2. Choosing a method and a dye to measure Δψm
The next considerations are (i) in which mode to use the dye, (ii) which dye to use, and (iii) what read-out method to use? As we discuss here, the best usage mode will be determined by the experimental requirements, which in turn will help determine the optimal dye for the studies, both of which in turn will help dictate the most suitable read-out for monitoring the fluorescent signal.
Typically, these dyes can be used in either quenching or nonquenching modes, as described in considerable detail elsewhere (2
). Briefly, quenching mode utilizes higher dye concentrations (50–100 nM to several micromolar, determined empirically), such that dye accumulates within mitochondria in sufficient concentration to form aggregates, thus quenching some of the fluorescent emissions of the aggregated dye. Under these conditions, once dye is loaded into mitochondria, a subsequent mitochondrial depolarization will result in release
of dye, thus unquenching the loaded probe, and (transiently) increasing
fluorescent signal. Once all probe in the mitochondria is unquenched, fluorescent signal will decline as dye continues to leave the mitochondria and the cell under continued depolarization. Conversely, a subsequent mitochondrial hyperpolarization will result in more dye entering mitochondria, causing further quenching and thus a relative decrease
in fluorescent signal. As such, and because quenching is a nonlinear event (15
), quenching mode can only be appropriately used for monitoring the dynamic or acute effects of experimental treatments on Δψm after
the dye has been loaded (i.e., Scenario 2 experiments). Furthermore, quenching mode is only appropriate for Scenario 2 studies when rapid and robust changes in Δψm
occur during the imaging period (2
). For following slower Δψm
changes in real time (slower Scenario 2 experiments), or for determining differences in Δψm
among two populations after an experimental treatment (Scenario 1), nonquenching mode (described in the next paragraph) should be used (2
). See for a trace demonstrating both mitochondrial hyperpolarization (using oligomycin) and depolarization (using FCCP) in rat cortical neurons loaded with Rhod123 in quenching mode.
In nonquenching mode, lower probe concentrations are used (0.5–30 nM, determined empirically) such that dye aggregation and quenching in the mitochondria does not occur. Therefore, depolarized (less negative) mitochondria will have lower cationic dye concentrations and lower fluorescence, and hyperpolarized (more negative) mitochondria will have higher dye concentrations and fluorescence. When using mitochondrial probes in nonquenching mode, acute experimental manipulations can be performed after probe loading (Scenario 2 experiments), or chronic treatments can be performed before dye loading to make a static comparison of pre-existing relative mitochondrial polarization between control and experimental treatments (i.e., Scenario 1 experiments). In summary, Scenario 2 (acute) experiments involve probe loading before the Δψm-affecting experimental treatments and can employ either quenching (if changes in Δψm are fast) or nonquenching modes (if changes in Δψm are slower), whereas Scenario 1 (chronic) experiments involve probe loading after Δψm-affecting experimental treatments and thereby should typically utilize probes in nonquenching modes (since quenching effects are nonlinear).
With the mode determined—quenching or nonquenching—the choice of probe then relates largely to their equilibration characteristics, which varies by probe and determines how each probe behaves in quenching or nonquenching modes. In addition to probe equilibration rates, which will be discussed in more detail below, each probe also has varying degrees of toxicity to mitochondria and cells, principally due to phototoxicity from singlet oxygen generation, and inhibition of the ETC. Each probe also varies in its degree of binding to mitochondria (i.e., non-Nernstian or non-ideal probe behavior), which will result in somewhat greater probe accumulation in mitochondria than would be predicted by the Nernst equation alone. For these reasons, it is always desirable to use the lowest possible probe concentration for any given experimental approach. Thus, as indicators of Δψm
, each probe has relative strengths and weaknesses for different experimental paradigms. briefly outlines fluorescent spectra, some key usage considerations, and potential pitfalls of each, and should be referred to in parallel with the further discussion of these issues (along with Box 1
For example, TMRM inhibits the ETC less, and binds mitochondria less than either Rhod123 or TMRE (in that order), although in practice, at the nonquenching (0.5–30 nM) and lower quenching (>50 nM–1 μM) doses used experimentally, these factors will be negligible for all of these probes, with the possible exception of inhibition of the ETC by TMRE if used above 1 μM (which is not a typical usage paradigm for this probe) (20
). For example, demonstrates that in the depolarized FCCP condition, 1 nM TMRE does not bind to mitochondria in any detectable amount; for these reasons, TMRE remains a common choice for nonquenching studies. Both TMRM and TMRE also equilibrate quickly, another factor making them well suited to nonquenching studies, but are somewhat more complicated to interpret in quenching studies (see below). Rhod123 has only slightly higher mitochondrial binding and ETC toxicity than TMRM, and is more slowly permeant, thus lessening the impact of plasma membrane potential (Δψp
) changes on dye redistribution during short-term experimentation and making Rhod123 often the preferred choice for acute quenching studies to monitor rapid real-time changes in Δψm
following acute cellular insults. DiOC6
(3) has been used most commonly for flow cytometry, but its higher toxicity toward mitochondrial respiration (see Reference 2
, and references therein), and its ultra-low (<1 nM) concentrations required to accurately measure Δψm
) somewhat limit the utility of this probe versus the others. All of the probes can be used for flow cytometry, typically in nonquenching mode.
