective inhibition by CsA would still be necessary to proveMTP involvement. Suitable molecules should meet thefollowing minimal requirements: (a) molecular mass lowerthan 1.5 kDa; (b) little or no hydrophobicity; and (c) lack ofutilization as a substrate by mitochondrial enzymes. Among theavailable fluorescent molecules, calcein has been selected asthe probe of choice to detect pore opening with imagingtechniques [113]. Although it is cell impermeant, calcein canbe easily loaded into cells by using its AM ester form whichis nonfluorescent. Once inside the cell the probe isdeesterified and trapped in its so-called free form, which isfluorescent ± a strategy commonly used to load cells withfluorescent probes.

The initial strategy in calcein utilization was based on thefinding of fluorescence voids corresponding to mitochondria inhepatocytes incubated with calcein-AM ester in the presence ofTMRM. The voids were interpreted as mitochondria excludingcalcein, and their filling with calcein was presented as aprocedure to monitor MTP in intact cells [113]. However, wefound that mitochondria are easily filled with calcein upon theincubation of many cell types with its AM ester. In addition,calcein fluorescence is quenched by TMRM within mitochon-dria, indicating that the voids were more likely to be producedby the intramitochondrial colocalization of the two dyes [57].More importantly, we found that the cytosolic signal can bequenched by incubating calcein-loaded cells with Co2+. Thisprocedure results in the appearance of mitochondria asglowing bodies over a dark background, and allows to studyMTP opening as the decrease of mitochondrially associatedcalcein fluorescence as a function of time. These studiesrevealed a spontaneous, slow decrease of mitochondrialcalcein fluorescence that was completely prevented by CsA,suggesting that MTP fluctuates rapidly between open and


closed states in intact cells [57]. It is noteworthy that nofluorescence changes could be detected by Dcm probes duringthe CsA-sensitive decrease of calcein fluorescence. AlthoughMTP opening must result in Dcm collapse, the response time ofavailable techniques is not fast enough to detect Dcm changesin the millisecond range, which is well above the open-closedtransitions of MTP [114]. MTP openings of short duration canrather be studied by the calcein loading-Co2+ quenchingmethod [57], a technique that should help characterizeconditions associated with MTP opening in situ, and to definethe causal relationships between pore opening, collapse of Dcm,perturbation of intracellular and mitochondrial Ca2+ home-ostasis, and release of apoptogenic proteins.

M I T O C H O N D R I A L P R O T E I N S I N V O L V E DI N C E L L D E A T H

The involvement of mitochondria in cell death has beeninvestigated for many years, particularly in relation to Ca2+

homeostasis. The idea that opening of the MTP could be afactor in ischemia-reperfusion and toxic damage, put forward inthe 1980s [115,116], is finding support from recent in vivostudies [117±120]. There is little doubt, however, that therecent impulse to mitochondrial studies in the context of celldeath came with the identification of mitochondrial proteinsthat participate in modulating the execution phase of apoptosis.These proteins can be grouped in two classes: (a) pro- and anti-apoptotic members of the Bcl-2 family that largely localize tothe outer mitochondrial membrane; and (b) proteins that may bereleased during apoptosis, like AIF and, quite unexpectedly,cytochrome c.

Bcl-2 family

This class of proteins includes both anti-apoptotic (such asBcl-2, Bcl-XL, Bcl-W and Mcl-1) and proapoptotic members(such as Bax, Bak and Bok). The two classes share highsequence homology except for the BH3 domain, which ispresent only in the proapoptotic proteins. Proteins of this familyassociate into dimers, and the cellular response to the deathsignals depends on the ratio of pro-to antiapoptotic molecules(reviewed in [121]). Studies of subcellular distribution indicatethat they localize both to the cytosol and to intracellularmembranes, including the nuclear envelope and the outermitochondrial membrane [122]. The consequences of locali-zation to the outer mitochondrial membrane appear to betwofold: (a) they provide docking sites for other proteinsinvolved in the death cascade, including the kinase Raf and thephosphatase calcineurin [123]; as these bound proteins remainenzymatically active, they can affect the formation ofheterodimers between Bcl-2 and other members of the super-family that lack the membrane insertion C-terminal domain,and only heterodimerize in the dephosphorylated form [121];(b) they appear to affect onset of the PT and/or release of AIF[124] and cytochrome c [125]; although the mechanismremains controversial (the relevant literature will be discussedin some detail below) it is intriguing that proapoptotic members(Bax) favor the PT and cytochrome c release while anti-apoptotic members (Bcl-2) make the PT more difficult, at leastwith some inducers; it has been suggested that these effectsmay be related to the mechanical properties conferred upon theouter membrane by the presence of Bcl-2 family proteins(reviewed in [18]).

It has been shown that Bcl-2 type proteins can form ionicchannels that are anion-selective for proapoptotic proteins andcation-selective for antiapoptotic proteins (reviewed in [126]).

The link between this channel activity, which has beenobserved under rather extreme conditions of pH in vitro, andthe role in cell death remains unclear. The outer membranelocation precludes direct effects on the Dcm and/or on thepermeability of the inner membrane, an issue that is not alwaysappreciated. In summary, we think that the reported effects ofBcl-2 family members on the probability of MTP opening andon cytochrome c release still await a mechanistic explanation.

Cytochrome c

In an effort to purify components required for in vitro activationof caspase 3, Liu et al. identified a fraction containingcytochrome c [7]. These Authors showed that in the presenceof dATP (mM) or ATP (mM) cytochrome c was able to activateprocaspase 9, followed by recruitment and activation ofprocaspase 3, the effector protease that cleaves lamin,poly(ADP-ribose) polymerase and fodrin [8]. Considerableevidence exists to show that cytochrome c release to the cytosoltakes place in a variety of models of apoptosis, yet a number ofproblems are apparent when the mechanistic aspects ofcytochrome c release are analyzed.

In a classical paper, Jacobs and Sanadi showed thatcytochrome c can be released by suspending mitochondria ina hypotonic medium, followed by a wash of the membranes inan isotonic saline medium [127]. The hypotonic treatment wasrequired to force mitochondrial swelling and outer membranerupture, while the salt wash was necessary to detachcytochrome c, which interacts with complexes III and IVmainly by virtue of electrostatic forces. Respiration wasinhibited after these treatments, but it could be restored bythe addition of exogenous cytochrome c [127]. Because ofthese observations, cytochrome c depletion has long beenknown to be a consequence of mitochondrial swelling in salt-containing media, irrespective of the cause of swelling; andstudies of ion transport in saline media are (or should be)routinely performed in cytochrome c-supplemented mito-chondria [128,129].

