ROLE, FEATURES, AND ORGANIZATION OF THE MITOCHONDRIA
Mitochondria are cell components involved in the survival of eukaryotic aerobic cells because they are the primary producers of ATP, regulators of ion homeostasis or redox state, and producers of free radicals. Mitochondria are directly involved in the transport of metabolites and proteins and in the metabolism of lipids and amino acids (1, 2). Like Janus, the mitochondrion shows two faces looking both forward and backward. It is involved in the maintenance of viability and vitality, but also plays a central role in the regulation of programmed cell death (3, 4). Thus, the mitochondrion may be considered the guardian of the gate between life and death.
The key role of mitochondria in the energy metabolism essential for the survival and proliferation of eukaryotic cells has been proven by extensive biochemical studies. Apoptosis plays a crucial role in the maintenance of homeostasis in multicellular organisms as well as in embryogenesis and metamorphosis of organs in evolution. There is an obvious link between mitochondrial dysfunction and a large number of diseases because mitochondria are present in most of the cells of an organism. Because mitochondria are ubiquitous systems, every tissue function may be affected. The most energy-dependent systems, such as the skeletal and cardiac muscles, nervous system, and renal and endocrinal systems, are frequently affected. In this context, the assessment of mitochondria or different indicators of their functionality is crucial.
Fixed cells have long been used for cytological investigations of the mitochondria; however, minimal attention has been paid to mitochondria in their native context: the living cell. Many approaches can be found in the literature, but authors typically study either mechanically isolated mitochondria or detergent-permeabilized cells. Their results and interpretations differ from studies in which mitochondria are assessed in intact cells, where the role of the cytosol as the external medium of the mitochondria must be taken into account. Indeed, intracellular organelles such as mitochondria possess function-related membrane potentials that far exceed that of the plasma membrane. The simultaneous measurement of both membrane potentials can provide important information about the role of electrical events in cell function (5). Moreover, the permeability transition pore (PTP), a dynamic multiprotein complex located at the contact site between the inner and the outer mitochondrial membranes (6), plays a crucial role in the metabolic coordination between the cytosol, the mitochondrial intermembrane space and the matrix because it participates in the regulation of matrix Ca2+, pH, mitochondrial transmembrane potential (ΔΨm), and volume (2). Conformational changes of the PTP do not provide similar consequences in isolated mitochondria or in intact cells. However, the effects of pharmacologic agents or recombinant proteins of interest are more easily studied in isolated mitochondria than in intact cells.
The development of techniques allowing high-resolution analysis of mitochondria without destroying the living cell, combined with the emergence of fluorescent probes (timeline, Fig. 1), allows the study of mitochondria and mitochondrial activities using flow and image cytometry.
The morphology of the mitochondria varies depending on cell type and activity. In most adherent cultured cell lines, mitochondria are organized in a dynamic network with an ovoid-shaped and a multi-branched structure (Fig. 2). During cell division, the organelle can switch between branched and fragmented morphologies. The cell cycle and metabolic state-dependent changes in mitochondrial morphology are controlled by a set of proteins that cause fission and fusion of the organelle mass (7, 8). Moreover, cytoskeletal changes that disrupt mitochondrial distribution may be the origin of the cell death phenomenon (Fig. 3) (9).
THE MITOCHONDRIAL TRANSMEMBRANE POTENTIAL (ΔΨm)
The proton pump of the electron transport chain (ETC) or mitochondrial respiratory chain establishes the mitochondrial membrane potential or proton motive force (Δp). The protons are pumped from the matrix space to the intermembrane space, creating a substantial pH and electrical gradient across the mitochondrial inner membrane. These protons eventually re-enter the matrix space via the ATPase, driving the synthesis of ATP. This potential is dependent on a variety of other mitochondrial functions and is involved in the apoptotic process (Fig. 4) (10, 11). A typical Δp of 180 mV comprises a membrane potential (ΔΨm) of 150 mV and a pH gradient (ΔpH) of −0.5 U (in an alkaline matrix): Δp = ΔΨm − 60ΔpH.
Different cationic fluorescent dyes can be used to measure the membrane potential of the individual cells because their essential attributes are membrane permeability and low membrane binding. Cations are attracted to the negative potential across the inner mitochondrial membrane, and thus, they preferentially accumulate into the mitochondria in living cells because they are highly efficient in crossing the hydrophobic membranes (12). A hydrophobic ionic dye will have some equilibrium distribution between the external aqueous medium, the cell plasma membrane, the cytosol and the membranes and aqueous compartments of intracellular organelles. The membrane potential plays a direct role in governing the distribution across the plasma membrane: the more negative the potential, the greater the accumulation of the positively charged dye. The membrane potential controls the average environment of the dye. The distribution of the dye is governed by the Nernst equation:
where V is the membrane potential, Z is the ion charge, R is the ideal gas constant, T is the absolute temperature, F is the Faraday constant, and Ci and Co are the concentrations inside and outside the cell, respectively (13).
A threshold concentration is recommended (characteristic of a particular probe), above which a fluorescence quenching occurs and the matrix signal becomes largely independent of any further accumulation, when cationic probes are in excess. In isolated mitochondria, the total fluorescence will decrease in response to mitochondrial hyperpolarization as a higher quantity of probe is accumulated into the matrix and quenched. In cells loaded with higher concentrations than this threshold, this phenomenon results in an artifact in the signal to ΔΨm, whereas sensitivity to ΔΨp remains the same. Therefore, it is necessary to consider the effect of probe redistribution across the plasma membrane. A temporary high fluorescence of the probe in cytoplasm, which will decay as the probe re-equilibrates across the plasma membrane, will be produced by the redistribution of the probe from the quenched mitochondrial matrix in response to rapid mitochondrial depolarization (14). Therefore, using cationic fluorescent probes requires using a third to a fourth of the initial loading concentration of the potential-indicating probe in the new buffer or medium (replaced after loading) to maintain its equilibrium distribution (15). Consequently, the ΔΨm assessment method differs between intact cells and isolated mitochondria because of the presence of the plasma membrane and its Δp. Then, the Δp can be minimized through the presence of nigericin or potassium ionophore.
Staining with ΔΨm-Insensitive Dye
Table 1 describes the features and precautions for the main ΔΨm-insensitive probes used. Some of the mitochondrial-specific dyes are considered “structural” dyes, as opposed to “functional” dyes, because they are capable of staining mitochondria regardless of their polarization status. The fluorescence of cells stained with this type of dye is directly proportional to their content of mitochondria (i.e., the mitochondrial mass). This is allowed by the affinity of these dyes with mitochondrial-specific components. For example, mitochondrial staining by Mito ID and Nonyl Acridine Orange (NAO) is attributed to their specific binding to cardiolipin in the inner mitochondrial membrane (16, 17). MitoTracker Green FM reacts with the free thiol groups of cystein residues belonging to mitochondrial proteins (18). The concentration of this dye is ∼300 times higher in the mitochondrial matrix than in the medium surrounding these organelles, which may be due to the pH of the inner mitochondrial membrane. This leads to covalent binding to a fraction of the mitochondrial proteins. The binding of this category of probe is mainly located in the inner mitochondrial membrane and suggests that the amount of fluorescence observed after cell staining is more correlated with the inner mitochondrial membrane quantity than with the full mitochondrial mass. This crucial point has to be carefully considered because the inner membrane topology may vary in different species, different cell types within the same species and different metabolic states within the same cells (19). Variations in staining intensity may sometimes be the result of a metabolic state transition and a shape modification of the inner mitochondrial membrane rather than a mitochondrial mass variation.
Table 1. Mitochondrial Specific Fluorescent Probes
Applications and/or precautions
Non-fluorescent until oxidized.