In contrast to these monochromatic probes, the emission spectra of JC-1 shifts from green to red with increasing concentration (i.e., aggregation) in the mitochondria, thus allowing for a dual-color (green/red) and ratiometric semiquantitative assessment of mitochondrial polarization states. With regard to quenching versus nonquenching, this makes JC-1 something of a hybrid, because its usage relies on readings from both aggregated (red) and non-aggregated (green) probe forms, but in this case probe aggregation results in spectral shift rather than quenching. For some cell types and assay conditions, JC-1 has been reported to be a more reliable indicator of Δψm
than other dyes described herein (21
). On the other hand, while the monomeric (i.e., green-emitting) form of JC-1 appears to faithfully track Δψm
under various experimental conditions, aggregate (i.e., red) JC-1 fluorescence may change independently of Δψm
, for example with H2
). Although pleasing to the eyes, this probe has several other traits and complexities that may render it less ideal for some applications. First, typical JC-1 usage as a ratiometric probe for Δψm
relies on aggregation (threshold) effects, which may render it ill-suited to making reliable distinctions between subtle gradations in Δψm
. It also means that in theory, and as shown in practice (22
) and from our own unpublished data, getting the probe to properly work in this fashion is highly sensitive to probe loading concentrations and loading times. Complicating this matter further, JC-1 is very photosensitive (2
) and its aggregates may not readily equilibrate with monomers. Moreover, all the mitochondrial probes are differentially permeant based on each probe’s distinct molecular structure, and JC-1 is slowly permeant. More specifically, while the monomer (green) form of JC-1 has been reported to equilibrate on a time scale similar to that of TMRM/TMRE (~ 15 min) (22
), the aggregate
(red) form of the dye—the formation of which is required in order to use JC-1 as a ratiometric probe for Δψm
—takes ~90 min to reach equilibration in cardiomyocytes (22
). Thus, since equilibration time is also closely linked to surface-to-volume (S/V) ratios, this means that in any cell populations where there may be significant heterogeneity in S/V ratios either across cells or within different subcellular regions (e.g., soma versus neurites in neurons), JC-1 may be particularly prone to indicating differences in Δψm
when in fact they do not exist. illustrates neuronal staining with JC-1, and also highlights the fact that, depending on localized dye concentrations, either the monomer (green) or the aggregate (red) forms can be seen in polarized mitochondria, which could reflect real subregional differences in Δψm
or it could simply reflect differential JC-1 equilibrium rates (and thus different aggregate formation rates) in subcellular regions with different S/V ratios. (See the legend for further description.) Thus results obtained with JC-1 must be evaluated with these issues in mind and interpreted with caution.
For these reasons, and assuming adequate equilibration time is allowed, an investigator may find that JC-1 is best suited to more coarse “yes/no” assessments with the goal of determining whether mitochondria populations are either largely polarized or largely unpolarized (e.g., apoptosis-type studies), rather than discriminating finer gradations or differences in membrane potential across regions or populations, for which TMRM is likely better suited. On the other hand, as the extensive literature base indicates, JC-1 has been elegantly used in many studies—often chronic Scenario 1 type studies, but acute studies also (e.g., 26,27)—and thus in some cases can be a useful complement to the other probes discussed herein.