It should be appreciated that cytochrome c release requiresboth swelling and a high ionic strength. Swelling in hypotonic,sucrose-based media will not release cytochrome c, whichunder these conditions remains bound to the inner membranedespite outer membrane rupture; and incubation in isotonicsaline media alone will not release cytochrome c because theouter membrane does not allow its diffusion. These require-ments for cytochrome c release in vitro should be kept in mind,and can probably explain some discrepancies in the field.Indeed, apparent cytochrome c release could be caused by thetechniques used to disrupt cells prior to organelle separation,particularly if homogenization is carried out in salt-containingmedia. Indeed, it has been reported that cytochrome c releasecould not be detected in the course of apoptosis induced by Fasligation when Jurkat cells were disrupted by nitrogen cavitationrather than mechanical homogenization [130]. This observationshould induce some caution when the evidence for cyto-chrome c release is based solely on cell fractionation ratherthan on in situ methods.

AIF

After in vivo treatment of mice with dexamethazone, asubpopulation of lymphoid cells maintaining a lower Dcm, asassessed with a fluorescent probe, could be sorted out whichwould then undergo apoptosis in culture. Addition of CsAprevented the fluorescence decrease and delayed apoptosis,suggesting PT involvement in the apoptotic cascade [6]. A


correlation has been subsequently established between onset ofthe PT in vitro and appearance of the nuclear signs of apoptosisin a reconstituted system where nuclei were incubated withisolated mitochondria. Nuclear degradation was observed whena PT had occurred, while nuclei remained intact whenmitochondria did not undergo a PT irrespective of the methodsused to induce or inhibit the pore [131,132]. These observationsstrengthened the general theory of the mitochondrial control ofapoptosis, and the proposal that the PT is a key event in theeffector phase of programmed cell death [50]. The link betweenopening of the PT and nuclear degradation has been identifiedby the Kroemer group in a protein, AIF, which is associatedwith markers of the mitochondrial intermembrane space. AIF isa protease acting through proteolytic activation of a nuclearendonuclease; and it is inhibited by N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (a caspase inhibitor) but not byspecific inhibitors of known Ca2+, serine, or cysteine proteasesincluding caspase-3, all of which are involved in the apoptoticcascade [124]. Cloning has revealed that AIF is a flavoproteinwith predicted mass of 57 kDa which displays a strikinghomology with bacterial ferredoxin and NADH oxido-reductases. The protein contains FAD, which is required forthe oxidoreductase but not for the apoptogenic activity [133].

This list of proteins is by no means conclusive, asmitochondria also contain other proteases, including pro-caspase-3 [134], caspase-9 [135] and calpain-like species [20]that might participate in a positive feedback loop, or eventrigger mitochondrial apoptotic responses that may includeMTP opening.

C Y T O C H R O M E c R E L E A S E , B a x , Dc m

A N D T H E P E R M E A B I L I T Y T R A N S I T I O N

Right after the discovery that cytochrome c release was an earlyevent in apoptosis, researchers have tried to define the pathwaysfor cytochrome c release and its relationships with mito-chondrial function, in particular with outer membrane integrity,maintenance of Dcm, and occurrence of a PT. Before theanalysis of experimental observations, however, a few keypoints need to be discussed.

The first issue is the relationship between respiration andDcm, a problem that has been thoroughly investigated inBioenergetics. This relationship is complex and nonlinear[136±138], which in turn poses obvious questions: how muchof cytochrome c must be released before a measurable decreaseof Dcm can be expected? Is cytochrome c release mediated by asmall fraction of mitochondria, initially at least, or rather by allmitochondria in the cell? These questions demand on one handa quantitative assessment of cytochrome c release (i.e. adetermination of both released and mitochondrially associatedcytochrome c), and on the other an assessment of probesensitivity both to homogeneous and non homogeneous Dcm

changes. Finally, and most importantly, mitochondria canmaintain a high Dcm by hydrolyzing glycolytic ATP evenwhen respiration is completely blocked, a condition where norelationship can be expected between maintenance of Dcm andcytochrome c release. A careful analysis of the relevantliterature suggests that these problems have not always beengiven due consideration; and that some conclusions need to bereassessed.

The widespread conviction that apoptotic cytochrome c releaseprecedes rather than follows mitochondrial depolarization, andtherefore that it cannot be caused by a PT, originated from twostudies reported in 1997 [125,139]. Wang et al. investigated themodel of staurosporine-induced apoptosis and measured Dcm

by rhodamine 123 staining, followed by visualization ofmitochondria by laser confocal microscopy. They found thatstaurosporine did cause mitochondrial depolarization, butrelease of cytochrome c occurred before a detectable decreaseof Dcm [125]. These experiments are not conclusive for threemain reasons: (a) cytochrome c release was assessed aftermechanical disruption of the cells, and it could have occurredduring homogenization because the buffer contained enoughsalts to release cytochrome c; (b) although the rhodamine 123signal responded to a high concentration of uncoupler, noreliable calibration is possible in these protocols, and theoccurrence of a depolarization smaller than that caused byFCCP (or occurring in just a fraction of the mitochondria)cannot be excluded; and (c) in these protocols Dcm could havebeen effectively maintained by the mitochondrial hydrolysis ofglycolytic ATP even if release of all of cytochrome c hadcaused complete respiratory inhibition. A more recent studywhere Dcm and a cytochrome c/green fluorescent proteinchimera were simultaneously imaged indeed suggests thatafter staurosporine treatment mitochondrial depolarizationoccurs at the same time as cytochrome c release [140].

In the study by Newmeyer et al. experiments were carriedout in a cell-free apoptotic system where cytochrome c isspontaneously released from mitochondria. Despite the releaseof cytochrome c changes of Dcm were not observed in assaysbased on retention of DiOC6(3) that were otherwise fullysensitive to the addition of a protonophore [139]. Theseresults are easily explained by the fact that 2 mm ATP and anATP-regenerating system based on phosphocreatine andcreatine kinase were present in the assay. Indeed, the Dcm

generated by ATP hydrolysis is obviously unaffected byelectron flow while it can still be dissipated by protonophoricuncouplers.

The idea that cytochrome c release occurs from fullyenergized mitochondria has been considerably reinforced byrecent studies centered on Bid, a BH3 domain-containingprotein that interacts with Bcl-2 and Bax. After cleavage bycaspase-8 activated by stimulation of cell surface deathreceptors, the C-terminal portion of cytosolic Bid is able toinsert into the mitochondrial outer membrane [141±143]. ThisBid fragment appears to cause cytochrome c release directly(i.e. in the absence of mitochondrial swelling) in a process thatis antagonized by overexpression of Bcl-2, and was presumedto occur in the absence of mitochondrial depolarization [141].This latter conclusion needs a reassessment, however, as thisstudy used CMTMRos, which causes respiratory inhibition andinduction of the PT while being largely insensitive to changesof Dcm in living cells [90].

