JC-1分析线粒体膜电位的方法
Andrea Cossarizza and Stefano Salvioli
Department of Biomedical Sciences
University of Modena School of Medicine
via Campi 287, 41100 Modena, Italy
phone +39 59 428.613
fax +39 59 428.623
E mail: cossariz@unimo.it
Introduction
The mitochondrial respiratory chain produces energy which is stored as an electrochemical gradient which consists of a transmembrane electrical potential, negative inside of about 180-200 mV, and a proton gradient of about 1 unit; this energy is then able to drive the synthesis of ATP, a crucial molecule for a consistent variety of intracellular processes. Several membrane permeable lipophilic cations, accumulated by living cells, organelles and liposomes exhibiting a negative interior membrane potential, have been used to study
. Such probes include those which exhibit optical and fluorescence activity after accumulation into energized systems, such as 3,3'-diehexiloxadicarbocyanine iodide [DiOC
6(3)], nonylacridine orange (NAO), safranine O, rhodamine-123 (Rh123) etc., radiolabelled probes, (
i.e., [
3H]methyltriphenyl-phosphonium, etc.) and unlabelled probes used with specific electrodes [
i.e., tetraphenyl-phosphonium ion (TPP+) etc.]. These systems have several possible disadvantages, including the: a) time required to achieve equilibrium distribution of a mitochondrial membrane probe; b) degree of passive (unspecific) binding of probes to a membrane component, such as in the case of NAO, which detects mitochondrial mass as it binds to cardiolipin (9), or Rh123, which has several energy-independent binding sites (10), or DiOC
6(3) which, notwithstanding its high capacity to bind other membranes than those of mitochondria and its low sensitivity to agents capable of depolarize such organelles (11,12), has been widely used in the last years for studies on
Dy; c) toxic effects of probes on mitochondrial functional integrity; d) sampling procedures; e) interference from light scattering changes and from absorption changes of mitochondrial components; f) requirement of large amounts of biological materials. TPP electrode affords an easy and precise tool to measure
D ydue to the:
i)low interference between bound TPP+ and the membrane; and
ii)lack of responses of the electrode to species different from TPP+. However, this method requires discrete amounts of biological samples and uptake of this lipophilic cation by intact mammalian cells is indeed a slow process.
To detect variations in D y at the single cell or at the single organelle level, a few years ago we have developed a new cytofluorimetric (FCM) technique by using the lipophilic cation 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1) (13-15). JC-1 is more advantageous over rhodamines and other carbocyanines, capable of entering selectively into mitochondria, since it changes reversibly its color from green to orange as membrane potentials increase (over values of about 80-100 mV). This property is due to the reversible formation of JC-1 aggregates upon membrane polarization that causes shifts in emitted light from 530 nm (i.e., emission of JC-1 monomeric form) to 590 nm (i.e., emission of J-aggregate) when excited at 490 nm; the color of the dye changes reversibly from green to greenish orange as the mitochondrial membrane becomes more polarized (16-18). Both colors can be detected using the filters commonly mounted in all flow cytometers, so that green emission can be analyzed in fluorescence channel 1 (FL1) and greenish orange emission in channel 2 (FL2). The main advantage of the use of JC-1 is that it can be both qualitative, considering the shift from green to orange fluorescence emission, and quantitative, considering the pure fluorescence intensity, which can be detected in both FL1 and FL2 channels.