Another consideration for probe choice and read-out mode, which relates very closely to probe equilibration kinetics, is which fluorescence intensity will be monitored. Changes in dye distribution in response to changes in Δψm
will alter fluorescence signal in the mitochondria (Fm
), in the cytoplasm (Fc
), in both of these together (i.e., whole cell or cell soma fluorescence) (Fwc
), and in the extracellular bath (Fec
). The direction and kinetics of dye response patterns will vary depending on exactly which of these F
signals is being tracked, and on the equilibration rate and mode (quenching or nonquenching) of the dye in use. Dye behaviors in response to acute changes in Δψm
have been expertly and thoroughly detailed elsewhere, and we refer the reader there for detailed discussion of these issues (2
) (and see Section 7). Similar concepts will apply to determining how these regional intra/extracellular F
intensities should vary in measurements of pre-existing
among different populations (i.e., Scenario 1 studies), keeping in mind that in this situation one is only viewing a static snapshot of dye distribution, rather than tracking distribution changes.
As a somewhat coarse oversimplification of these issues for the sake of clarity to the first-time user, we might suggest the following:
- For short time scale (minutes) studies to monitor rapid step changes in Δψm, using Rhod123 in quenching mode and monitoring Fwc is almost invariably a reliable choice. Rhod123’s slow equilibration may allow plasma membrane (PM) effects to be ignored over short time scales (2) and provides robust, relatively long-lasting whole-cell unquenching or quenching signals which are difficult to miss. In this paradigm, depolarization of Δψm leads to increased Fwc as Rhod123 unquenches from the mitochondria into the cytoplasm, whereas a hyperpolarization of Δψm leads to decreased Fwc as Rhod123 quenches from cytoplasm into the mitochondria. TMRE/TMRM can be used for these purposes as well, but these dyes’ shorter equilibration times means that PM effects must be accounted for, and make for much more transient unquenching (Δψm depolarization) or quenching (Δψm hyperpolarization) bursts in Fwc following step changes in Δψm, due to faster dye re-equilibration across the PM (compare and Figure 2B in Reference 19). This makes the initial change in Fwc easier to miss if capture frequency is not sufficient, and could lead to misinterpretation (i.e., opposite interpretation) of data. Another somewhat advanced distinction that Figure 2 in Reference 19 and Box 2 in Reference 2 nicely make clear is that in quenching mode, it is effectively the change in dye concentration in the cytoplasm/soma as dye is unquenched from or quenched into mitochondria that is responsible for the unquenching or quenching fluorescence burst—hence why Fwc and not Fm is monitored in this mode. This occurs because of the much greater S/V ratio of the mitochondrial inner membrane-matrix versus the PM-cytoplasm interface—i.e., dye leaves or enters mitochondria faster than dye leaves/enters the cell to compensate for this change in intracellular dye concentration—and is why the ΔF unquenching or quenching burst is easier to miss with faster equilibrating dyes like TMRM/TMRE versus slower equilibrating dyes like Rhod123.
- For longer time scale (e.g., several tens of minutes to hours) but still ongoing real-time monitoring of Δψm, using the low-ETC inhibiting and low-binding TMRM in nonquenching mode (1–30 nM, best determined empirically) is often a good choice (28).
- For measuring pre-existing Δψm across different chronically treated populations (Scenario 1 experiments), TMRM/TMRE in nonquenching mode will again usually provide good results. Again, TMRM is reported to exhibit somewhat less ETC inhibition and mitochondrial binding versus TMRE (20), but at low-nanomolar nonquenching concentrations, neither dye should have effects of concern in these areas.
In either of these latter nonquenching experimental paradigms (paradigm 2 or 3), the majority of TMRM/TMRE intracellular fluorescent signal will arise from polarized mitochondria, and therefore Fm can be monitored, which will correlate directly with changes in Δψm: depolarization of Δψm decreases Fm, and hyperpolarization of Δψm increases Fm. However, because in nonquenching mode the ratio of Fm/Fc is very high when mitochondria are polarized, if the read-out method lacks sufficient resolution to distinguish Fm from Fwc (e.g., flow cytometry or plate reader), then Fwc can serve as a satisfactory surrogate for Fm.