Irrespective of the issue of mechanism, a number of studieshave shown that mitochondrial overexpression of Bcl-2decreases release of cytochrome c and of AIF, the latter effecthaving been ascribed to inhibition of the PT [124]. Conversely,it has been reported that mitochondrial overexpression of theproapoptotic Bax facilitates cell death because it facilitates themitochondrial release of cytochrome c. Whether the lattereffect is due to a PT is perhaps the single most controversialissue of recent literature.

Pastorino et al. have reported that overexpression of Bax in aJurkat T cell line by stable transfection caused mitochondrialdepolarization, cytochrome c release, caspase 3 activation, andcell death that could be completely prevented by a combinationof CsA and of the phospholipase A2 inhibitor, aristolochic acid[144]. Similar results were reported by Bradham et al. in amodel where stimulation of the tumor necrosis factor a receptorinduced mitochondrial depolarization, cytochrome c release,


caspase activation and cell death, all events that could beprevented by CsA [145]. These findings led to the conclusionthat MTP opening was responsible for cytochrome c releaseand the subsequent downstream events.

In a similar study where Bax was overexpressed in HeLa andCOS cells, and cytochrome c release was assessed both in situand in isolated mitochondria, Eskes et al. reached just theopposite conclusion, as Bax-induced release of cytochome cwas not prevented by CsA either in intact cells or in isolatedmitochondria [146]. After 15 h of treatment CsA did notprevent release of cytochrome c, but this is not necessarily anargument against involvement of the PT at earlier time points,or even later in the cell death sequence. It is well establishedthat inhibition by CsA is transient in long time-frameexperiments even with isolated mitochondria [42], and Eskeset al. [146] did not assess whether release of cytochrome c byBax was accompanied (and therefore potentially caused) byswelling. Indeed, whether CsA inhibits the PT in vitro is noteasy to predict, and it remains possible that Bax overexpressionaffected the PT sensitivity to CsA directly, or by varying theretention of factors that are essential for MTP inhibition byCsA like ADP [147]. In summary, we feel that involvementof the PT in cytochrome c release cannot be ruled out atpresent. It remains quite possible that different mechanisms(both PT-dependent and PT-independent) may be operating indifferent experimental systems, or at different time points; orthat subtle, inapparent differences exist that will eventuallyexplain what today appear to be unresolved experimentaldiscrepancies.

In conclusion, we believe that the renewed enthusiasm aboutmitochondria in cell death is fully justified. The key role ofmitochondria in energy production, the mitochondrial local-ization of pro- and anti-apoptotic proteins, and the prominentposition of mitochondria in cellular Ca2+ homeostasis allsuggest potential points of regulation. We hope that this reviewwill help address the mechanistic issues that still need to besolved.

A C K N O W L E D G E M E N T S

Research in our laboratories is supported by the Ministero per l'Universita 0

e la Ricerca Scientifica e Tecnologica, Projects `Bioenergetics and

Membrane Transport' (P. B.) and `Favouring myocardium viability to

necrosis'. A challenge in ischemic heart disease: molecular mechanisms

and clinical relevance' (F. D. L.); the Consiglio Nazionale delle Ricerche

(Dotazione Centro Biomembrane and Grant n. 98.00413.CT04 to P. B.);

Telethon-Italy Grant No. 1141 (P. B.); the National Institutes of Health

(USA) Grant No. 1R21 Gm58792 (P. B.); and the Giovanni Armenise-

Harvard Foundation (P. B.).

R E F E R E N C E S

1. Lemasters, J.J., DiGiuseppi, J., Nieminen, A.L. & Herman, B. (1987)

Blebbing, free Ca2+ and mitochondrial membrane potential pre-

ceding cell death in hepatocytes. Nature 325, 78±81.

2. Hockenbery, D.M., NunÄez, G., Milliman, C., Schreiber, R.D. &

Korsmeyer, S.J. (1990) Bcl-2 is an inner mitochondrial membrane

protein that blocks programmed cell death. Nature 348, 334±336.

3. Pastorino, J.G., Snyder, J.W., Serroni, A., Hoek, J.B. & Farber, J.L.

(1993) Cyclosporin and carnitine prevent the anoxic death of

cultured hepatocytes by inhibiting the mitochondrial permeability

transition. J. Biol. Chem. 268, 13791±13798.

4. Imberti, R., Nieminen, A.-L., Herman, B. & Lemasters, J.J. (1993)

Mitochondrial and glycolytic dysfunction in lethal injury to

hepatocytes by t-butylhydroperoxide: protection by fructose,

cyclosporin A and trifluoperazine. J. Pharmacol. Exp. Ther. 265,

392±400.

5. Ankarcrona, M., Dypbukt, J.M., Bonfoco, E., Zhivotovsky, B.,

Orrenius, S., Lipton, S.A. & Nicotera, P. (1995) Glutamate-induced

neuronal death: a succession of necrosis or apoptosis depending on

mitochondrial function. Neuron 15, 961±973.

6. Zamzami, N., Marchetti, P., Castedo, M., Decaudin, D., Macho, A.,

Hirsch, T., Susin, S.A., Petit, P.X., Mignotte, B. & Kroemer, G.

(1995) Sequential reduction of mitochondrial transmembrane

potential and generation of reactive oxygen species in early

programmed cell death. J. Exp. Med. 182, 367±377.

7. Liu, X., Kim, C.N., Yang, J., Jemmerson, R. & Wang, X. (1996)

Induction of apoptotic program in cell-free extracts: requirement for

dATP and cytochrome c. Cell 86, 147±157.

8. Zou, H., Henzel, W.J., Liu, X., Lutschg, A. & Wang, X. (1997)

Apaf-1, a human protein homologous to C. elegans CED-4,

participates in cytochrome c-dependent activation of caspase-3.

Cell 90, 405±413.

9. Mignotte, B. & Vayssiere, J.L. (1998) Mitochondria and apoptosis.

Eur. J. Biochem. 252, 1±15.

10. Bernardi, P. (1998) Mitochondria in cell death (preface). Biochim.

Biophys. Acta 1366, 1±2.

11. Ichas, F. & Mazat, J.-P. (1998) From calcium signalling to cell death:

two conformations for the mitochondrial permeability transition

pore. Biochim. Biophys. Acta 1366, 33±50.

12. Lenaz, G. (1998) Role of mitochondria in oxidative stress and aging.

Biochim. Biophys. Acta 1366, 53±67.

13. Di Lisa, F., MenaboÁ, R., Canton, M. & Petronilli, V. (1998) The role

of mitochondria in the salvage and the injury of the ischemic

myocardium. Biochim. Biophys. Acta 1366, 69±78.