MTP insensitive probes
MitoTracker Green FM
Free thiol groups of cystein residues belonging to mitochondrial proteins; Non-fluorescent in aqueous solution, becomes fluorescent once accumulated in the lipid environment of mitochondria; immediately after the addition of the probe without a wash step; Accumulates in mitochondria regardless of MMP in certain cell types.
Mitochondrial mass, monitoring of mitochondrial morphology; more photostable than rhodamine 123.
Cell-permeant; mitochondrial selective; Thiol-reactive chloromethyl groups probably responsible for keeping the dye after fixation.
A subsequent permeabilization with cold acetone could maintain the staining pattern.
MitoTracker Deep Red 633
Mitochondrial-selective, binding to cardiolipin in the innermitochondrial membrane; minimally cytotoxic and phototoxic; superior photostability.
Mitochondrial mass, imaging of mitochondrial morphology adapted for both live and fixed cells.
Source: Enzo life sciences
MTP sensitive probes
Carbocyanine derivative, amphiphilic, accumulates on hyperpolarized membranes including ER and plasmatic membranes.
Membrane potential measurements; not specific to mitochondria; inhibits respiration and stains Golgi at high concentration. cell/dye ratio, cell size variability or magnitude of plasma membrane potential may affect the fluorescence intensity.
Green fluorescence is essentially invariant with membrane potential, whereas red fluorescence significantly increases when membranes are hyperpolarized.
Tetramethylrosamine ethyl ester (TMRE)
Lipophilic; diffuse across the cell membrane; stain specifically polarized mitochondria rather than other intracellular organelles;
Responds to the Nernst equation: reversible accumulation and quantitatively related to the contrast between intra- and extracellular fluorescences that must be taken into account during the MMP measurement in intact cells
Some of these dyes are capable of forming covalent bonds with mitochondrial proteins. This property makes them compatible with a fixation step after cell staining. This is of particular interest when biological samples exhibit an infectious risk or when samples have to be analyzed after a time delay. In these cases, the best fixation reagent is formaldehyde. For a subsequent permeabilization step, the use of acetone allows intracellular labeling with antibodies; this stage also improves the retention of the dye. However, as described by the supplier, permeabilization or fixation procedures cannot be applied to cells stained with MitoTracker Green FM.
Cell staining with ΔΨm-insensitive probes can sometimes be influenced by changes in ΔΨm. Therefore, rigorous controls have to be made for all cell lines and all protocols in order to evaluate the sensitivity of this type of dye to ΔΨm. The use of respiratory uncouplers, such as carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) or carbonyl cyanide m-chlorophenylhydrazone (CCCP) and hyperpolarization induced by nigericin, is a good way to evaluate the existence of this potential pitfall.
Staining with ΔΨm-Sensitive Dye
Table 1 describes the features and precautions for the main ΔΨm-sensitive probes used.
Most of the fluorochromes used belong to the following families of molecules:
Rosamine dyes: orange, red and infra-red-fluorescent MitoTrackers (23).
Cationic dyes are widely used as mitochondrial probes. They accumulate within the cell and preferentially localize in the mitochondrial matrix, induced by the membrane potential established by the Nernst equation that could become.
Thus, a plasma membrane potential of −60 mV and a mitochondrial potential of −180 mV will result in a tenfold accumulation of the cationic probe in the cytosol and a 10,000-fold accumulation within the mitochondria (12).
In the rhodamine family, Rh123 is widely used for mitochondrial staining because of its rapid cellular uptake and equilibration (20). However, Rh123 is not well retained by cells. Washing steps and efflux lead to a rapid decrease of cell fluorescence, and some quenching phenomena have been observed when the dye is used at high concentrations (20, 24). Rh123 was the first probe used to assess ΔΨm and it is also commonly used for the ER and Golgi apparatus. Even though Rh123 is widely used, this dye appears to assess ΔΨm incorrectly due to its relatively high non-specific binding to the mitochondria (24–26). Moreover, results are sometimes not sufficiently reproducible without using relatively high amounts of mitochondria (1, 27). The fluorescent Rh123 then accumulates in the mitochondrial membranes. Because of its non-specific binding, the use of Rh123 for determining mitochondrial membrane potential is not recommended. The same consideration applies to all fluorochromes cited in the present table under the heading: “Non-fluorescent until oxidized.” RedoxSensor Red CC-1 Stain is yet another example because it is sensitive to the superoxide anion (28).
In 1988, Ehrenberg et al. described new dyes synthesized by the esterification of tetramethylrhodamine: TMRM and TMRE. These dyes are non-toxic, highly fluorescent and neither form aggregates nor display binding-dependent changes in fluorescence efficiency (13). TMRE and TMRM can be used at relatively low concentrations for the specific staining of the mitochondria with no quenching effect (24, 29). Their low membrane partition coefficient allows them to preferentially stain specifically polarized mitochondria rather than other intracellular organelles (29). Their reversibility to the Nernst equation must be taken into account during the measurement of membrane potential in intact cells. Using flow cytometry, the analysis can be performed immediately after labeling, with or without cell wash steps. In microscopy, in time-lapse experiments following the fluorescence intensity relative to the mitochondrial membrane potential and if the extracellular medium is changed, Nernst's equilibrium must be maintained by conserving a minimal concentration of TMRM in the replacing cell medium. We estimated this concentration by spectrofluorimetry to be a third to a fourth of the initial concentration, in accordance with the literature (15)
DiOC6(3), a carbocyanine dye from the DiOC family, cannot be exclusive to the mitochondrial membrane potential measurement in intact cells, except by dissipating the plasmatic and ER membrane potentials as well as the ΔpH. DiOC6(3) is widely used as a cytofluorometric ΔΨm indicator. To produce rigorous and reproducible results, the dye and the cell concentrations have to be monitored with care. When DiOC6(3) is used at low concentrations (10–20 nM), this dye rapidly reaches equilibrium in the mitochondria with low quenching effects. The use of higher concentrations (more than 50 nM) may result in non-mitochondrial staining (plasma membrane and endoplasmic reticulum) and in fluorescence quenching. Moreover, similar to probes previously mentioned, the use of relatively high concentrations of dye leads to a high accumulation in the inner mitochondrial membrane, resulting in a quenching of the fluorescence and, occasionally, in an inhibition of either mitochondrial respiration or complex I (except for TMRM and TMRE) (30). Many factors, such as the cell/dye ratio, cell size variability or the magnitude of plasma membrane potential may affect the fluorescence intensity of cells stained with DiOC6(3) (31). If the cell population studied is homogeneous and asynchronous, this potential pitfall is not of crucial interest; otherwise, internal standards and normalization procedures have to be used to allow the detection of ΔΨm dissipation and DiOC6(3) fluorescence decrease (30).
JC-1 and JC-9 are two other members of the cyanine family and have a polychromatic fluorescence emission: a green fluorescence in a monomeric state and a red one in an aggregate state (22). At a low concentration, in normal healthy cells, JC-1 aggregates in the mitochondria in a potential-dependent manner. JC-1 undergoes aggregation within the mitochondrial matrix above a critical threshold concentration, and J-aggregates are formed when the ΔΨm is over −140 mV. The intensity of green monomer fluorescence in the matrix is potential-independent until ΔΨm falls below that value (14). In apoptotic cells exhibiting a dissipated ΔΨm, this dye does not stain mitochondria and remains located in the cytosol in a monomeric form (32). The green fluorescence of JC-9 is essentially invariant with the membrane potential, whereas red fluorescence significantly increases when membranes are hyperpolarized. The percentage of cells with a low mitochondrial membrane potential could be evaluated on a biparametric histogram with green vs. red fluorescence or using a red to green ratio as a derived parameter (33, 34). However, this method implies that short-term changes in the plasma membrane potential can be ignored: the dynamic change in the fluorescence ratio in response to a plasma-membrane depolarization will be a complex function of the rate at which J-aggregates dissociate, the rate at which monomers re-equilibrate across the inner mitochondrial membrane and any slow re-equilibration across the plasma membrane (14).