Finally, the mode and dye, and thus the signal that will be monitored (Fwc or Fm), will further help dictate the choice of read-out method. Certainly for quenching studies, or other studies requiring single-cell or single-mitochondria resolution, wide field fluorescence microscopy (WFM) (with CCD image capture) or confocal microscopy are reliable and proven choices. With either approach, light/laser intensities should be minimized as much as possible to prevent dye photobleaching and phototoxicity, and if at all possible, limited only to periods of actual image capture/image scanning. In addition, with either microscopy method, for acute studies the exposure (WFM) or scan (confocal) time for a single image must be fast enough to capture the kinetics of the dye response, and to minimize light exposure and prevent motion artifacts. Multiphoton microscopy is also being increasingly applied to studies with mitochondrial probes for Δψm, which will in theory not only allow real-time in vivo studies under some conditions (provided attention is given to all the considerations discussed herein, many of which are amplified in vivo), but should help reduce photobleaching and phototoxicity effects as well. Finally, read-out of Fwc by flow cytometry or fluorescent plate reader—while they lack the resolution of microscopy—may be effective choices for some population studies.
3. Other key dye loading, detection, and monitoring considerations
Regardless of experimental approach and dye method employed, correct interpretation of results requires a keen understanding of all the factors that can affect dye loading, distribution, and equilibrium. For chronic Scenario 1 experiments in particular, where each treatment group is not serving as its own internal baseline against which to measure fluctuations in dye fluorescence, this multitude of considerations in aggregate threatens to become all the more confounding, and thus (perhaps counterintuitively) obtaining reliable results in such chronic studies is sometimes more technically difficult than using the dyes in acute Scenario 2 studies.
For example, the concentration of dye loaded into mitochondria will be a function of, at minimum: bath dye concentration; Δψp; Δψm; mitochondrial size or mass (e.g., S/V ratios); and loading time (at least until equilibrium is reached). As detailed below, these variables must be controlled for in order to obtain interpretable results. For example, if the experimental treatment changes Δψp or mitochondrial mass, then the mitochondrial dye concentration—and thus Fm (and/or Fwc)—in experimental samples will differ from control independent of variations in Δψm. Despite these caveats, using low nonquenching dye concentrations and leaving the dye in the bath during image acquisition (i.e., no washout step; see next paragraph) should allow the investigator to make reproducible qualitative comparisons of post-treatment Δψm across different cell populations, assuming that total dye loading times and total light exposure times are precisely matched among the experimental conditions under comparison and that other factors discussed here are given due attention. In these kinds of Scenario 1 studies, leaving the dye in the bath helps avoid additional confounding experimental factors, such as if the experimental treatment were to affect the rate at which dye unbinds and/or repartitions across the mitochondrial and PM once dye is removed from the buffer—which in turn would change mitochondrial fluorescence independently of Δψm.
Another consideration is loading procedures. With perfectly free diffusion, the preferred approach would be to leave the dye in the perfusion bath during the course of the experiment so that the probe may equilibrate and re-equilibrate in Nernstian fashion with changes in Δψm. This approach, with dye continuously in the perfusion bath, has been frequently prescribed, tested, and applied in many published works. However, since all these dyes bind mitochondria to varying degrees, under many experimental paradigms a perfect Nernstian equilibrium may never be reached as dye remains in the perfusion buffer during the experiment. Despite this caveat, we have found that when TMRM or TMRE is loaded at sufficiently low concentrations (usually 1 nM) and left in the perfusion bath, the mitochondrial fluorescence intensity (Fm) will stop increasing after ~15–30 min (depending on sample conditions), meaning that an equilibrium—even if not a strictly Nernstian equilibrium due to some degree of unavoidable probe binding—has been reached. At this time, we find Fm will respond appropriately to known manipulations of Δψm.