14. Halestrap, A.P., Kerr, P.M., Javadov, S. & Woodfield, K.-Y. (1998)

Elucidating the molecular mechanism of the permeability transition

pore and its role in reperfusion injury. Biochim. Biophys. Acta 1366,

79±94.

15. Nicholls, D.G. & Budd, S.L. (1998) Mitochondria and glutamate

neurotoxicity. Biochim. Biophys. Acta 1366, 97±112.

16. Montal, M. (1998) Mitochondria, glutamate neurotoxicity and the

death cascade. Biochim. Biophys. Acta 1366, 113±126.

17. Reed, J.C., Jurgensmeier, J. & Matsuyama, S. (1998) Bcl-2 family

proteins and mitochondria. Biochim. Biophys. Acta 1366, 127±137.

18. Cai, J., Yang, J. & Jones, D.P. (1998) Mitochondrial control of

apoptosis: the role of cytochrome c. Biochim. Biophys. Acta 1366,

139±149.

19. Susin, S.A., Zamzami, N. & Kroemer, G. (1998) Mitochondria as

regulators of apoptosis: doubts no more. Biochim. Biophys. Acta

1366, 151±165.

20. Gores, G.J., Miyoshi, H., Botla, R., Aguilar, H.I. & Bronk, S.F. (1998)

Induction of the mitochondrial permeability transition as a

mechanism of liver injury during cholestasis: a potential role for

mitochondrial proteases. Biochim. Biophys. Acta 1366, 167±175.

21. Lemasters, J.J., Nieminen, A.L., Qian, T., Trost, L., Elmore, S.P.,

Nishimura, Y., Crowe, R.A., Cascio, W.E., Bradham, C.A., Brenner,

D.A. & Herman, B. (1998) The mitochondrial permeability

transition in cell death: a common mechanism in necrosis, apoptosis

and autophagy. Biochim. Biophys. Acta 1366, 177±196.

22. Di Mauro, S., Bonilla, E., Davidson, M., Hirano, M. & Schon, E.A.

(1998) Mitochondria in neuromuscular disorders. Biochim. Biophys.

Acta 1366, 199±210.

23. Beal, M.F. (1998) Mitochondrial dysfunction in neurodegenerative

diseases. Biochim. Biophys. Acta 1366, 211±223.

24. Schapira, A.H.V. (1998) Mitochondrial dysfunction in neuro-

degenerative disorders. Biochim. Biophys. Acta 1366, 225±233.

25. Mitchell, P. (1966) Chemiosmotic Coupling in Oxidative and

Photosynthetic Phosphorylation. Glynn Research, Bodmin, Corn-

wall, England.

26. Mitchell, P. (1979) Keilin's respiratory chain concept and its

chemiosmotic consequences. Science 206, 1148±1159.

27. Azzone, G.F., Pietrobon, D. & Zoratti, M. (1984) Determination of the


proton electrochemical gradient across biological membranes. Curr.

Top. Bioenerg. 13, 1±77.

28. Green, D.R. & Reed, J.C. (1998) Mitochondria and apoptosis. Science

281, 1309±1312.

29. Brierley, G.P., Baysal, K. & Jung, D.W. (1994) Cation transport

systems in mitochondria: Na+ and K+ uniports and exchangers.

J. Bioenerg. Biomembr. 26, 519±526.

30. Garlid, K.D. (1996) Cation transport in mitochondria ± the potassium

cycle. Biochim. Biophys. Acta 1275, 123±126.

31. Bernardi, P. (1999) Mitochondrial transport of cations: channels,

exchangers and permeability transition. Physiol. Rev. 79 (in press).

32. Pozzan, T., Bragadin, M. & Azzone, G.F. (1977) Disequilibrium

between steady-state Ca2+ accumulation ratio and membrane

potential in mitochondria. Pathway and role of Ca2+ efflux.

Biochemistry 16, 5618±5625.

33. Sparagna, G.C., Gunter, K.K., Sheu, S.S. & Gunter, T.E. (1995)

Mitochondrial calcium uptake from physiological-type pulses of

calcium. A description of the rapid uptake mode. J. Biol. Chem. 270,

27510±27515.

34. Gunter, T.E., Gunter, K.K., Sheu, S.S. & Gavin, C.E. (1994)

Mitochondrial calcium transport: physiological and pathological

relevance. Am. J. Physiol. 267, C313±C339.

35. Gunter, T.E., Buntinas, L. & Gunter, K.K. (1998) The Ca2+ transport

mechanisms of mitochondria and Ca2+ uptake from physiological-

type Ca2+ transients. Biochim. Biophys. Acta 1399, 5±15.

36. Hunter, D.R. & Haworth, R.A. (1979) The Ca2+-induced membrane

transition in mitochondria. I. The protective mechanisms. Arch.

Biochem. Biophys. 195, 453±459.

37. Haworth, R.A. & Hunter, D.R. (1979) The Ca2+-induced membrane

transition in mitochondria. II. Nature of the Ca2+ trigger site. Arch.


38. Hunter, D.R. & Haworth, R.A. (1979) The Ca2+-induced membrane

transition in mitochondria. III. Transitional Ca2+ release. Arch.


39. Massari, S. & Azzone, G.F. (1972) The equivalent pore radius of

intact and damaged mitochondria and the mechanism of active

shrinkage. Biochim. Biophys. Acta 283, 23±29.

40. Bernardi, P. (1992) Modulation of the mitochondrial cyclosporin

A-sensitive permeability transition pore by the proton electro-

chemical gradient. Evidence that the pore can be opened by

membrane depolarization. J. Biol. Chem. 267, 8834±8839.

41. Bernardi, P., Vassanelli, S., Veronese, P., Colonna, R., Szabo, I. &

Zoratti, M. (1992) Modulation of the mitochondrial permeability

transition pore. Effect of protons and divalent cations. J. Biol. Chem.

267, 2934±2939.

42. Broekemeier, K.M. & Pfeiffer, D.R. (1995) Inhibition of the

mitochondrial permeability transition by cyclosporin A during

long time frame experiments: relationship between pore opening

and the activity of mitochondrial phospholipases. Biochemistry 34,

16440±16449.

43. Bernardi, P., Veronese, P. & Petronilli, V. (1993) Modulation of the

mitochondrial cyclosporin A-sensitive permeability transition pore.

I. Evidence for two separate Me2+ binding sites with opposing

effects on the pore open probability. J. Biol. Chem. 268, 1005±1010.

44. Connern, C.P. & Halestrap, A.P. (1996) Chaotropic agents and

increased matrix volume enhance binding of mitochondrial

cyclophilin to the inner mitochondrial membrane and sensitize the

mitochondrial permeability transition to [Ca2+]. Biochemistry 35,

8172±8180.