Other available dyes, such as MitoTracker CMXRos and tetramethyl rhodamine derivatives, have high sensitivity to changes in ΔΨm values, even if the changes of fluorescence emission of these dyes in response to changes in ΔΨm are quantitative (increase or decrease of a given color) and not qualitative (change in colors). Conventional fluorescent dyes are easily washed out of cells once the membrane potential is lost. MitoTracker dyes have been developed to overcome this limitation and can be used prior to a fixation step. They are cell-permeant and passively diffuse across the plasma membrane and accumulate in active mitochondria. They contain a thiol-reactive chloromethyl moiety responsible for keeping the dye associated with the mitochondria after an aldehyde-based fixation. Reduced MitoTracker CMTMRos and CMXRos (also called MitoTracker Orange and MitoTracker Red, respectively) belong to the rosamine molecule family and possess the advantage of being retained by the cell after formaldehyde fixation. This is of particular interest in the study of biohazardous samples and in the use of protocols where intracellular antigens have to be detected. Their orange or red fluorescence makes them suitable for multicolor protocols.
Quantification of Mitochondrial Potential: From Arbitrary Units of Fluorescence Intensity to Millivolts
Flow cytometry provides estimates of the magnitude of ΔΨm for a large number of individual cells, even in heterogeneous cell populations. ΔΨm determines the distribution of cations between the cytosol and the mitochondrial matrix, and ΔΨp determines the distribution between the cytosol and the extracellular medium. The accumulation of lipophilic cations from the medium to the mitochondria then depends on ΔΨm and ΔΨp. Consequently, both potentials determine the magnitude of the fluorescence at a given dye concentration, depending on cell density and the amount of mitochondrial membranes. Calibration methods with valinomycin and potassium are often used to provide accurate measurements (30, 35, 36). The addition of valinomycin hyperpolarizes the cells if the extracellular potassium concentration is higher than the intracellular one, and depolarizes the cells if inversed. The addition of valinomycin, equilibrated with a potential-sensitive dye in the media with a range of K+ concentrations, establishes a calibration curve correlating the fluorescence to ΔΨm, which can be calculated from the Nernst equation. Then, from measurements of intracellular and extracellular dye concentrations using this equation, ΔΨm can be accurately calculated. However, flow cytometry measures the amount, not the concentration, of the dye in each cell. This implies that the intracellular concentration could be obtained by dividing the fluorescence value by the cell's volume estimated from forward scatter measurements or other methods. In this case, flow cytometry does not allow the determination of the extracellular concentration (36). Confocal microscopy can be an alternative method to determine the ratio of fluorescence between internal and external mitochondrial compartments, which should be proportional to the ratio of the respective dye concentrations. The potential across the membrane separating the compartments can then be calculated by introducing this ratio into the Nernst equation (37), which was done by Lemasters et al. for the determination of the electrical potential in millivolts:
[Cation]out and [Cation]in are the monovalent cationic fluorophore concentrations in the extracellular space and within the cell, respectively (15). ΔΨ can be determined by differences of Ψ between the compartments:
ΔΨp = Ψcytosolic − Ψextracellular and ΔΨm = Ψmitochondrial − Ψcytosolic. Under normal conditions in most cells, the plasmalemmal ΔΨ ranges from −30 to −100 mV and the mitochondrial ΔΨ ranges from −120 to −160 mV. Because the ΔΨ are additive, mitochondria are more negative than extracellular space by 150 to 260 mV. This ΔΨ corresponds to the cation concentration ratios between the mitochondria and the exterior of the cell that can exceed 10,000:1. Such a variation cannot be captured in digital images using a 256 gray-level scale.
Finally, no published results have validated an absolute quantification of the mitochondrial membrane potential from the fluorescence intensities to the expressed values in millivolts, and only a semi-quantitative approach has been allowed.
Detection of Reactive Oxygen Species/Reactive Nitrogen Species (ROS/RNS) in the Cell and/or the Mitochondria
Reactive species (Table 2) are continuously generated in normal cellular processes and are essential to life. ROS and RNS play crucial roles in gene activation, cellular growth, and chemical reactions in the cell, and they are major components of the defense against bacteria and viruses. These beneficial effects could explain the failure of antioxidant therapy (38). However, when overproduced or when antioxidants become depleted, these reactive species cause oxidative stress, leading to cellular damage that can be irreversible. As the main producers of free radicals in most cells, mitochondria play a key role in several pathologies involving oxidative stress (e.g., diabetes, aging, apoptosis, cancer, and neurodegenerative diseases).
Table 2. Reactive Species Generated in the Cell
Reactive oxygen species (ROS)
Reactive nitrogen species (RNS)
Superoxide anion O2•−
Nitric oxide •NO
Hydroperoxyl radical HO2•
Nitrogen dioxide •NO2
Hydroxyl radical HO•
Peroxynitrite anion ONOO−
Peroxyl radical ROO•
Peroxynitrous acid ONOOH
Alkoxyl radical RO•
Nitrosoperoxycarbonate anion ONOOCO2−
Hydrogen peroxide H2O2
Nitronium cation NO2+
Singlet oxygen 1O2
Dinitrogen trioxide N2O3
Hypochlorous acid HOCl
ROS can be produced in the mitochondrial ETC by electron reduction of molecular oxygen by electron donors such as flavoproteins and semiubiquinone species (Fig. 5) (39). The use of inhibitors of the ETC such as rotenone, myxothiazol or antimycine A has allowed the identification of Complex I and III as sites of ROS production (40). Complex I produces superoxide exclusively into the mitochondrial matrix, whereas Complex III produces superoxide into the matrix as well as into the intermembrane space. The reactive oxygen species O2•− (superoxide), H2O2 (hydrogen peroxide) and HO• (hydroxyl radical) are the result of an incomplete oxygen reduction. H2O2 is the major radical species measured in isolated mitochondria. It is essentially produced by the dismutation of the superoxide anion by the superoxide dismutases (SOD): cytosolic CuZn SOD or mitochondrial MnSOD. H2O2 can then be converted into H2O by antioxidant enzymes such as catalase or glutathione peroxydase.
Even if mitochondria are major producers of ROS, many radicals from other origins are present in the cytosol of the cell and must be taken into account for the assessment of ROS/RNS in living cells. Figure 6 shows the different radical species and their compartmentation in the cell.
The short lifetime of reactive species and the variety of antioxidant systems existing in vivo that are able to capture these species make them difficult to detect. Fluorescent probes are excellent indicators of reactive species due to their high sensitivity (Table 3). Many precautions must be taken, and they are listed in the review of Wardman (41). The aim of this article is to present a non-exhaustive number of guidelines to provide researchers with enough knowledge about what they are measuring in their experiments, as the ideal technique has not yet been published.
Table 3. Reactive Species Specific Fluorescent Probes
Radical species detected
Precautions of use
Non-fluorescent until oxidized.
Probes for ROS/RNS detection
Amplex Red (=Resorufin)
H2O2 in presence of HRP
Matricial O2•− is converted to H2O2 and then diffuses to the intermembrane space to be detected by Amplex Red.
Preferable to use in isolated mitochondria because of the presence of xanthine/xanthine oxidase producing peroxide and NADPH interacting with HRP. Addition of SOD eliminates this reaction.
