Biosci Rep (2007) 27:11–21 DOI 10.1007/s10540-007-9033-4 ORIGINAL PAPER
Evaluating Mitochondrial Membrane Potential in Cells Giancarlo Solaini Æ Gianluca Sgarbi Æ Giorgio Lenaz Æ Alessandra Baracca
Published online: 12 May 2007 The Biochemical Society 2007
Abstract Permeant cationic fluorescent probes are widely employed to monitor mitochondrial transmembrane potential and its changes. The application of such potential-dependent probes in conjunction with both fluorescence microscopy and fluorescence spectroscopy allows the monitoring of mitochondrial membrane potential in individual living cells as well as in large population of cells. These approaches to the analysis of membrane potential is of extremely high value to obtain insights into both the basic energy metabolism and its dysfunction in pathologic cells. However, the use of fluorescent molecules to probe biological phenomena must follow the awareness of some principles of fluorescence emission, quenching, and quantum yield since it is a very sensitive tool, but because of this extremely high sensitivity it is also strongly affected by the environment. In addition, the instruments used to monitor fluorescence and its changes in biological systems have also to be employed with cautions due to technical limits that may affect the signals. We have therefore undertaken to review the most currently used analytical methods, providing a summary of practical tips that should precede data acquisition and subsequent analysis. Furthermore, we discuss the application and feasibility of various techniques and discuss their respective strength and weakness.
Keywords Membrane potential Mitochondria Cells Fluorescent probes
Introduction Mitochondria are by far the main producers of ATP in eukaryotic aerobic cells. In addition mitochondria play other important roles in cell physiology and pathology,
G. Solaini (&) G. Sgarbi G. Lenaz A. Baracca Dipartimento di Biochimica, Universita` di Bologna, Via Irnerio 48, Bologna 40126, Italy e-mail:
[email protected]
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including participation in ions homeostasis, regulation of the cell redox state (including reactive radical biology), transport of metabolites, including import of proteins synthesized in the cytosol, lipid and amino acid metabolism, cell death. These important functions are highly dependent on the electrochemical transmembrane potential (proton motive force: Dp = Dw – 2.3(RT/F)DpH), a physico-chemical parameter consisting of two components: Dw, the total transmembrane electrical potential (voltage gradient), and DpH, the proton gradient that is physiologically generated across the inner mitochondrial membrane by the respiratory chain activity (Mitchell and Moyle 1969; Nicholls and Ferguson 2002). To establish the role played by Dp in pathophysiology it is extremely important to be able to measure it or at least its changes, accurately. To precisely quantify Dp in cells is very difficult and time-consuming since it requires the separate estimation of Dw and DpH. These parameters can be determined using either microelectrodes (Labajova et al. 2006) or equilibrium distributions of permeant cations and weak acids (for a recent comment, see Nicholls 2005); the latter approach being essential when only small biological samples are available. Since Dw is the major component of the proton motive force, and eventually its pH component can be experimentally further minimized by nigericin, an electroneutral K+/ H+ ionophore (Nicholls and Ferguson 2002), much research has focused on the monitoring of Dw. This parameter can be monitored using fluorescent lipophilic cations, that accumulate in the mitochondrial matrix driven by the mitochondrial membrane potential and are soluble in both the inner mitochondrial membrane and matrix space (Kinnally et al. 1978; Jackson and Nicholls 1986; Lemasters et al. 1995). This approach is rapid, it appears easy to be set, and it can supply acceptable data in cells. However, interpretation of data is not straightforward and it can be easily misleading: some notions on the fluorescence phenomenon have to be known, the method must be properly applied using the probe at very low concentrations, the probe should not interfere with mitochondrial functions, and the cells should not contain molecules or proteins capable of interfering with the dynamics or the fluorescence quantum yield of the probe (Lakowicz 1983; Montana et al. 1989). In intact cells a further complication may occur and one must be aware of: the electrical potential of the plasma membrane may change during the experiment. This parameter can be evaluated, and estimates of changes have been reported (Farkas et al. 1989; Rottenberg and Wu 1998; Nicholls 2006). The most popular fluorescent dyes used as Dw probes belong to various families of compounds, including Rhodamine dyes (Emaus et al. 1986; Baracca et al. 2003; Nicholls 2006), and Carbocyanins that include 5,5¢,6,6¢-tetrachloro-1,1¢,3,3¢-tetraethylbenzimidazlocarbocyanine iodide (JC-1) and 3,3¢-dihexyloxacarbocyanine iodide (DiOC6(3)) (Rottenberg and Wu 1998; Reers et al. 1991), merocyanines (Kalenak et al. 1991), oxonols (Cooper et al. 1990). Once the probe has been chosen, the fluorescence or its changes in normal or pathologic cells may be monitored by different techniques, including cytofluorometry, confocal microscopy, fluorescence microscopy, and fluorescence spectroscopy. In this review, we will briefly examine the techniques listed above, with a particular emphasis on fluorescence spectroscopy as based on the use of Rhodamine-123 (R-123), and suggest some practical means to approach Dw monitoring.