45. Nicolli, A., Basso, E., Petronilli, V., Wenger, R.M. & Bernardi, P.

(1996) Interactions of cyclophilin with the mitochondrial inner

membrane and regulation of the permeability transition pore, a

cyclosporin A-sensitive channel. J. Biol. Chem. 271, 2185±2192.

46. Chernyak, B.V. & Bernardi, P. (1996) The mitochondrial permeability

transition pore is modulated by oxidative agents through both

pyridine nucleotides and glutathione at two separate sites. Eur. J.

Biochem. 238, 623±630.

47. Costantini, P., Chernyak, B.V., Petronilli, V. & Bernardi, P. (1996)

Modulation of the mitochondrial permeability transition pore by

pyridine nucleotides and dithiol oxidation at two separate sites.

J. Biol. Chem. 271, 6746±6751.

48. Fontaine, E., Eriksson, O., Ichas, F. & Bernardi, P. (1998) Regulation

of the permeability transition pore in skeletal muscle mitochondria.

Modulation by electron flow through the respiratory chain complex

I. J. Biol. Chem. 273, 12662±12668.

49. Fontaine, E., Ichas, F. & Bernardi, P. (1998) A ubiquinone-binding

site regulates the mitochondrial permeability transition pore. J. Biol.

Chem. 273, 25734±25740.

50. Kroemer, G., Dallaporta, B. & Resche-Rigon, M. (1998) The

mitochondrial death/life regulator in apoptosis and necrosis. Annu.

Rev. Physiol. 60, 619±642.

51. Vinogradov, A., Scarpa, A. & Chance, B. (1972) Calcium and

pyridine nucleotide interaction in mitochondrial membranes. Arch.


52. Scott Boyer, C., Moore, G.A. & Moldeus, P. (1993) Submitochondrial

localization of the NAD+ glycohydrolase. Implications for the

role of pyridine nucleotide hydrolysis in mitochondrial calcium

fluxes. J. Biol. Chem. 268, 4016±4020.

53. Di Lisa, F. & Bernardi, P. (1998) Mitochondrial function as a

determinant of recovery or death in cell response to injury. Mol. Cell

Biochem. 184, 379±391.

54. Pfeiffer, D.R., Gudz, T.I., Novgorodov, S.A. & Erdahl, W.L. (1995)

The peptide mastoparan is a potent facilitator of the mitochondrial

permeability transition. J. Biol. Chem. 270, 4923±4932.

55. Ichas, F., Jouaville, L.S. & Mazat, J.-P. (1997) Mitochondria are

excitable organelles capable of generating and conveying electrical

and calcium signals. Cell 89, 1145±1153.

56. HuÈser, J., Rechenmacher, C.E. & Blatter, L.A. (1998) Imaging the

permeability pore transition in single mitochondria. Biophys. J. 74,

2129±2137.

57. Petronilli, V., Miotto, G., Canton, M., Colonna, R., Bernardi, P. & Di

Lisa, F. (1999) Transient and long-lasting openings of the

mitochondrial permeability transition pore can be monitored

directly in intact cells by changes of mitochondrial calcein

fluorescence. Biophys. J. 76, 725±734.

58. Rossi, E. & Azzone, G.F. (1970) The mechanism of ion translocation

in mitochondria. 3. Coupling of K+ efflux with ATP synthesis. Eur.

J. Biochem. 12, 319±327.

59. Azzone, G.F. & Azzi, A. (1965) Volume changes in liver

mitochondria. Proc. Natl Acad. Sci. USA 53, 1084±1089.

60. Mannella, C.A., Marko, M., Penczek, P., Barnard, D. & Frank, J.

(1994) The internal compartmentation of rat-liver mitochondria:

tomographic study using the high-voltage transmission electron

microscope. Microsc. Res. Technical 27, 278±283.

61. Perkins, G., Renken, C., Martone, M.E., Young, S.J., Ellisman, M. &

Frey, T. (1997) Electron tomography of neuronal mitochondria:

three-dimensional structure and organization of cristae and

membrane contacts. J. Struct. Biol. 119, 260±272.

62. Mannella, C.A., Marko, M. & Buttle, K. (1997) Reconsidering

mitochondrial structure: new views of an old organelle. Trends.

Biochem. Sci. 22, 37±38.

63. Sottocasa, G.L., Kuylenstierna, B., Ernster, L. & Bergstrand, A.

(1967) An electron-transport system associated with the outer

membrane of liver mitochondria. J. Cell Biol. 32, 415±438.

64. Bernardi, P. & Azzone, G.F. (1981) Cytochrome c as an electron

shuttle between the outer and inner mitochondrial membranes.

J. Biol. Chem. 256, 7187±7192.

65. Loew, L.M., Carrington, W., Tuft, R.A. & Fay, F.S. (1994)

Physiological cytosolic Ca2+ transients evoke concurrent mitochon-

drial depolarizations. Proc. Natl Acad. Sci. USA 91, 12579±12583.

66. Zoeteweij, J.P., van de Water, B., de Bont, H.J., Mulder, G.J. &

Nagelkerke, J.F. (1993) Calcium-induced cytotoxicity in hepato-

cytes after exposure to extracellular ATP is dependent on inorganic

phosphate. Effects on mitochondrial calcium. J. Biol. Chem. 268,

3384±3388.

67. Schinder, A.F., Olson, E.C., Spitzer, N.C. & Montal, M. (1996)

Mitochondrial dysfunction is a primary event in glutamate

neurotoxicity. J. Neurosci. 16, 6125±6133.


68. White, R.J. & Reynolds, I.J. (1996) Mitochondrial depolarization in

glutamate-stimulated neurons: an early signal specific to excitotoxin

exposure. J. Neurosci. 16, 5688±5697.

69. Budd, S.L. & Nicholls, D.G. (1996) A reevaluation of the role of

mitochondria in neuronal Ca2+ homeostasis. J. Neurochem. 66,

403±411.

70. Leyssens, A., Nowicky, A.V., Patterson, L., Crompton, M. & Duchen,

M.R. (1996) The relationship between mitochondrial state, ATP

hydrolysis, [Mg2+]i and [Ca2+]i studied in isolated rat cardio-

myocytes. J. Physiol. (London) 496, 111±128.

71. Delcamp, T.J., Dales, C., Ralenkotter, L., Cole, P.S. & Hadley, R.W.

(1998) Intramitochondrial [Ca2+] and membrane potential in

ventricular myocytes exposed to anoxia-reoxygenation. Am. J.

Physiol. 275, H484±H494.

72. Hoek, J.B., Nicholls, D.G. & Williamson, J.R. (1980) Determination

of the mitochondrial protonmotive force in isolated hepatocytes.