H2DCFDA; Carboxy- H2DCFDA; and CM-H2DCFDA; OxyBURST Green- H2DCFDA
H2O2, HOO−, ONOO−. Idem and HO−, O2•−
Their functional moiety for oxidation is phenolic, oxidized to a partially quinone-like structure; DCFH2 is oxidized to its radical DCFH•. DCFH2 has a very low reactivity toward superoxide radicals on the dismutation product, hydrogen peroxide.
Oxidation initiated by peroxinitrite. A decrease in binding specificity with HO• with high DCFH2 concentrations. Measuring the rate constants for the reaction of HO•, NO2• and CO3•− with DCFH2 is important to assess the reactivity of the probe toward peroxinitrite. The pH-dependance of DCF fluorescence and its cellular loss due to negative electrical charge are significant drawbacks.
DAF-FM diacetate is cell-permeant and passively diffuses across cellular membranes. Once deacetylated by esterases inside the cells, it becomes fluorescent DAF-FM.
NO is often co-localized with O2•− and ONOO−,so DAF fluorescence must be considered as an RNS indicator that is more than NO-specific (126).
Passively diffuses across cell membranes, where it is oxidized to Rh123 in mitochondria. Its positive electrical charge facilitates the intracellular accumulation due to negative cell membrane and mitochondrial potentials with reference to extracellular space.
Unreactive toward O2•− and H2O2 in the absence of catalyst. Oxidized indirectly by peroxinitrite via decomposition (radical) product. Fluorescence independant of pH.
355 cyto; 518 DNA
420 cyto*; 605 DNA
More reactive than DCFH2 or RhH2 toward O2•−, Reacts with H2O2 via peroxidase catalysts or heme proteins to produce components interfering with O2•− detection. As with MitoSox, the reactivity with O2•− is limited by the difficulty in assessing intracellular levels of the O2•− product; that is, these probes are limited despite the fact that other probes are not commercialized (127).
Uncontrolled fluorescence enhancement due to intercalation of 2-hydroethidium with mitochondrial DNA has been addressed. Reduction of cytochrome c by HE and catalysis of superoxide disproportion was reported. Unlike NO•, superoxide can accomplish the initial oxidation step. It is probable that the response of hydroethidium to superoxide can be altered by the flux of other radicals from the initial oxidation step
Live-cell permeant that is rapidly and selectively targeted to the mitochondria, where it is oxidized by superoxide. MitoSOX is not oxidized by other ROS/RNS-generating systems, and the oxidation of the probe is prevented by superoxide dismutase
The oxidation product becomes highly fluorescent upon binding to nucleic acids because MitoSOX is a derivative of ethidium bromide.
Following Wardman, a list of traps can be established:
Lack of selectivity of probes based on reduced dyes, which require a catalyst for reaction;
The absence of leakage when loading into cells;
The presence of intracellular anti-oxidants that can compete with reactive species leading to changes in fluorescence, possibly reflecting changes in competing antioxidants rather than the free radical generation rate.
Dyes that are stable in a reduced state, but that can be oxidized by the species of interest, are widely used as probes for free radical studies. The oxidation is accompanied by a change in binding to form highly fluorescent moieties. Reduced dyes are often non-fluorescent, whereas their oxidized products are highly fluorescent. The peroxidase-coupled oxidation of a probe results in the formation of a fluorescent product and defines the principle of quantitatively determining H2O2 formation in biological samples such as isolated mitochondria. The total formation of H2O2 can be accurately assessed without compartmentation effects because this ROS species most likely passes through all biological membranes by means of aquaporin channels (39).
The Amplex Red reagent (10-acetyl-3,7-dihydroxyphenoxazine) reacts stoichiometrically with H2O2 in the presence of horseradish peroxidase (HRP) to produce highly fluorescent resorufin. This probe is described and widely used to detect H2O2. When used in intact cells, Amplex Red can be combined with other ROS probes such as Carboxy-H2DCFDA.
Fluorescein probes such as (Dichloro)dihydrofluorescein H2DCFDA (or DCFH2) and its analogs are the most commonly used (41). The dye with the suffix DA (diacetate) is improved in the CM- and carboxy-forms that allow a better uptake in cells. DCFH2 is actually not a useful probe for the two most common ROS in biology such as H2O2 and O2•−, although it is still being perceived as such.
Dihydrorhodamine (RhH2) is the uncharged and nonfluorescent reduction product of Rh123. Hypochlorous acid oxidizes RhH2 to Rh123 with much higher efficiency than with DCFH2 (41).
Hydroethidine or dihydroethidine are much more reactive than DCFH2 or RhH2 toward superoxide radicals. The MitoSOX Red mitochondrial superoxide indicator is a novel fluorogenic dye for the high detection of superoxide in the mitochondria of live cells. Nitric oxide indicators (DAF-FM and DAF-2) are essentially non-fluorescent until they react with low concentrations of NO (probably NO+) to form a fluorescent benzotriazole (Table 3). DAF-2 was described by the manufacturer as the most successful indicator for nitric oxide, but less photostable than DAF-FM, whose NO adduct spectra is independent of pH above pH 5.5. These reagents are able to quantify low concentrations of nitric oxide.
To conclude this part of the assessment for reactive species, it appears necessary to establish a list of important details that must be considered before using a probe to assess the production of ROS/RNSs in a biological system:
A first point deals with the system to be considered: isolated (whether or not it is permeabilized) mitochondria, or living cells or tissues.
Peroxidase-like catalysis of DCFH2 and/or RhH2 oxidation has been reported by the most common HRP and cytochrome c, a general catalyst for the oxidation of a variety of organic molecules and nitrite by H2O2. Other catalysts reported to have peroxidase-like activities with DCFH2 include free iron, hemoglobin and myoglobin, catalase, CuZnSOD, xanthine oxidase, and lipoxygenase. Proteins such as SOD and xanthine oxydase effect their peroxydase-like activity involving CO3•− as an intermediate.
The reader should find a number of technical points exhaustively listed in the review by Wardman et al. (41), for example, probe reactivity, specificity and distribution; the presence of probe intermediates; the need for a catalyst; the effects of antioxidants; photochemical reactions or instrumental artifacts.
EXAMPLES OF APPLICATIONS
Mitochondria play a central part in the apoptotic process: they act as central regulators of the intrinsic apoptotic pathways and effector-mediators of the extrinsic apoptotic pathways (3). Under physiological conditions, mitochondria exhibit a high ΔΨm, and specific proteins are maintained within the intermembrane space. Proapoptotic signals may induce the destabilization of the mitochondrial outer membranes, the formation of pores (including PTP opening) via the direct binding of Bax, Bak and BH3-only proteins to specific components of the PTP (42, 43). This leads to the dissipation of ΔΨm and the release of intermembrane-specific proteins including cytochrome c, which activates the apoptosome and therefore the caspase cascade, and other proteins that have important proapoptotic functions (such as smac/diablo, Omi/Htra2, apoptosis-inducing factor (AIF), and endonuclease G) (4). The ΔΨm dissipation defines the early stage of apoptosis preceding DNA fragmentation, ROS production and the increase of membrane permeability (44, 45). These events [ΔΨm dissipation and mitochondrial membrane permeabilization (MMP)] occur in most cases of apoptosis. Interestingly, MMP is always followed by ΔΨm dissipation; however, ΔΨm dissipation is not always caused by MMP, and cytochrome c release has been observed even in the absence of ΔΨm dissipation (3, 46). Additionally, transient ΔΨm dissipation does not always cause cell death, and partial mitochondrial membrane permeabilization may not inevitably lead to a lethal event (47). Several methods have been developed to study apoptosis, and we describe here the techniques that focus on the mitochondrial events.