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Measurements of mitochondrial membrane potential Several cationic dyes distribute electrophoretically into the mitochondrial matrix in response to the electrical potential across the inner mitochondrial membrane (Chen 1988; Smith 1990; Dykens and Stout 2001; Rottenberg 1984). The accumulation takes place as a consequence of their charge and of their solubility in both the inner membrane lipids and the matrix aqueous space. For the above reasons, these dyes have been extensively employed to measure the mitochondrial electrical potential exploiting their spectroscopic properties or, alternatively, after isotopic labelling (Rottenberg 1984; Jackson and Nicholls 1986; Zanotti and Azzone 1980; Emaus et al. 1986). When properly calibrated, methods based on the use of these probes allow to estimate the size of Dw (Emaus et al. 1986; Juan et al. 1994; Nicholls and Ward 2000). Without calibration, changes in optical signal of the probe can be used to monitor the qualitative response of Dw to perturbation by external chemical or physical agents. It must be born in mind that fluorescence measurements are different when they are performed in intact cells or in both isolated mitochondria and detergent-permeabilized cells. The measurement in intact cells is based on the fluorescence enhancement due to uptake of the probe in the cell and hence in the mitochondria, whereas in isolated organelles the potential-dependent uptake induces fluorescence quenching (the probe not taken up is still present in the medium). The reason for the quenching is probably due to aggregation or stacking of the dye after accumulation (Emaus et al. 1986). Incidentally, the fact that dye accumulation induces fluorescence quenching suggests that the fluorescence enhancement observed in intact cells must be less then expected by the extent of accumulation. Fluorescence microscopy and laser-scanning confocal fluorescence microscopy These powerful tools provide suggestive images of mitochondria within a cell. They find a widespread application due to the attractiveness of the images, to the possibility of seeing even single mitochondria, their distribution and their organization as reticular networks or single organelles in a cell. This is particularly important also because it allows to evaluate the heterogeneity of the mitochondrial membrane potential within single cells (Johnson et al. 1981; Nakayama et al. 2002). In addition, these techniques allow to monitor co-localization of molecules and associated phenomena within single cells. However, this high resolution is also a flaw since the imaging field is rather limited, therefore it is difficult to obtain reliable Dw average estimates in cell populations. Moreover, microscopy techniques allow to distinguish between different states of Dw in cells, but fail to provide quantitative information due to the extreme difficulty of calibrating the method, since it is almost impossible to impose well-defined ionic concentration difference between the mitochondrial matrix and the cytosol in intact cells. Another disadvantage of the fluorescence microscope is that it cannot provide spatial resolution below the diffraction limit of specific specimen features, the detection of fluorescent molecules below such limits is readily achieved using confocal microscopy. This technique offers several advantages over conventional optical microscopy, including controllable depth of field, the elimination of image degrading out-of-focus information, and the ability to collect serial optical sections from thick specimens. The key to the confocal approach is the use of spatial filtering to eliminate out-of-focus light or flare in specimens that are thicker than the plane of focus. There has been a
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tremendous explosion in the popularity of confocal microscopy in recent years, due in part to the relative ease with which extremely high-quality images can be obtained from specimens prepared for conventional optical microscopy (Schatten and Pawley 1988; Toescu and Verkhratsky 2000). Among the most significant technical challenges for performing successful live-cell imaging experiments is to maintain the cells in a healthy state and functioning normally on the microscope stage while being illuminated in the presence of synthetic fluorophores. Quantitative three-dimensional imaging in fluorescence microscopy is often complicated by artifacts due to specimen preparation (coverslip thickness, quantum efficiency, and the specimen embedding medium), controllable and uncontrollable experimental variables (bleaching artifacts, inner filter phenomena), or configuration problems with the microscope, including laser system, optical component and alignment immersion oil (Ubl et al. 1996; Nicholls and Ward 2000). Moreover, it has to be noticed that the laser beam used to excite the probes can affect the biological system directly or by producing large amounts of radical species, therefore faithful images of functional mitochondria must be observed rapidly. Another limitation in this technique is that it currently can only be performed on intact cells. This limitation is especially significant in the study of the functioning of the mitochondria either exposed or not to xenobiotics or to conditions requiring high-energy demand (i.e. under state 3 respiration), which is probably central to the mitochondrial response, particularly when studying pathological cells. Finally, only qualitative changes in Dw of individual mitochondria in cells can be detected by this technique. The ratio of fluorescence intensity between the mitochondria and adjacent mitochondria-free cytoplasm can in theory be put into the Nerst equation to derive Dw. In practice, the resolution of the confocal microscope is insufficient to image just the mitochondrial matrix (Nicholls and Ward 2000). Applications of the described microscopy techniques in studying the mitochondrial potential of human cells harboring the mtDNA 8993T > G mutation are shown (Fig. 1). In Fig. 1A, images of permeabilized fibroblasts loaded with TMRM from a patient (bottom) and a control (top) under de-energized, state 3 and state 4 respiratory conditions are reported. In Fig. 1B confocal microscopy images of intact lymphocytes loaded with JC-1 under different energetic conditions are shown. Flow cytometry This technique offers the advantage of being able to estimate the intracellular fluorescence of cells in the culture media and to evaluate heterogeneity of a cell population due to the different levels of the mitochondrial membrane potential in single cells. Therefore, flow cytometry is conveniently used when one wishes to compare Dw in two populations of cells (Rottenberg and Wu 1998; Cossarizza et al. 1993; Juan et al. 1994; Dubot et al. 2004). A problem may arise with many types of cells: cells are required to be in suspension, and require either scraping or trypsinization, which induce oxidative stress, therefore affecting the membrane structure and function. An additional disadvantage of flow cytometry is that often the cytometer is at room temperature rather than 37C, which can give variable data depending on cells and probes used. Finally, well reproducible data are obtained particularly when analyzing intact cells, which will limit the study of functioning mitochondria under conditions of high energy demand, as discussed above for confocal microscopy. However, it has to be noticed that Plasek et al. (2005) developed a flow-cytometric method for monitoring mitochondrial
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+ FCCP
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State 4
+ FCCP
no addition
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8993T>G
B Control
8993T>G
Fig. 1 (A) It is presented a typical fluorescence microscopy image of digitonin permeabilized adherent fibroblast cells under different metabolic conditions obtained from one control and one patient harboring the 8993T > G mutation of the mitochondrial DNA (Baracca et al. 2000). Fibroblasts were incubated for 7 min with 0.2 lM TMRM, 20 lg/ml digitonin, 10 mM glucose, 5 U/ml hexokinase, 1 lg/ml rotenone, 0.4 mM ADP and 20 mM succinate (State 3 respiration) in presence of 0.4 lM oligomycin (state 4 respiration) or 2 lM FCCP (uncoupled respiration). It is shown that the mutant cells have slightly higher DW than controls during ADP phosphorylation, whereas fully uncoupled cells of both mutant and controls have similar DW, and also oligomycin-inhibited ATP synthase in mutant and control cells have the same mitochondrial membrane potential. (B) Confocal microscopy analysis of DW in living lymphocytes of one control and one 8993T > G patient. Membrane potential is monitored by means of the fluorescent probe JC-1. Lymphocytes were harvested, centrifuged 400g for 5 min and resuspended at a concentration of 105 cells/ml in RPMI 1640 supplemented with 10% heat inactivated FBS and 100 U/ ml Penicillin-100 lg/ml Streptomycin medium (complete medium). Lymphocytes were then seeded on a glass coverslip coated with polylysine for 3 h in the incubator to allow the adhesion of the cells. The samples were then incubated with 2 lM JC-1 in complete medium for 30 min, and washed twice with PBS, mounted and sealed-up on microscope slides and immediately processed. DW in presence of 2 lM FCCP, in endogenous metabolic conditions and in presence of 0.4 lM oligomycin