J. Biol. Chem. 255, 1458±1464.

73. Wan, B., Doumen, C., Duszynski, J., Salama, G. & LaNoue, K.F.

(1993) A method of determining electrical potential gradient across

mitochondrial membrane in perfused rat hearts. Am. J. Physiol. 265,

H445±H452.

74. Rottenberg, H. & Wu, S. (1998) Quantitative assay by flow cytometry

of the mitochondrial membrane potential in intact cells. Biochim.

Biophys. Acta 1404, 393±404.

75. Fadok, V.A., Voelker, D.R., Campbell, P.A., Cohen, J.J., Bratton, D.L.

& Henson, P.M. (1992) Exposure of phosphatidylserine on the

surface of apoptotic lymphocytes triggers specific recognition and

removal by macrophages. J. Immunol. 148, 2207±2216.

76. Terasaki, M. (1989) Fluorescent labeling of endoplasmic reticulum.

Methods Cell Biol. 29, 125±135.

77. Petriz, J. & Garcia-Lopez, J. (1997) Flow cytometric analysis of

P-glycoprotein function using rhodamine 123. Leukemia 11,

1124±1130.

78. Eytan, G.D., Regev, R., Oren, G., Hurwitz, C.D. & Assaraf, Y.G.

(1997) Efficiency of P-glycoprotein-mediated exclusion of rho-

damine dyes from multidrug-resistant cells is determined by their

passive transmembrane movement rate. Eur. J. Biochem. 248,

104±112.

79. Litman, T., Zeuthen, T., Skovsgaard, T. & Stein, W.D. (1997)

Structure-activity relationships of P-glycoprotein interacting drugs:

kinetic characterization of their effects on ATPase activity. Biochim.

Biophys. Acta 1361, 159±168.

80. Atlante, A., Passarella, S., Quagliariello, E., Moreno, G. & Salet, C.

(1989) Haematoporphyrin derivative (Photofrin II) photosen-

sitization of isolated mitochondria: inhibition of ADP/ATP trans-

locator. J. Photochem. Photobiol. B4, 35±46.

81. Salet, C. & Moreno, G. (1990) Photosensitization of mitochondria.

Molecular and cellular aspects. J. Photochem. Photobiol. B5,

133±150.

82. Salet, C., Moreno, G., Ricchelli, F. & Bernardi, P. (1997) Singlet

oxygen produced by photodynamic action causes inactivation of the

mitochondrial permeability transition pore. J. Biol. Chem. 272,

21938±21943.

83. Bernal, S.D., Shapiro, H.M. & Chen, L.B. (1982) Monitoring the

effect of anti-cancer drugs on L1210 cells by a mitochondrial probe,

rhodamine-123. Int. J. Cancer 30, 219±224.

84. Davis, S., Weiss, M.J., Wong, J.R., Lampidis, T.J. & Chen, L.B.

(1985) Mitochondrial and plasma membrane potentials cause

unusual accumulation and retention of rhodamine 123 by human

breast adenocarcinoma-derived MCF-7 cells. J. Biol. Chem. 260,

13844±13850.

85. Arcadi, J.A. (1998) The effect of rhodamine-123 on 3 prostate tumors

from the rat. J. Urol. 160, 2402±2406.

86. Gulliya, K.S., Sharma, R., Liu, H.W., Arnold, L. & Matthews, J.L.

(1995) Relationship of mitochondrial function and cellular adeno-

sine triphosphate levels to pMC540 and merodantoin cytotoxicity in

MCF-7 human breast cancer cells. Anticancer Drugs 6, 545±552.

87. Rottenberg, H. & Wu, S. (1997) Mitochondrial dysfunction in

lymphocytes from old mice: enhanced activation of the permeability

transition. Biochem. Biophys. Res. Commun. 240, 68±74.

88. Macho, A., Decaudin, D., Castedo, M., Hirsch, T., Susin, S.A.,

Zamzami, N. & Kroemer, G. (1996) Chloromethyl-X-Rosamine is

an aldehyde-fixable potential-sensitive fluorochrome for the detec-

tion of early apoptosis. Cytometry 25, 333±340.

89. Bossy-Wetzel, E., Newmeyer, D.D. & Green, D.R. (1998) Mito-

chondrial cytochrome c release in apoptosis occurs upstream of

DEVD-specific caspase activation and independently of mito-

chondrial transmembrane depolarization. EMBO J. 17, 37±49.

90. Scorrano, L., Petronilli, V., Colonna, R., Di Lisa, F. & Bernardi, P.

(1999) Chloromethyltetramethylrosamine (Mitotracker OrangeTM)

induces the mitochondrial permeability transition and inhibits

respiratory complex I. Implications for the mechanism of cyto-

chrome c release. J. Biol. Chem. 274, in press.

91. Reers, M., Smith, T.W. & Chen, L.B. (1991) J-aggregate formation of

a carbocyanine as a quantitative fluorescent indicator of membrane

potential. Biochemistry 30, 4480±4486.

92. Cossarizza, A., Ceccarelli, D. & Masini, A. (1996) Functional

heterogeneity of an isolated mitochondrial population revealed by

cytofluorometric analysis at the single organelle level. Exp. Cell

Res. 222, 84±94.

93. Di Lisa, F., Blank, P.S., Colonna, R., Gambassi, G., Silverman, H.S.,

Stern, M.D. & Hansford, R.G. (1995) Mitochondrial membrane

potential in single living adult rat cardiac myocytes exposed to

anoxia or metabolic inhibition. J. Physiol. (London) 486, 1±13.

94. Miyata, H., Silverman, H.S., Sollott, S.J., Lakatta, E.G., Stern, M.D.

& Hansford, R.G. (1991) Measurement of mitochondrial free Ca2+

concentration in living single rat cardiac myocytes. Am. J. Physiol.

261, H1223±H1234.

95. Miyata, H., Lakatta, E.G., Stern, M.D. & Silverman, H.S. (1992)

Relation of mitochondrial and cytosolic free calcium to cardiac

myocyte recovery after exposure to anoxia. Circ. Res. 71, 605±613.

96. Rizzuto, R., Simpson, A.W., Brini, M. & Pozzan, T. (1992) Rapid

changes of mitochondrial Ca2+ revealed by specifically targeted

recombinant aequorin. Nature 358, 325±327.

97. Rutter, G.A., Burnett, P., Rizzuto, R., Brini, M., Murgia, M., Pozzan,

T., Tavare, J.M. & Denton, R.M. (1996) Subcellular imaging of

intramitochondrial Ca2+ with recombinant targeted aequorin:

significance for the regulation of pyruvate dehydrogenase activity.

Proc. Natl Acad. Sci. USA 93, 5489±5494.