Mitochondrial Membrane Potential Dissipation
Considering the techniques that focus on mitochondrial events, the assessment of mitochondrial depolarization is the most commonly used method for the identification of apoptotic cells by cytometry (48). Early apoptotic cells are characterized by ΔΨm dissipation and the preservation of cell membrane integrity. Thus, when cells are stained with a potential-sensitive mitochondrial specific probe such as DiOC6(3), TMRE, Rh123, MitoTraker Red, or TMRM, the staining and the fluorescence intensity are directly correlated with the mitochondrial polarization status. Viable cells appear as a highly fluorescent population, whereas apoptotic cells exhibit a lower fluorescence. Usually, this mitochondrial staining is associated with the use of an exclusion dye to check the integrity of the plasma membrane. Classic distributions of viable and apoptotic cells are represented in Figure 7.
It should be noted that the mitochondrial polarization status is not the only parameter that influences the incorporation of these cationic dyes into the cell; it is also dependant on other factors such as the cell size, the cell shape, the mitochondrial density and structure (ovoid shaped or multi-branched), the activity of the multiple drug resistance pump, and the plasma membrane permeability.
The JC-1 probe differs from other molecules because of its multi-spectral characteristics (see paragraph entitled “Staining with ΔΨm sensitive dye”). It has been used for the detection of cells exhibiting mitochondrial alterations during apoptosis by flow and imaging cytometry (49–51). The aggregation/monomerization phenomenon is due to variations in ΔΨm and leads to a lower plasma membrane polarization dependence of this dye than with DiOC6(3) (31). The red-to-green fluorescence ratio allows a normalization of the signal that avoids the influence of mitochondrial size, shape and density on the signal.
Moreover, we recommend the use of CCCP-treated cells as a depolarization control. The fluorescence intensities of apoptotic cells and CCCP-treated cells need to be identical.
Reduced ΔΨm is not an absolute indicator of apoptosis. It may be the result of an inhibition of the mitochondrial respiration that could occur during quiescence or following cellular stress, leading to a transient mitochondrial membrane permeabilization. The presence of apoptotic cells needs to be confirmed by a more specific technique such as the detection of activated caspases.
Cytochrome c Release
As a result of mitochondrial membrane permeabilization, cytochrome-c, a key component of the respiratory chain located in the intermembrane space, is released from the mitochondria to the cytosol (52). Released cytochrome c acts as a caspase activator and may serve as a target for the identification of apoptotic cells. Fluorescence microscopy of immunolabeled cells has been the first cytometric technique developed for the identification of apoptotic cells on the basis of cytochrome c release. The release of this mitochondrial component was highlighted by the staining pattern: viable cells were characterized by a punctiform staining corresponding to the mitochondrial localization of cytochrome c, and apoptotic cells exhibited a more diffuse staining due to the release of cytochrome c from the mitochondria to the cytosol. Dual staining may offer an accurate identification of apoptotic cells and may provide additional information about the apoptotic process. For example, counterstaining with a nuclear probe and/or antibodies specific to apoptotic actors such as active caspases allows the study of interconnected events (e.g., changes in the nuclear morphology and/or cascade of activation of caspases) (48). However, a co-localization study with a dual staining of cytochrome c and another protein that always remains within the mitochondria (also termed “mitochondrial sessile protein”; e.g., voltage-dependent anion selective channel) allows a precise monitoring of cytochrome c release (4). These techniques require fixation and permeabilization steps. To visualize the release of cytochrome c in living cells, they can be transfected with a cytochrome c—GFP fusion protein and can be observed by video-microscopy (53). This allows a kinetic study of the apoptotic process and gives information that is not generally obtained with a unique immuno-staining.
However, microscopic studies prove rather useless in the analysis of a large number of cells. In this case, flow cytometry will be preferred. However, the use of this technique is limited because of the necessity of distinguishing between the mitochondrial and the cytosolic localization of cytochrome c. Two different approaches help to solve this problem. The first method is based on the specificity of different cytochrome c-specific monoclonal antibodies: one of the antibodies (clone 7H8.2C12) cannot detect cytosolic cytochrome c after the induction of apoptosis and can only detect mitochondrial cytochrome c (54). Staining apoptotic cells with this antibody results in a decrease in the quantity of bound antibodies within the cell compared to the staining of viable cells (55). Another clone can be used as a control to detect the total amount of cytochrome c. The second method is based on a permeabilization method that causes all of the cytosolic contents to wash out from the cell with the exception of mitochondrial cytochrome c. In this method, the permeabilizing reagent digitonin is used at low concentration. After fixation, permeabilization and staining, the intact cells appear as the brightest cell population, and cells that have released cytochrome c exhibit a diminished fluorescence (56). These methods are off the beaten track and not ready for widespread use. A rigorous determination of the methodology needs to be done for every type of cell and apoptogenic stimulus.
Visualization of PTP Opening by Confocal Microscopy
Mitochondrial permeability maintains the rate of ATP synthesis in living cells. Under certain conditions, the opening of the mitochondrial permeability transition pore (PTP) causes the inner membrane to be permeable to molecules up to 1,500 Da. The opening of the PTP results in a rapid collapse of the membrane potential, an equilibration of ion gradients and a loss of metabolites (57). Consequently, mitochondrial transition permeability constitutes a fundamental step in the signaling cascade leading to apoptosis. Matrix calcium is the most important factor regulating the mitochondrial transition permeability, whereas cyclosporine A has been identified as a potent inhibitor of the pore transition.
To monitor the mitochondrial inner membrane permeabilization and/or the transient opening of the PTP, the calcein-cobalt technique can be used. The mitochondrial inner membrane is characterized by its quasi-impermeability to water and ions, including protons. However, the calcein acetoxymethyl ester, because of its high liposolubility, can diffuse into all subcellular components including mitochondria. Cells are then loaded with both calcein acetoxymethyl ester and its quencher cobalt (Co2+) that distributes in cells but not in the mitochondria (58). Viable cells stained and analyzed by microscopy appear with a punctiform fluorescence located within the cytosol as a result of the unquenched fluorescence of calcein that has diffused into the mitochondria. The mitochondrial inner membrane permeabilization or the transient opening of the PTP allow the uptake of Co2+ within the mitochondria, leading to the quenching of calcein fluorescence and a decompartmentation (Fig. 8). This technique can also be used with flow cytometry where cells with a permeabilized inner membrane exhibit lower fluorescence intensity than intact cells (48).
Colocalization of TMRM and NAD(P)H Fluorescence
A novel method to assess the opening of the PTP by confocal microscopy consists of a time-lapse measurement followed by NAD(P)H autofluorescence and TMRM. Based on the method described by Dumas et al., the monitoring of both intracellular NAD(P)H distribution and mitochondrial potential intensity indicates the PTP state (59). Under normal conditions of viable cells with closed PTP, the autofluorescence of NAD(P)H is mainly intra-mitochondrial and co-localized with TMRM fluorescence. A transient opening of the PTP induces NAD(P)H diffusion out of the mitochondria that can be visualized as an increase in the area and fluorescence intensity of NAD(P)H (Fig. 9). In the case of a transient PTP opening, the mitochondrial potential-dependent TMRM signal remains stable. Only an irreversible mitochondrial depolarization, considered as the point-of-no-return of apoptosis, leads to a decrease in TMRM fluorescence.
Proteins Involved in the PTP Opening
Bax and Bak have been described as the main effector proteins implicated in the PTP opening but a large number of regulatory protein, including the BH3-only proteins family members, also have a crucial role in the mitochondrial apoptosis regulation (42, 60). Several techniques were developed in order to assess these proteins expression including cytometry. A very complete review has recently been published on this subject (43). Here we just bring to mind that the apoptotic regulatory processes imply homo- and hetero-dimerizations of the Bcl-2 family members. Detection of a single kind of protein is also poor-informative compared to protein–protein interaction studies allowed bay the use of fluorescence resonance energy transfer-based cytometry.