On the other hand, under some experimental conditions the user may find that when probe remains in the perfusion bath, Fm may continue to increase with time, and never reach a stable equilibrium. There are several reasons this could occur. First, dye concentrations may be too high, and the user can determine whether the problem is corrected by lower dye loading concentrations. Second, the sample type may be such that unavoidable probe binding is very difficult to saturate, as might occur with very high cell densities, or tissue slices, for example. In this case, lower probe concentrations and/or lower cell densities can be explored. Other ways to remedy a continuously increasing Fm when probe remains in the perfusion bath include loading the mitochondrial dye for 15–30 min, followed by washout, then performing image acquisition with reduced concentrations or no dye in the perfusion buffer. (In some cases, by lowering fluorescence background, this approach may also improve signal-to-noise ratio.) Under these conditions, once bath dye concentrations are reduced or removed, mitochondria-loaded dye will repartition to equilibrium across both the mitochondrial and plasma membranes. Thus even if dye is completely removed from the perfusion buffer after dye loading, probe will still remain in the imaging bath buffer due to this re-equilibration. For these reasons, whenever the dye concentration in the imaging buffer is lower than the dye concentration in the loading buffer, it is imperative that the dye is allowed to fully re-equilibrate before imaging. This is particularly important for slow-equilibrating dyes such as JC-1. Despite the merit of this approach in some situations, at least for experiments in nonquenching mode, it may be preferable to maintain equivalent dye concentrations in loading and imaging buffers whenever possible to eliminate any possibility of misinterpreting Nernstian dye redistribution as an experimentally induced effect.
The converse situation may also exist whereby the mitochondrial dye loads slowly, and never appears to reach an equilibrium state. Often this may indicate efflux of the dye through rhodamine-sensitive multi-drug resistance transporters (MDR) (particularly in tumor cell lines) (6
), which can be confirmed by blocking the MDR with verapamil or cyclosporin H.
Loading, equilibration, and binding considerations aside, there are other reasons that Fm
may change independently of Δψm
. Namely, these cationic probes also have varying degrees of phototoxicity, and others have shown that light exposure by itself can actually increase
fluorescent TMRM signal, for reasons thought to be related to oxidant production (25
). We have observed similar phenomena with TMRE under some experimental conditions (unpublished data). In other cases, significant and rapid photobleaching of these dyes may be observed with light exposure. In either case, minimizing total light exposure times is critical. For all these reasons, under Scenario 1 experiments, because experimental treatments are applied before probe loading, it is imperative that dye loading times and total duration of light exposure be precisely matched for any experimental conditions or individual fields under comparison. For example, following dye loading, the simple act of looking at fields of view under the microscope using mercury or xenon lamp illumination—unless total light exposure time and recovery time is identical
for each condition (nearly impossible to achieve in practice)—will often impact the dye signal enough to render comparisons of Fm
among the experimental conditions meaningless after such observation. Therefore for such Scenario 1 population studies, either fully automated microscopy (with automated field selection and image capture, such that specimens are only
subject to low-intensity light during brief sub-second image exposures) or other read-out methods such as flow cytometry or a fluorescent plate reader may be most appropriate to minimize the confounds of light exposure on Fm
With due attention to these kinds of details, verifiable and reproducible results can be obtained regardless of the approach chosen.