98. Rizzuto, R., Pinton, P., Carrington, W., Fay, F.S., Fogarty, K.E.,

Lifshitz, L.M., Tuft, R.A. & Pozzan, T. (1998) Close contacts with

the endoplasmic reticulum as determinants of mitochondrial Ca2+

responses. Science 280, 1763±1766.

99. Minta, A., Kao, J.P. & Tsien, R.Y. (1989) Fluorescent indicators for

cytosolic calcium based on rhodamine and fluorescein chromo-

phores. J. Biol. Chem. 264, 8171±8178.

100. Hajnoczky, G., Robb-Gaspers, L.D., Seitz, M.B. & Thomas, A.P.

(1995) Decoding of cytosolic calcium oscillations in the mito-

chondria. Cell 82, 415±424.

101. Boitier, E., Rea, R. & Duchen, M.R. (1999) Mitochondria exert a

negative feedback on the propagation of intracellular Ca2+ waves in

rat cortical astrocytes. J. Cell Biol. 145, 795±808.

102. Bowser, D.N., Minamikawa, T., Nagley, P. & Williams, D.A. (1998)

Role of mitochondria in calcium regulation of spontaneously

contracting cardiac muscle cells. Biophys. J. 75, 2004±2014.

103. Werth, J.L. & Thayer, S.A. (1994) Mitochondria buffer physio-

logical calcium loads in cultured rat dorsal root ganglion neurons.

J. Neurosci. 14, 348±356.

104. Jouaville, L.S., Ichas, F., Holmuhamedov, E.L., Camacho, P. &

Lechleiter, J.D. (1995) Synchronization of calcium waves by

mitochondrial substrates in Xenopus laevis oocytes. Nature 377,

438±441.

105. Babcock, D.F., Herrington, J., Goodwin, P.C., Park, Y.B. & Hille, B.

(1997) Mitochondrial participation in the intracellular Ca2+ network.

J. Cell Biol. 136, 833±844.

106. Di Lisa, F., Gambassi, G., Spurgeon, H. & Hansford, R.G. (1993)


Intramitochondrial free calcium in cardiac myocytes in relation to

dehydrogenase activation. Cardiovasc. Res. 27, 1840±1844.

107. McCormack, J.G., Halestrap, A.P. & Denton, R.M. (1990) Role of

calcium ions in regulation of mammalian intramitochondrial

metabolism. Physiol. Rev. 70, 391±425.

108. Hansford, R.G. (1994) Physiological role of mitochondrial Ca2+

transport. J. Bioenerg. Biomembr. 26, 495±508.

109. Robb-Gaspers, L.D., Rutter, G.A., Burnett, P., Hajnoczky, G., Denton,

R.M. & Thomas, A.P. (1998) Coupling between cytosolic and

mitochondrial calcium oscillations: role in the regulation of hepatic

metabolism. Biochim. Biophys. Acta 1366, 17±32.

110. Di Lisa, F., Fan, C.Z., Gambassi, G., Hogue, B.A., Kudryashova, I. &

Hansford, R.G. (1993) Altered pyruvate dehydrogenase control and

mitochondrial free Ca2+ in hearts of cardiomyopathic hamsters. Am.

J. Physiol. 264, H2188±H2197.

111. Bennett, W.M. & Norman, D.J. (1986) Action and toxicity of

cyclosporine. Annu. Rev. Med. 37, 215±224.

112. Petronilli, V., Cola, C., Massari, S., Colonna, R. & Bernardi, P. (1993)

Physiological effectors modify voltage sensing by the cyclosporin

A-sensitive permeability transition pore of mitochondria. J. Biol.

Chem. 268, 21939±21945.

113. Nieminen, A.L., Saylor, A.K., Tesfai, S.A., Herman, B. & Lemasters,

J.J. (1995) Contribution of the mitochondrial permeability transition

to lethal injury after exposure of hepatocytes to t-butylhydroper-

oxide. Biochem. J. 307, 99±106.

114. Szabo, I. & Zoratti, M. (1991) The giant channel of the inner

mitochondrial membrane is inhibited by cyclosporin A. J. Biol.

Chem. 266, 3376±3379.

115. Crompton, M., Costi, A. & Hayat, L. (1987) Evidence for the

presence of a reversible Ca2+-dependent pore activated by oxidative

stress in heart mitochondria. Biochem. J. 245, 915±918.

116. Crompton, M. & Costi, A. (1988) Kinetic evidence for a heart

mitochondrial pore activated by Ca2+, inorganic phosphate and

oxidative stress. A potential mechanism for mitochondrial dysfunc-

tion during cellular Ca2+ overload. Eur. J. Biochem. 178, 489±501.

117. Folbergrova, J., Li, P.A., Uchino, H., Smith, M.L. & SiesjoÈ, B.K.

(1997) Changes in the bioenergetic state of rat hippocampus during

2.5 min of ischemia, and prevention of cell damage by cyclosporin

A in hyperglycemic subjects. Exp. Brain Res. 114, 44±50.

118. Li, P.A., Uchino, H., Elmer, E. & SiesjoÈ, B.K. (1997) Amelioration

by cyclosporin A of brain damage following 5 or 10 min of

ischemia in rats subjected to preischemic hyperglycemia. Brain

Res. 753, 133±140.

119. Friberg, H., Ferrand-Drake, M., Bengtsson, F., Halestrap, A.P. &

Wieloch, T. (1998) Cyclosporin A, but not FK 506, protects

mitochondria and neurons against hypoglycemic damage and

implicates the mitochondrial permeability transition in cell death.

J. Neurosci. 18, 5151±5159.

120. Kondo, Y., Asanuma, M., Iwata, E., Kondo, F., Miyazaki, I. & Ogawa,

N. (1999) Early treatment with cyclosporin A ameliorates the

reduction of muscarinic acetylcholine receptors in gerbil hippo-

campus after transient forebrain ischemia. Neurochem. Res. 24,

9±13.

121. Reed, J.C. (1997) Double identity for proteins of the Bcl-2 family.

Nature 387, 773±776.

122. Riparbelli, M.G., Callaini, G., Tripodi, S.A., Cintorino, M., Tosi, P. &

Dallai, R. (1995) Localization of the Bcl-2 protein to the outer

mitochondrial membrane by electron microscopy. Exp. Cell Res.

221, 363±369.

123. Wang, H.G., Rapp, U.R. & Reed, J.C. (1996) Bcl-2 targets the protein

kinase Raf-1 to mitochondria. Cell 87, 629±638.

124. Susin, S.A., Zamzami, N., Castedo, M., Hirsch, T., Marchetti, P.,

Macho, A., Daugas, E., Geuskens, M. & Kroemer, G. (1996)

Bcl-2 inhibits the mitochondrial release of an apoptogenic

protease. J. Exp. Med. 184, 1331±1341.