Recently, Kroemer et al. provided recommendations to authors, reviewers and editors of scientific journals for the abandonment of confusing expressions that are frequently used. Instead of terms such as “percent apoptosis,” “percent necrosis,” “percent cell death,” or “percent cell survival,” they recommended the use of more descriptive, more precise and less confusing terms such as the “percent of cells with a low membrane potential” and the “percent of cells exhibiting cytochrome c release” (61).
Using targeted fluorescent probes, the analysis of the mitochondrial compartment can reveal modulations in mitochondrial mass, regardless of whether they are associated with distinctive ultrastructural and/or functional changes. There are several situations (i.e., cell types, experimental conditions, etc.) in which mitochondrial mass evolve, and only some examples are provided in this part of the review.
A prerequisite for the evaluation of mitochondrial mass is that the dye used here is insensitive to ΔΨm: the cell fluorescence intensity after staining should not be linked to the polarization status of the mitochondria. However, some authors have described that the mitochondrial membrane staining with NAO, MitoFluor Green, and MitoTracker Green, may respond to ΔΨm variations (62–64). In this case, these probes are not reliable indicators of the mitochondrial mass and it is important to take this point in account. As mentioned in the paragraph about non-potentiometric probes, the insensitivity of the dye to ΔΨm must be checked using uncouplers or nigericin.
NAO is one of the most commonly used dyes for the monitoring of mitochondrial mass. It has been used concomitantly with Rh123 or a MitoTracker for the study of the variations of mitochondrial mass and/or mitochondrial function during the cell cycle. For example, the induction of quiescence in monolayers or spheroids induced an enhancement of mitochondrial mass per unit of cell volume for all cells while the mitochondrial function (activity/mass) decreased with quiescence (65). Another study by Sweet and Sing et al. dealt with changes in mitochondrial mass during the cell cycle: ATP changes as a function of cell cycle progression but cannot be predicted by changes in mitochondrial mass or membrane potential (66).
Using MitoTracker Green FM for mitochondrial mass estimation and Propidium Iodide for DNA content measurement, Arakaki et al. suggested that protein factors modulating mitochondrial morphology, size, distribution, and mass are involved in mitochondrial functions related to metabolism, differentiation and cycle progression (67).
Because an unblemished solution allowing the mitochondrial mass evaluation does not exist, it is important to keep in mind the limits of all these methods.
Mitochondria and Cell Proliferation
Mechanisms that control the mitochondria during progression through the cell cycle of mammalian cells remain poorly known (68, 69). The first study was undertaken by Darzynkiewicz et al. on cultured lymphocytes stimulated with phytohemagglutinin. Flow cytometric analysis revealed a fluorescence increase during stimulation with a maximal fluorochrome uptake, concomitant with the peak of DNA synthesis, mitotic activity and increase in cellular RNA content (70). To determine the pattern of mitochondrial increase per cell over the cell cycle, James and Bohamn established the mitochondrial specificity and stoichiometry of Rh123 (71). They showed a near-linear synthesis of mitochondrial mass over the cell cycle and culture age in HL60 cells. Benel et al. compared Rh123 and NAO for their respective capabilities in distinguishing between the activity and biogenesis of mitochondria in non-proliferating vs. exponentially proliferating cells. The authors suggest that the different evolution of Rh123 and NAO uptake could be due to the existence of two levels of controls for mitochondrial activity: an enhanced respiratory activity to provide the energy required for growth, and a mitochondrial biogenesis (72). Using TMRM to assess membrane potential and PI for cell cycle analysis, Schieke et al. demonstrated that cell cycle progression through G1 is associated with a significant increase in mitochondrial membrane potential (ΔΨm) and respiration (73). They suggested a coupling of mitochondrial bioenergetics and G1 progression and described the mTOR signaling pathway as a potential molecular coordinator of these two processes.
Kennady et al. studied the relationship between mitochondrial size and cell cycle phases using flow cytometry and image cytometry for rat fibroblast synchronization and labeling with Hoechst 33342 and Rh123 (74). Based on the fluorescence pulse width analysis, two populations of mitochondria were detected in viable cells: early G1 phase cells with the smallest mitochondria and mitotic phase cells with the largest mitochondria, suggesting that mitochondrial size increased during cell cycle progression. Another study by Yang et al. revealed two distinct Rh123-stained populations of mitochondria isolated from maize leaves. Further labeling of mitochondria stained with Janus Green B and Rh123 followed by microscopic observation revealed different sizes of mitochondria as determined by pulsed-field gel electrophoresis from the mitochondrial DNA. These results suggested that several types of mitochondria coexist with different physiological statuses, mass, genomic DNA sizes, and functions during the growth and development steps of the maize leaf (75). Mitra et al. have studied the potential regulatory role of mitochondria in cell cycle control by cyclin E or other cyclins (76). They examined the mitochondrial membrane potential (assessed by TMRE) per unit of mitochondrial mass (using MitoTracker Green) at different cell cycle stages. They demonstrated that (i) when mitochondrial potential is monitored in cells at different cell cycle stages, the membrane potential is greatest at G1-S; (ii) a giant mitochondrial network occurs during the G1-to-S transition in mito-NRKcells, from normal rat kidney, stably expressing red fluorescent protein (RFP) targeted to the mitochondrial matrix; (iii) a mitochondrial depolarization specifically blocks the G1-to-S cell cycle progression in a p53-dependent manner; (iv) the presence of hyperfused mitochondria induces cyclin E buildup; and (v) hyperfused mitochondria induces G0 cells to enter the S phase.
ROS and RNS produced by the mitochondrial ETC are involved in a variety of cellular processes for the continuous adaptation of the cell to its environment. Described as deleterious processes in most cases, the oxidative stress phenomenon is essential to the existence and development of the cell. Measurement of ROS production constitutes a fundamental aspect to evaluate the cell state exposed to different stress situations. However, for in vivo experiments, the presence of intracellular antioxidants must be considered for the interpretation of the results. Indeed, antioxidants are often at the location of radical formation to compete with the target of the radicals, which are extremely short-lived species (38). Batandier et al. studied methodological aspects of the determination of mitochondrial ROS. They presented variances or interferences and limits of ROS measurement methods in order to (i) avoid artifactual conclusions and (ii) improve researchers' knowledge of the physiological and pathological mechanisms of such phenomenon (77). Kohen and Nyska reviewed the oxidation of biological systems and provided different topics that are essential for the understanding of oxidative stress as well as methodological tools for its quantification (38). These aspects of ROS production must be clearly understood before undertaking ROS/RNS quantification in cellular systems. Tarpey proposed methods for the detection of reactive metabolites of oxygen and nitrogen (78). Soh presented advances in fluorescent probes for the detection of ROS and mentioned Fluorescence resonance energy transfer (FRET) as an exploitable approach in cellular ROS monitoring (79). This work completes the review by Gomes, who presented fluorescent probes as excellent sensors of ROS (80). The first part of the present article described these different approaches and fluorescent probes for the quantification of oxidative stress.