4. Use agents that affect Δψm to confirm directional dye behavior
Whatever method of dye usage one chooses for their experiments, it is critical to validate the approach in their experimental system using agents that affect Δψm
to confirm that directional changes in mitochondrial fluorescence can be interpreted as predicted. For example, oligomycin (usually 2–5 μg/mL) will slightly hyperpolarize mitochondria by blocking proton re-entry via the ATP-synthase (7
). Under nonquenching conditions, this will translate to a slight increase
, and a slight (transient) decrease
under quenching conditions. It is important to note, however, that in unhealthy mitochondria with compromised energetics—for example, inhibited respiration and/or with proton leaks—the Δψm
may be maintained by reversal of the ATP-synthase, in which case oligomycin treatment will cause increased interior [H+
] and mitochondrial depolarization (7
). Thus protonophores such as FCCP or carbonyl cyanide m-chloro phenyl hydrazone (CCCP) (usually ~0.25–5 μM), which depolarize mitochondria by increasing their permeability to protons, should be co-employed to further confirm dye behavior. In nonquenching mode, these compounds will decrease Fm
. In quenching mode, FCCP/CCCP will transiently increase Fwc
as quenched dye exits the mitochondria and becomes unquenched (thus brighter) in the cytoplasm, followed by a delayed decrease in Fwc
as dye continues to exit mitochondria and cytoplasm and re-equilibrates across the PM with continued depolarization (see Box 2 in Reference 2
). Following protonophore treatment, the duration of the transient increase in mitochondrial fluorescence caused by dye unquenching will differ with varying equilibration rates of each probe. These changes are well described in more detail elsewhere (2
). The investigator should be aware of additional important considerations for FCCP/CCCP, namely that while lower concentrations (e.g., 250 nM) will specifically collapse Δψm
, high concentrations (>2.5 μM) (likely to be variable with cell type) will also significantly diminish Δψp
)—an important consideration if the investigator wishes to selectively collapse Δψm
but not Δψp
. Findings of another study suggested that in cerebellar granule neurons, FCCP may depolarize Δψp
at least in part by glutamate release and NMDA receptor activation (28
), perhaps either secondary to or downstream of its protonophoric or metabolic effects on Δψm
. Addition of NMDA receptor antagonists/glutamate antagonists such as MK-801 may help determine if FCCP is acting in this fashion in a particular model system. These examples illustrate additional factors, likely to vary with cell type, that must be taken into account when interpreting effects of FCCP/CCCP on Δψm
5. Control or account for changes in Δψp
As discussed above, a cell’s Δψp will impact the amount of these cationic dyes entering the cytoplasmic space, thereby also affecting how much dye is available for mitochondrial uptake. Therefore, when using probes for Δψm, it is imperative to employ additional controls or modeling to ensure that altered Δψp is not responsible for any observed changes in Fm under the experimental conditions. Conversely, a change in Δψp that only occurs with experimental treatment could mask a legitimate (opposing) change in Δψm.
One approach is to use a complementary probe under parallel experimental conditions, such as bis-(1,3-dibutylbarbituric acid) trimethine oxonol [DiBAC4
(3)], a relatively specific probe for Δψp
. An increase in the anionic DiBAC can be interpreted as a depolarization of Δψp
, whereas decreased intracellular DiBAC fluorescence represents an increase in Δψp
). In Scenario 1 experiments, this approach can be used to obtain a “static” snapshot of whether Δψp
is also altered by experimental treatment, to help validate whether or not observed changes in Fm
are due to altered Δψm
alone, or to changes in Δψp
(or both) (5
Another clever approach is to obviate the dependence of Fm
by monitoring the ratio of fluorescence intensity between mitochondria-rich and mitochondria-poor regions of single cells (31
); this method and the methods described herein may prove quite useful for many laboratories. However, it has been pointed out that this method requires nonquenching concentrations of probe (and thus may not be suitable for all experimental needs) and that estimating the rather small differences in fluorescence intensity between background and mitochondria-poor areas may introduce some measurement error (18
Advanced methodologies have also been demonstrated for addressing relative contributions of Δψm
in real-time Scenario 2 type experiments, which include using computer modeling to simulate and isolate the effects of changes in Δψm
), parallel tracking of real-time changes in Δψp
using DiBAC (28
), and a method for simultaneous real-time tracking of both Δψm
using TMRM and a proprietary probe for monitoring Δψp
(PMPI, Molecular Devices, Sunnyvale, CA) (18
; also see Reference 33
These methods elegantly demonstrate, for example, that KCl or other cell-depolarizing treatments are indeed helpful for illustrating the impact of Δψp
on mitochondrial dye equilibration behaviors in the context of real-time Scenario 2 type studies and simulation modeling (i.e., when the mitochondrial dyes are loaded before addition of KCl) (18
). On the other hand, because regulation of Δψp
are intricately intertwined with intracellular ionic compositions and with each other—and because a depolarization of Δψp