Clearly, D y has been previously studied by flow cytometry, mostly by evaluating the changes in fluorescence intensity of cells stained with different, cationic dyes. Researchers used first Rh123 (19-21), then other molecules such as DiOC6(3) (22). Typically, the signal coming from cells whose mitochondria had a low potential was much lower than that of control samples, and in a classical histogram depolarized populations go to the left. However, after the shift to the left the peaks (i.e. that of controls and treated cells) are not always perfectly separate, the operator has to decide "by eye" where the population of cells with depolarized mitochondria begins. These two fluorescent probes have this and other problems. Rh123 binding to mitochondria is difficult to calculate when the cell has a certain mitochondrial heterogeneity due, for example, to a high number of mature or immature organelles, as occurs in a continuously growing cell line. Moreover, different mitochondrial binding sites for Rh123 exist, i.e. sites which are freely accessible whatever the energy status of the mitochondria and sites which are hidden in the energized state and freely accessible in the deenergized form of the organelle. This has been attributed to different maturative states of the organelles. Thus, in a single cell, organelles can have different Rh123 binding sites with consequent different fluorescence emissions. As a result, it is very difficult to ascertain whether or not mitochondria bind Rh123 in an energy-dependent or energy-independent manner. However, the probe is perfect when used in association with propidium iodide, as this combination allows a clear and elegant distinction between dead and living cells (4).
DiOC6(3) is more reliable for analysis of plasmamembrane potential rather than for studies on DY. Indeed, the first application of this probe in FCM was for the analysis of plasmamembrane potential (23). After this, DiOC6(3) was used in isolated mitochondria to detect D y changes (24). Any cationic molecule goes to negative sites, and can be released when the negative charge decreases. If that molecule is fluorescent, the signal decreases when the membrane potential of the organelle is lost. Fluorescent molecules present in intact cells have a different behaviour. In our hands, DiOC6(3) reacted properly when U937 cells were treated with FCCP, but such behaviour was not observed in cells treated with valinomycin. Moreover, when cells were kept in the presence of plasmamembrane depolarizing agents such as ouabain or high doses of extracellular K+, a consistent decrease in DiOC6(3) fluorescence was noted, indicating a consistent sensitivity of the probe for plasmamembrane (12). This behaviour was not totally unexpected, as it is known that this probe can bind several membranes other than mitochondria, as also reported in the Handbook of Fluorescent Probes and Research Chemicals (edited by the Company that produces and sells this reagent, i.e. Molecular Probes, Eugene, OR, USA). Thus, using this probe, it is very difficult, if not impossible, to distinguish between depolarization of plasmamembrane or changes in DY in several physiological or pathological conditions, such as apoptosis, when both events can take place.
JC-1 staining
JC-1 fluorescent probe, plastic tubes for FACS analysis, complete medium, i.e. RPMI added with 10% fetal calf serum, PBS.2.2. Methodology
1. Harvest cells (at least 2x105) from experimental samples, bring total volume up to 1 mL of fresh complete medium.
2. Stain cell suspension with 2.5 mg/mL JC-1. Shake cell suspension until the dye is well dissolved, giving a uniform red-violet color. To do this, it is also possible to vortex vigorously the suspension immediately after the addition of the probe.
3. Keep the samples in a dark place at room temperature for 15-20 minutes. The duration of the staining depends upon the cell type, but in our hands all the cells used (lymphocytes, cell lines of different origin, fibroblasts, keratinocytes, hepatocytes, etc.) responded quite well to the treatment. Wash twice centrifuging at 500 g for 5 min with a double volume of PBS.
4. Resuspend in 0.3 mL of PBS, then analyze immediatly with the flow cytometer, typically equipped with a 488 nm argon laser. Set the value of photomultiplier (PMT) detecting the signal in FL1 at about 390 V, and FL2 PMT at 320 V; FL1-FL2 compensation should be around 4.0%, while FL2-FL1 compensation around 10.6%. This is however the classical setting of the instrument we use in our laboratory, and it has to be taken into account that, as each instrument has a different sensitivity, a different setting can be necessary to obtain an optimal signal. Concerning instruments, the staining has been tested on several different apparatus such as an Excel, from Coulter (in Bergen, Norway), an Elite (Coulter) in Paris, some FACSCAN, a FACSTAR Plus and a FACSCalibur, from Becton Dickinson (in Krakow, Poland, or Modena and Venice, Italy), a Biorad Brite and a Partec (in Krakow too), and they work perfectly as well. Obviously, compensations have to be set in a different way.