Mitochondria are highly dynamic systems capable of fusion and fragmentation that can occur concurrently. These phenomena regularly alternate in the cell cycle (81). Mitochondrial fusion remains a largely unknown process. Legros et al. used a green and a red fluorescent protein targeted to the mitochondrial matrix to demonstrate that mitochondrial fusion in human cells is efficient and achieves complete mixing of matrix contents within 12 h. They showed that fusion requires mitochondrial potential and is mediated by mitofusins (82). Mitochondrial fusion represents a rescue mechanism for impaired mitochondria by the mixing of contents (proteins, lipids and mitochondrial DNA) and the unification of the mitochondrial compartment, permitting it to maintain its functionality and play roles in cellular development, aging and energy dissipation (83, 84). Fluorescent methods are described to assess mitochondrial fission and fusion in cells; however, none of the fluorescent probes described above is mentioned for this application. Most methods cited in the literature present other fluorescent technologies based on fluorescent proteins targeted to the mitochondrial compartment. Inspired by cell-cell fusion strategies employed in yeast, mitoRFP (mitochondrially targeted RFP) or mitoGFP (mitochondrially targeted GFP) has been applied to monitor the mixing of mammalian mitochondrial matrices and to demonstrate that fusion is dependent on mitochondrial membrane potential but not on mitochondrial respiration (82, 85). New technologies were developed to mimic in vivo conditions: Fluorescence recovery after photobleaching (FRAP) makes it possible to follow the refilling of bleached areas with mitochondrially targeted fluorescent molecules from neighboring unbleached (fluorescent) areas to demonstrate that the mitochondria are physically connected (86). The photoactivable mitoGFP (variant of aequora Victoria GFP, in which a single amino-acid substitution renders a stable 100-fold increase in green fluorescence after laser excitation) facilitates the ability to monitor the fission and fusion of individual mitochondria. The photoconvertible fluorescent protein Kaede was also used to determine whether mitochondrial fusion occurs in plants via the conversion of green to red fluorescence of labeled mitochondria within a cell (87).
Aged and injured mitochondria may enter an autophagic process called “mitophagy” that participates in the mitochondrial homeostasis and the recycling of some mitochondrial components such as amino acids and nutriments (88). Briefly, this very rapid process (less than 10 min) consists of the sequestration of the mitochondria by isolation membranes to form an autophagosome. After the dissipation of ΔΨm, autophagosomes acidify and fuse with lysosomal vesicles to form autolysosomes. Then, the mitochondria are entirely hydrolyzed, and their molecular components are recycled. The sequestration of the mitochondrial content is also important to avoid the release of proapoptotic proteins that may activate the cell death pathway. Mitophagy typically occurs when cells are in conditions of nutrient deprivation and is inhibited after nutrient replenishment [for review, see (89–92)].
Studies of mitophagy by flow cytometry appear to be very complex because of the low number of mitochondria simultaneously located within the cell chondriome. However, microscopic techniques are well suited to reveal the physiological changes of a single organelle diluted in a pool of mitochondria. To detect autophagic mitochondria, cells can be co-stained with MitoTraker Green to localize all the mitochondria, LysoTracker Red for the labeling of acidic intracellular compartments and TMRM to monitor the ΔΨm (93). The co-localization of MitoTraker Green and TMRM in polarized mitochondria allows a fluorescence resonance energy transfer between these two dyes that give rise to red fluorescence emission with blue light excitation. In the case of mitochondrial depolarization, the TMRM is released from individual mitochondria. Because of its covalent binding to protein thiols after uptake, MitoTraker Green is retained in the depolarized mitochondria, and its fluorescence emission becomes unquenched. A chromatic change from a red to a green fluorescence demonstrates the appearance of a spontaneous mitochondrial depolarization resulting from an autophagic process. A more specific detection of mitophagy can be assessed with the addition of LysoTracker Red (93). This dye stains lysosomes, endosomes and autophagosomes because of its acidotropic properties. Elmore et al. used this strategy to identify different stages in the mitophagic process including (i) normal mitochondria with quenched MitoTraker Green fluorescence; (ii) a population of depolarized mitochondria with unquenched MitoTraker Green fluorescence and a low red fluorescence, identifying autophagic mitochondria that are entrapped inside an acidic autophagic vacuole; (iii) a population of depolarized mitochondria with unquenched MitoTraker Green fluorescence and a bright red fluorescence underlining lysosomal/autophagosomal vacuole formation surrounding the mitochondria (93). A dual staining with MitoTraker Green and LysoTracker Red is easier to set up and allows sufficient detection of mitophagy (94).
Monodansylcadaverine can also be used to stain autophagosomes. Its quantum yield is significantly enhanced when hydrophobic interactions with lipids are formed from unhydrolyzed mitochondrial lipid membranes contained in autophagosomes (93, 95). Monodansylcadaverine is excitable by UV light and emits a blue fluorescence.
Resting or quiescent cells are characterized by a non-proliferative status and a low metabolic activity. This particular state of the cell cycle is accompanied by a low rate of transcription (decrease in the content of mRNA and transcription-associated proteins), a low rate of protein synthesis, a low rate of metabolism (decrease in the mitochondrial mass and/or decrease in the mitochondrial polarization level) and a reduction in cell size. Many cell types are concerned by cellular quiescence, and their entry in this state is fundamental for the control of their physiological properties. Quiescence might also be fundamental for the preservation of cell integrity by improving cell resistance to oxidative stress, DNA damage and stress response proteins (96–98). The quiescence of hematopoietic stem cells is also crucial for the preservation of their stemness (i.e., repression of differentiation, self-renewal, and tissue repopulation) (99–101).
The detection of quiescent cells by flow cytometry is also of particular interest, and methodologies using mitochondrial staining have been developed for many years to identify this subpopulation. Darzynkiewicz et al. conducted the first studies on resting lymphocytes after cell staining with Rh123, where resting cells were characterized by their low uptake of the vital dye (70). In most cases, this low uptake may be due to a diminished mitochondrial mass and/or a lower mitochondrial polarization in quiescent cells than in proliferating cells. The same staining strategy has been used for the detection and isolation of hematopoietic stem cells in multiple species (102–105). Currently, the identification of hematopoietic stem cells following Rh123 staining appears to be the main application of quiescence studies using mitochondrial-specific probes. However, this technique may provide a valuable method for the characterization of other stem cells such as cancer stem cells.
Mitochondria, pH, and Other Ions
The regulation of protons and other ion concentrations is crucial for mitochondrial functionality. Different methods were developed to evaluate the concentration of ions inside the mitochondria. Takahashi et al. described a method allowing the measurement of intramitochondrial pH with SNARF-1 pH indicator (excitation max 576 nm, emission max 635 nm, no fixable) (106). After staining with 10 μM of the dye, highly selective localization of the staining was obtained with a prolonged period of incubation without any dye, leading to a passive diffusion of the cytosolic SNARF-1 out of the cell and the conservation of the dye located inside the mitochondria. SNARF-1 fluorescence emission, analyzed by confocal microscopy, only responds to variations in mitochondrial pH. The same authors have described an alternative method that uses a mitochondrial-targeted yellow fluorescent protein. This system combines the advantages of specific mitochondrial localization, high-fluorophore quantum yield, and an extinction coefficient with an appropriate pKa for measuring mitochondrial pH (106).
The CoroNa Red Na+ indicator is an analog of the Ca2+ chelator BAPTA and is weakly fluorescent in the absence of Na+. Its fluorescence (excitation max 547 nm, emission max 570 nm, no fixable) increases (∼15 fold) upon binding to Na+. Colocalization studies using MitoTracker Green FM have shown that CoroNa Red spontaneously localizes in the mitochondria. This probe has been used for the monitoring of Na+ transients in the mitochondria of astrocytes and MDCK cells (107–109).
Some ions, particularly calcium, have strong effects on the regulation of enzymatic activities or metabolic systems, including mitochondrial Krebs cycle dehydrogenases in many cell types. The presence of calcium-transporters in the mitochondria allows these organelles to control the calcium cycle in the cell. Indeed, an excess of calcium in the mitochondria causes the opening of the mitochondrial permeability transition pore, eventually leading to cell death. Fluorescent approaches to evaluate the rate of calcium influx or efflux in mitochondria can be coupled with other methods to correlate mitochondrial activity and calcium effects.