will severely limit the amount of dye available for mitochondrial uptake—investigators may find that for Scenario 1 studies (i.e., dye is loaded after experimental manipulation), loading the mitochondrial dye in the presence of KCl as a control for distinguishing the relative contributions of Δψp
to an experimentally observed Fm
is of marginal diagnostic value. Rather, the other approaches described in this section should yield more useful information as to whether or not changes in Δψp
are causing experimentally observed changes in Fm
7. Utilize multiple approaches to quantifying fluorescent changes
Particularly if not employing software modeling of dye distribution behavior (see Section 5 above, and Section 9 below), investigators can consider using multiple analytical approaches to quantify changes in mitochondrial fluorescence under their experimental conditions. The most commonly employed approach is simply monitoring Fwc (in quenching mode) or Fm (in nonquenching mode) as described in Section 3 above. With changes in Δψm (or Δψp), changes in Fm will in turn be accompanied by opposing changes in Fc and Fec fluorescence, as probe redistributes among mitochondrial, cytoplasmic, and extracellular compartments. However, Fc signal is small, especially at lower dye concentrations, and moreover, Fc changes that accompany Fm changes will be (often rapidly) transient as dye redistributes across the PM. Therefore, an approach that we have found useful is to instead monitor Fec along with Fm (). If Fm and Fec are not mirroring each other in an opposing manner with oligomycin and FCCP treatment, for example, then the probes are not behaving in a fashion that will accurately predict Δψm and this methodological flaw must be identified and corrected before proceeding.
Acute tracking of nonquenching mitochondrial TMRE
In addition, performing quantitative numerical analyses beyond measuring raw intensity changes in Fm
can also further validate the interpretation of results. For example, examining the coefficient of variance of pixel intensities (CV = standard deviation/mean of Fwc
pixel intensities)—a representation of the spatial heterogeneity of intracellular dye distribution between cytoplasmic and mitochondrial compartments—can further confirm results obtained from looking at Fm
) and has the added benefit of helping to obviate any effects of a changing Δψp
. By this method, depolarization of Δψm
will lead to a more uniform intracellular distribution of probe as dye exits the mitochondrial pool, resulting in greater spatial homogeneity of the Fwc
fluorescent signal, and thus a reduction in the variation of individual pixel values around the mean (i.e., a reduced CV). Hyperpolarization of Δψm
will, in contrast, result in increased CV. While CV, like Fm
, is still sensitive to changes in cell morphology or mitochondrial localization which must be accounted for (see Section 8), it nonetheless may provide a useful complementary analysis to confirm observed changes in Fm
8. Control for changes in mitochondrial morphology, localization, or mass
Under some experimental conditions, it will be desirable to further ensure that any observed fluorescence changes correspond to changes in Δψm, rather than to changes in mitochondrial morphology, localization, or mass, changes in which can by themselves affect readings of Fm or Fwc. These controls are particularly important for nonquenching Scenario 1 type experiments in which dye fluorescence measurements are taken after experimental treatment using ensemble averaging (e.g., low-resolution analysis of dye fluorescence over whole cells) or over cell populations. Moreover, because of S/V ratios discussed in Section 2, more slowly permeant dyes like JC-1 used in a nonquenching experiment of this nature will still be more sensitive to the effects of changes in mitochondrial morphology or mass (i.e., volume). In fairness, even with best-practice efforts at some of these approaches below, we must note that quantifying mitochondrial morphology and/or mass can be a very difficult thing for any investigator do reliably, since (i) none of the assays themselves is without inherent sources of error, just as (ii) no one approach will be a perfect correlate for total mitochondrial mass/content under all conditions.
The first-line approach to control for these factors is to use qualitative (and if possible, quantitative, using image processing) visual assessment of whether changes in any of these parameters are occurring under the experimental conditions. Another complementary approach is to assess mitochondrial staining with Mitotracker (Molecular Probes, Invitrogen, Carlsbad, CA, USA) or other mitochondria binding fluorescent probes to determine whether these probes show intensity differences under experimental conditions identical to those used for monitoring Δψm
. A key caveat to this approach is that many probes traditionally used in this fashion (e.g., Mitotracker probes or nonyl acridine orange) exhibit varying sensitivity to Δψm
and/or inhibition of the ETC (35
). Nonetheless, if used judiciously with these caveats and limitations in mind, mitochondria tracking probes of this nature may provide some relative impression of whether mitochondrial mass, morphology, or localization is changing under the experimental conditions and to determine if these factors may be contributing to changes in Fm
observed with TMRM, Rhod123, JC-1, or similar probes.