depolarization in permeabilized human skin fibroblasts exposed to TMRM. Interestingly, the authors could assess the differences between mitochondrial membrane potential in different populations of cells in the absolute scale of millivolts. In addition, a recent paper reports the use of real-time cytometry to analyze the membrane potential
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of isolated mitochondria and its variation in presence of exogenous effectors (Lecoeur et al. 2004), which is promising to evaluate the effects of compounds on Dw in a large number of individual mitochondria. Figure 2 reports the cytofluorometric analysis of Dw in intact lymphocytes of patients (n = 5) harboring the mtDNA 8993T > G transversion and controls (n = 12). It is shown that the mutation decreases significantly the JC-1 fluorescence-525/fluorescence575 ratio, indicating an increased Dw in patient lymphocytes. Fluorescence spectroscopy
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The use of fluorescence spectroscopy to investigate membrane potential has long being applied, dating back some three decades. The cells are suspended in a cuvette and analyzed in a relatively inexpensive fluorometer that cannot offer intracellular and/or intercellular Dw heterogeneity as more recent and high tech instruments like cytofluorometers and confocal microscopes can, but it can supply the most quantitative information on the average mitochondrial membrane potential and on its variation in large population of cells. Moreover, it allows the analysis of Dw in permeabilized cells in conditions of both low and high energy requirement as those corresponding to state 3 and state 4 respiration. In cells or isolated organelles, Dw can be monitored by two approaches called ‘‘quench mode’’ and ‘‘non-quench mode’’. The latter is not widely used since it offers a
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Fig. 2 Cytofluorometric analysis of DW in intact lymphocytes using the ratiometric fluorescent probe JC1. Lymphocytes (3 · 105 cells/ml) were loaded with 2 lM JC-1 in complete medium and incubated at 37C for 30 min followed by a sudden injection into the cytofluorometer. TOP The diagrams show the results of a typical cytofluorometric analysis (abscissa shows the ratio of fluorescence intensity FL525/ FL575 and ordinate reports the relative number of cells) of one control (a) and one T > G patient (b). BOTTOM Mean value ±SD of lymphocytes from all individuals examined (12 controls and 5 patients)
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considerable sensitivity being capable to detect a small, physiological depolarization of mitochondria (few mV) seen as an increase of fluorescence, following release of probe from the quenched matrix to the unquenched cytoplasm. However, this method can easily generate artifacts if for instance (i) the quench threshold at which the mitochondrial fluorescence ceases to be proportional to the accumulated probe is achieved or if (ii) the probe binds to endogenous compounds when internalized into the mitochondrial matrix. Therefore, the quench mode is more widely used, particularly because it allows to monitor even small changes in Dw, with a good level of reliability, as it will be shown below. Among the fluorescent molecules used as probes of Dw, Rhodamines present some advantages, including the possibility of loading cells with ester derivatives, Tetramethylor Tethraehyl-rhodamine ester (TMRM or TMRE, respectively), taken up suddenly by the cells, the relatively low sensitivity to cells environment, and a strong fluorescence quantum yield, that allows their use at very low concentrations (Ehrenberg et al. 1988). This minimizes several possible interferences and problems: low concentration reduces possible inhibition of enzymes, reduces possible effects on the membrane structure, avoids significant osmotic swelling of mitochondria due to accumulation of the probe and of its counterion. Therefore, we will describe in more details the use of Rhodamine 123 to evaluate Dw and its changes under steady state and dynamic conditions. Johnson et al. (1980) first reported Rhodamine 123 as an appropriate probe to localize mitochondria in living cells, and a few years later Emaus et al. (1986) exhaustively described the spectral and metabolic properties of Rhodamine 123 (RH123) when used as a probe of transmembrane electrical potential in isolated rat liver mitochondria. These authors first showed that RH-123 accumulates in response to Dw, and that this accumulation is followed by both RH-123 diminished fluorescence due to self-quenching and red shift. The method is based on the evaluation of RH-123 steady state fluorescence in ‘‘quench mode’’, and has proven useful in many laboratories for monitoring Dw and its changes in both organelles and cells. We also used successfully this method, but under certain circumstances we found it unsatisfactory for two strictly connected