Zoccarato et al. used Fura-2 and Calcium Green 5N to demonstrate that Ca2+ acts as an inhibitor of both H2O2 removal and succinate-supported H2O2 production. The use of high and low affinity Ca2+ indicators Fura-2 and Calcium Green 5N, respectively, can give an indication of the magnitude of free Ca2+ concentration in the incubation medium (110).
Rhod-2 is a non-fluorescent probe for Ca2+ binding and becomes fluorescent (excitation 556 nm and emission 576 nm) with increasing Ca2+ concentration. Using confocal microscopy, Anmann et al. showed a significant increase in the fluorescence intensity of Rhod-2 in the mitochondrial matrix in the presence of elevated concentrations of calcium. The mitochondrial localization was imaged from the autofluorescence of mitochondrial flavoproteins in cardiomyocytes (111). The main difficulty in measuring mitochondrial ion concentration is the lack of specificity of the utilized dyes in terms of localization. Lysosomal staining was observed in some cases (112). More recently, fluorescent fusion proteins were developed for the estimation of calcium and zinc concentrations in the mitochondria (113, 114). This new strategy offers a valuable method for the specific assessment of ionic dynamic changes.
Using Mitochondrial Probes for Multidrug Resistance Assessment
Drug resistance is one of the major obstacles to the successful treatment of cancer by chemotherapy, and one way for tumoral cells to survive antitumor drug administration is by the MDR phenotype, which can be the result of a variety of mechanisms (115). However, the main mechanism is the overexpression of drug-transmembrane transporter proteins (116). The best-known involved proteins are the P-glycoprotein (P-gp) (117), encoded by the mdr1 gene (118); the multidrug resistance-associated protein MRP-1 (119), encoded by the mrp1 gene; and the breast cancer resistance protein BCRP (120). They belong to the ABC super family of transporters and use ATP hydrolysis to extrude various drugs from cancer cells (121)
The activity of P-gp tends to lower the intracellular concentration of its substrates. The efflux of fluorescent probes is currently used to evaluate P-gp activity in various cell types using fluorochromes/P-gp substrates such as NAO, Dimethyloxadicarbocyanine iodide (DiOC2), and Rh 123, allowing an assessment of P-gp activity (122–124). Using flow cytometry, Marques-Santos described the use of MitoTracker Green FM for the assessment of the mitochondrial mass of a human erythroleukemic cell line and its counterpart overexpressing P-gp. Special attention must be paid to the mitochondrial properties of chemoresistant cells when fluorescent dyes are involved. MitoTracker Green, a probe used to evaluate mitochondrial mass, is a P-gp substrate, and its staining profile is dependent on the activity of this protein. Many factors, such as P-gp expression or activity, alterations in mitochondrial lipid and protein content, mitochondrial ABC protein expression or MDR phenotype, can lead to a misinterpretation of the data. The author suggests that attention should be given to the expression of P-gp when performing an evaluation of mitochondrial properties with fluorochromes specific to mitochondria.
Different methods using fluorescent probes have been described to assess mitochondrial compartments in living or detergent-permeablized cells or isolated mitochondria (1). Many errors can occur regardless of which fluorescent probe is used, including the interference with cell or mitochondrial metabolism, photo-induced damages, and probe binding (14, 125).
The different morphological states of heterogeneous organelles such as mitochondria reflect their different activities. The term “mitochondria(l) activity” is used to characterize respiring mitochondria that have a functional, active ETC to provide ATP. These organelles can function differently depending on their position and whether they are located near structures where energy is required (125).
As for the machinery of ETC, it must be noted that the ΔΨm varies depending on the function of the cell metabolism and the function of the given substrates provided to the ETC. For example, the mitochondrial membrane potential increases in the presence of fatty acids compared to a carbon-hydrate diet (unpublished results).
During apoptosis or in response to depolarizing agents, some mitochondria lose their membrane potential whereas others do not, even in the same cell from the same culture.
Staining with mitochondrial-specific probes induces an inhibition of mitochondrial respiration. This inhibition is dependent on the probe and its concentration. No inhibitory effects were observed with TMRM, whereas the strongest effects were observed with the DiOC6(3). Rh123 and TMRE exert intermediate inhibitory effects.
It is important to use low amounts of fluorescent probes to avoid the problems related to their toxicity, as they are capable of affecting several mitochondrial functions (e.g., uncoupling action, inhibition of ETC, and mitochondrial respiration).
For these reasons, the morphological and functional heterogeneity of mitochondria must be taken into account in the interpretation of data.
For this reason, cytometric techniques such as flow cytometry and confocal microscopy offer several advantages. Flow cytometry evaluates only the fluorescence associated with single particles using low amounts of biological samples and dye, thus avoiding quenching and light-scattering variations, molecules that are not incorporated into cells or organelles that are not analyzed. Flow cytometry allows the quantification of morphological and functional characteristics of whole cells or organelles. An additional advantage of some flow cytometers is the possibility to sort highly purified cell populations, in which mitochondria can be studied.
Despite the fact that flow cytometry requires cell suspension (trypsinization must be performed before incubation with the fluorescent probe because of the changes induced on the membrane structure and function that can affect the membrane potential), this reproducible method allows the detection of different levels of mitochondrial ΔΨm in a heterogeneous population of a large number of cells. Fluorescent and confocal microscopy provides precise images of the mitochondria within the cell (can visualize even a single mitochondrion), and allows the visualization of their distribution and organization as reticular networks in a majority of cell types (30). However, confocal microscopy allows measurements of only a few cells at a precise time. Therefore, it does not seem fit for the analysis of a large population of cells and presents limitations when studying a homogeneous cell population.
In addition to the numerous fluorescent probes, diverse sets of kits have been developed by several companies to probe mitochondria. Kits allow the measurement of ΔΨm as well as the opening of the permeability transition pore. They offer a fast, simple, and convenient alternative for labeling and assaying mitochondria, they contain all the required reagents and they are convenient for fluorescence observation by fluorescence microscopy or measurement by flow cytometry. In most cases, protocols do not require tricks to work for adherent cells and cells in suspension; however, kits are often tested on a limited number of cells. They are characterized by a short sampling time but remain expensive as compared with “homemade” reagents, and it is often necessary to verify their innocuousness on each cellular type. Even if kits provide the key advantage of containing all of the materials required for the experiments, one must keep in mind that “if you want something done right, do it yourself!”
In conclusion, the cytometric assessment of mitochondria using fluorescent probes requires the consideration of
the sampling procedures;
the incubation time and conditions to achieve equilibrium of the mitochondrial membrane probe;
the degree of non-specific binding of the probe in a non-ΔΨm-dependent manner;
the effect of the probe on mitochondrial functional integrity;
interference from light-scattering changes and from absorption changes of mitochondrial components.
“The ideal chemical probe […] would be highly reactive at low concentrations: specific, sensitive, without other reactivity, non toxic, well characterized chemically, easy to load into organelles, cells or tissues without subsequent leakage or unwanted diffusion, excretion or metabolism, readily available, easy to use without specialized apparatuses and cheap” (41).
This article reviewed some fluorescent probes used to assess the mitochondrial compartment by measuring its activity. However, questions regarding their specificity and sensitivity in biological samples must be kept in mind. Electrochemical electrodes were recently developed to monitor NO production (e.g., [126)]; however, the results appeared difficult to reproduce.
The authors thank Dr. W. Patton from Enzo life sciences for information regarding the Mito-D; Prof. Valdur Saks, Dr. Frederic Lamarche, and Dr. Sandrine Lablanche for the images of cardiomyocytes, astrocytes, and INS-1 cells, respectively; Prof. Eric Fontaine, Dr. Cécile Batandier and Dr. Serge Bottari for their scientific advice as well as Christophe Cottet for reviewing the English version of this article.