Another alternative to help control for mitochondrial localization and shape is the use of mitochondria-targeted fluorescent proteins [e.g., mtGFP (37
)], which have been used for mitochondrial tracking and fusion/fission studies, among others (39
). In many cases, these may prove more reliable than Mitotracker probes for assessing mitochondrial shape and localization, although mention of a few possible drawbacks and cautions is warranted. First, because these will require transfection or similar method to induce expression in cells, not all cells will be transfected, with transfection percentages depending on cell type. Second, because expression levels of the mitochondria-targeted fluorescent proteins will vary with many parameters besides mitochondrial mass and because their fluorescence intensity can also vary with many factors (e.g., with pH for some variants, see Section 10 and case study in Box 2
), these tools are best used for assessment of mitochondrial localization and shape, rather than mass. Finally, if they are used simultaneously
with any of the mitochondrial probes discussed herein, the investigator must be absolutely certain the two fluorophores can be spectrally separated with no cross-talk and that they do not interfere with the mitochondrial probe behavior, either of which will confound results.
Other means to assess relative mitochondrial mass under parallel experimental conditions include measuring the ratio of mitochondrial to nuclear DNA (mtDNA/nDNA), immunoblot analysis for key mitochondrial proteins (e.g., cytochrome c, Complex V), and/or analysis of mitochondrial protein content or weight following mitochondrial isolation procedures. Operating on a similar principle, one can also measure activity of the mitochondria-exclusive tricarboxylic acid (TCA) cycle enzyme citrate synthase (41
), readily done with commercially available kits (Sigma Aldrich, St. Louis, MO, USA).
To further control for changes in mitochondrial localization that may affect quantification of mitochondrial dye signal when using ensemble-averaging approaches, Fm and Fec under each experimental condition can be assessed and compared before and after addition of a protonophore (FCCP or CCCP). Analyzing the net change in Fm versus Fec before and after complete mitochondrial depolarization can help to control for influences of mitochondrial mass or localization on Fm or Fwc when using ensemble averaging approaches. However, controls for Δψp must still be performed.
10. Employ complementary tools to assess the ΔpHm component of Δp
With their utility as indicators of Δψm
, coupled with the proton’s central role in mitochondrial bioenergetics, it is tempting to assume that distribution behavior of these mitochondrial dyes specifically reflects the H+
gradient between the mitochondrial exterior and interior (i.e., ΔpHm
, the pH component of Δp (recall from the Introduction that Δp = Δψm
). In other words, one might be tempted to conclude that increased dye uptake (hyperpolarized mitochondria) also indicates decreased mitochondrial proton concentration ([H+
), or that decreased mitochondrial dye uptake (depolarized mitochondria) reflects increased [H+
. To do so would be to equate Δψm
, when in fact these are each separate contributors to Δp (see the Introduction), or to assume that Δψm
will always follow ΔpHm
. On the contrary, because Δψm
, and because changes in Δψm
may often but do not necessarily follow changes in ΔpHm
, additional tools have been developed to assess the ΔpHm
component of Δp. For example, mitochondria-targeted pH sensitive fluorescent fusion reporter proteins whose fluorescence intensity varies with pH (42
), or pH-sensitive probes like SNARF-1 targeted to mitochondria (44
), can be used to directly assess mitochondrial pH. These tools can be used together with assessments of cytosolic pH using similar cytosol-targeted pH-sensitive fluorescent fusion reporter proteins (42
) or the pH-sensitive fluorophore 2′-7′-bis(carboxyethyl)-5(6)-carboxyfluorescein acetoxymethyl ester (BCECF-AM) (43
), to directly calculate ΔpHm
across the mitochondrial membrane. Precisely because cationic dyes for monitoring Δψm
can only respond to charge gradients and not specifically to proton gradients, these additional tools are useful or even imperative (see case study in Box 2
) complementary methods by which to assess both Δψm
as independent and sometimes divergent components of Δp.