125. Yang, J., Liu, X., Bhalla, K., Kim, C.N., Ibrado, A.M., Cai, J., Peng,

T.I., Jones, D.P. & Wang, X. (1997) Prevention of apoptosis by

Bcl-2: release of cytochrome c from mitochondria blocked. Science

275, 1129±1132.

126. Schendel, S., Montal, M. & Reed, J.C. (1998) Bcl-2 family proteins as

ion channels. Cell Death Diff. 5, 372±380.

127. Jacobs, E.E. & Sanadi, D.R. (1960) The reversible removal of

cytochrome c from mitochondria. J. Biol. Chem. 235, 531±534.

128. Bernardi, P., Pozzan, M. & Azzone, G.F. (1982) Mitochondrial

oscillation and activation of H+/cation exchange. J. Bioenerg.

Biomembr. 14, 387±403.

129. Bernardi, P., Angrilli, A., Ambrosin, V. & Azzone, G.F. (1989)

Activation of latent K+ uniport in mitochondria treated with the

ionophore A23187. J. Biol. Chem. 264, 18902±18906.

130. Adachi, S., Gottlieb, R.A. & Babior, B.M. (1998) Lack of release

of cytochrome C from mitochondria into cytosol early in the

course of Fas-mediated apoptosis of Jurkat cells. J. Biol. Chem.

273, 19892±19894.

131. Zamzami, N., Marchetti, P., Castedo, M., Hirsch, T., Susin, S.A.,

Masse, B. & Kroemer, G. (1996) Inhibitors of permeability

transition interfere with the disruption of the mitochondrial

transmembrane potential during apoptosis. FEBS Lett. 384, 53±57.

132. Zamzami, N., Susin, S.A., Marchetti, P., Hirsch, T., Gomez

Monterrey, I., Castedo, M. & Kroemer, G. (1996) Mitochondrial

control of nuclear apoptosis. J. Exp. Med. 183, 1533±1544.

133. Susin, S.A., Lorenzo, H.K., Zamzami, N., Marzo, I., Snow, B.E.,

Brothers, G.M., Mangion, J., Jacotot, E., Costantini, P., Loeffler, M.,

Larochette, N., Goodlett, D.R., Aebersold, R., Siderovski, D.P.,

Penninger, J.M. & Kroemer, G. (1999) Molecular characterization

of mitochondrial apoptosis-inducing factor. Nature 397, 441±446.

134. Mancini, M., Nicholson, D.W., Roy, S., Thornberry, N.A., Peterson,

E.P., Casciola, R.L. & Rosen, A. (1998) The caspase-3 precursor has

a cytosolic and mitochondrial distribution: implications for

apoptotic signaling. J. Cell Biol. 140, 1485±1495.

135. Krajewski, S., Krajewska, M., Ellerby, L.M., Welsh, K., Xie, Z.,

Deveraux, Q.L., Salvesen, G.S., Bredesen, D.E., Rosenthal, R.E.,

Fiskum, G. & Reed, J.C. (1999) Release of caspase-9 from

mitochondria during neuronal apoptosis and cerebral ischemia.

Proc. Natl Acad. Sci. USA 96, 5752±5757.

136. Nicholls, D.G. (1974) The influence of respiration and ATP hydrolysis

on the proton-electrochemical gradient across the inner membrane

of rat-liver mitochondria as determined by ion distribution. Eur. J.

Biochem. 50, 305±315.

137. Pietrobon, D., Azzone, G.F. & Walz, D. (1981) Effect of funiculosin

and antimycin A on the redox-driven H+-pumps in mitochondria: on

the nature of `leaks'. Eur. J. Biochem. 117, 389±394.

138. Luvisetto, S., Schmehl, I., Intravaia, E., Conti, E. & Azzone, G.F.

(1992) Mechanism of loss of thermodynamic control in mito-

chondria due to hyperthyroidism and temperature. J. Biol. Chem.

267, 15348±15355.

139. Kluck, R.M., Bossy-Wetzel, E., Green, D.R. & Newmeyer, D.D.

(1997) The release of cytochrome c from mitochondria: a primary

site for Bcl-2 regulation of apoptosis. Science 275, 1132±1136.

140. Heiskanen, K.M., Bhat, M.B., Wang, H.-W., Ma, J. & Nieminen,

A.-L. (1999) Mitochondrial depolarization accompanies cyto-

chrome c release during apoptosis in PC6 cells. J. Biol. Chem.

274, 5654±5658.

141. Li, H., Zhu, H., Xu, C.J. & Yuan, J. (1998) Cleavage of BID by

caspase 8 mediates the mitochondrial damage in the Fas pathway of

apoptosis. Cell 94, 491±501.

142. Luo, X., Budihardjo, I., Zou, H., Slaughter, C. & Wang, X. (1998)

Bid, a Bcl2 interacting protein, mediates cytochrome c release from

mitochondria in response to activation of cell surface death

receptors. Cell 94, 481±490.

143. Gross, A., Yin, X.M., Wang, K., Wei, M.C., Jockel, J., Milliman, C.,

Erdjument-Bromage, H., Tempst, P. & Korsmeyer, S.J. (1999)

Caspase cleaved BID targets mitochondria and is required for

cytochrome c release, while BCL-XL prevents this release but not

tumor necrosis factor-R1/Fas death. J. Biol. Chem. 274, 1156±1163.

144. Pastorino, J.G., Chen, S.T., Tafani, M., Snyder, J.W. & Farber, J.L.

(1998) The overexpression of Bax produces cell death upon

induction of the mitochondrial permeability transition. J. Biol.

Chem. 273, 7770±7775.


145. Bradham, C.A., Qian, T., Streetz, K., Trautwein, C., Brenner,

D.A. & Lemasters, J.J. (1998) The mitochondrial permeability

transition is required for tumor necrosis factor alpha-mediated

apoptosis and cytochrome c release. Mol. Cell. Biol. 18,

6353±6364.

146. Eskes, R., Antonsson, B., Osen-Sand, A., Montessuit, S., Richter, C.,

Sadoul, R., Mazzei, G., Nichols, A. & Martinou, J.-C. (1998)

Bax-induced cytochrome c release from mitochondria is indepen-

dent of the permeability transition pore but highly dependent on

Mg2+ ions. J. Cell Biol. 143, 217±224.

147. Novgorodov, S.A., Gudz, T.I., Milgrom, Y.M. & Brierley, G.P. (1992)

The permeability transition in heart mitochondria is regulated

synergistically by ADP and cyclosporin A. J. Biol. Chem. 267,

16274±16282.

Documents

Mitochondria and cell death : Mechanistic aspects and methodological issues