reasons: results were not sufficiently reproducible unless relatively high amounts of mitochondria were present in the samples to be examined. Since we were studying mitochondrial functions and dysfunctions in cells of patients affected by mitochondrial pathologies and the amount of tissue was very limited, we set up a more reproducible and sensitive method than that based on the evaluation of steady state Dw. This method allows good reproducible results even using as low as 70 lg protein (Baracca et al. 2003). It is based on the measurement of RH-123 fluorescence quenching kinetics and it allows to compare Dw and its changes induced by substrates and/or effectors in samples taken from patients and control individuals. Figure 3 reports traces of RH-123 fluorescence quenching in human digitonin-permebilized lymphocytes under both state 3 (+ succinate + ADP) and state 4 (+ oligomycin) respiration (panel A). Panel B shows a typical kinetic of RH-123 fluorescence quenching in lymphocytes carrying the mtDNA 8993T > G transversion, in controls (both under state 3 respiration) and in the presence of oligomycin, that blocking the proton transport through the F1F0-ATPase complex, induces iperpolarization. An exhaustive presentation of the analytical method, calibration details and discussion are reported in Baracca et al. (2003) and in Sgarbi et al. (2006). In conclusion, the two methods based on measurements of steady state and kinetics of RH-123 fluorescence quenching complement each other, being preferable the former if the availability of samples is not limited and if analysis of more than one effector on
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A
Succinate + ADP AntimycinA
Fluorescence (AU)
Oligomycin
1 min
Fluorescence (AU)
B Succinate + ADP
control 8993T>G
+ oligomycin 1 min Fig. 3 Typical example of mitochondrial membrane potential evaluation in digitonin permeabilized lymphocytes. Lymphocytes were cultured in complete medium for 24 h, then DW was assayed using the potentiometric fluorescent probe Rhodamine 123. (A) Control cells (5 · 106 cells/ml) were preincubated with an ADP regenerating system (10 mM glucose + 5 U/ml hexokinase), 33 nM cyclosporin A, 1 lg/ml rotenone, 50 nM Rhodamine 123, 20 lg/ml digitonin, 0.4 mM ADP and 20 mM succinate to induce state 3 respiration. The following addition of 0.4 lM oligomycin induced state 4 respiration. Finally, the DW was dissipated adding 1 lg/ml Antimycin A that blocked respiratory chain at complex III. (B) RH-123 fluorescence kinetics of mtDNA 8993T > G and control permeabilized lymphocytes in the above medium, either or not preincubated with 0.4 lM oligomycin. Note that the traces provide both the extent of RH-123 fluorescence quenching allowing steady state evaluation of DW and the RH-123 fluorescence quenching rate, that allows to perform the kinetic evaluation of DW following calibration of the methods, as detailed in Emaus et al. (1986) and in Baracca et al. (2003), respectively
the same sample has to be assayed, whereas the latter has to be preferred when the amount of samples is scarce and/or changes of Dw induced by single effectors have to be compared among different cell samples. As a final remark it has to be recalled that though Rhodamines are very good probes of mitochondrial membrane potential being readily sequestered by functioning mitochondria, they are subsequently washed out of the cells once Dw is even transiently lost. This characteristic suggest to perform with great caution experiments in which cells preloaded with Rhodamines must be treated with reagents that affect the inner mitochondrial membrane permeability (Bernardi et al. 1999).
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Conclusions This review presents the most widespread methods and fluorescent probes used to evaluate mitochondrial membrane potential and its changes in living or detergentpermeabilized cells, or in isolated mitochondria. It also provides some useful insights concerning their limits and their correct use particularly when they are applied to studies of biological phenomena in pathological cells, where careful preventive assay of interfering endogenous or exogenous (rescuing agents) molecules can give rise to dramatic artifacts or false results (Vergun and Reynolds 2004; O’Reilly et al. 2003). Whatever method one uses to monitor Dw, it is necessary to think carefully about how the method works, what is likely to mystify it, and eventually how quantitative it can be. The introduction of imaging techniques to the study of mitochondrial membrane potential has been a significant advance, however being aware of the limitations of the current technology is warranted (Sack 2006; Le et al. 2006; Kuznetsov et al. 2006; Jayaraman 2005).
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