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CHAPTER 14

Imaging of Mitochondrial Polarization and Depolarization with Cationic Fluorophores

John J. Lemasters and Venkat K. Ramshesh

Center for Cell Death, Injury and Regeneration, and Departments of Pharmaceutical Sciences and Biochemistry & Molecular Biology Medical University of South Carolina, Charleston, South Carolina 29425

I. Introduction II. Quantitative Imaging of DC with Fluorescent Cations A. Nernstian Distribution of Cationic Fluorophores B. Cellular Loading of Potential-Indicating Fluorophores C. Image Acquisition and Processing D. Nonideal Characteristics of Fluorophores III. Visualization of Depolarized Mitochondria A. Covalent Adduct Formation by MitoTracker Probes B. FRET Between Cationic Fluorophores C. FRET Between MitoTracker Green FM and TMRM D. Use of FRET to Distinguish Depolarized from Polarized Mitochondria IV. Conclusion References

I. Introduction

Mitochondrial respiration generates an electrochemical gradient of protons made up mostly of a negative electrical potential diVerence (ÁÉ) across the mitochondrial inner membrane. In living cells, the plasma membrane also generates a negative inside ÁÉ. Electrophysiological techniques have long been used to measure plasmalemmal ÁÉ, but mitochondria are in general too small to impale with a microelectrode. Instead, mitochondrial ÁÉ must be estimated from the equilibrium distribution of

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membrane-permeant cations, which accumulate electrophoretically into polarized cells and mitochondria (Hoek et al., 1980). By using fluorescent lipophilic cations, ÁÉ in individual cells and mitochondria can also be visualized and quantified (Chacon et al., 1994; Ehrenberg et al., 1988; Farkas et al., 1989; Zahrebelski et al., 1995). By coupling fluorescent cation uptake with fluorescence resonance energy transfer (FRET), mitochondria undergoing depolarization can also be revealed selectively (Elmore et al., 2001, 2004).

II. Quantitative Imaging of DC with Fluorescent Cations

A. Nernstian Distribution of Cationic Fluorophores Membrane-permeant monovalent cationic fluorophores, such as rhodamine 123, tetramethylrhodamine methyl ester (TMRM), and many others, accumulate electrophoretically within cells in response to the electrical potential É of subcellular compartments (Ehrenberg et al., 1988; Emaus et al., 1986; Johnson et al., 1981). At equilibrium, É is related to fluorescent cation uptake by the Nernst equation: Fin É ¼ À59 log ð1Þ Fout where É is electrical potential in millivolts, Fout is the monovalent cationic fluorophore concentration in the extracellular space (electrical ground), and Fin is the fluorophore concentration at any point within the cell. By using confocal/ multiphoton microscopy to quantify intracellular fluorophore distribution, maps of intracellular É can be generated using Eq. (1). From diVerences of É between compartments, membrane potentials (ÁÉ) can be determined, specifically plasmalemmal ÁÉ (cytosolic minus extracellular É) and mitochondrial ÁÉ (mitochondrial minus cytosolic É). Under normal conditions in most cells, plasmalemmal ÁÉ ranges from À30 to À100 mV and mitochondrial ÁÉ ranges from À120 to À160 mV. Since these ÁÉ's are additive, mitochondria are 150­260 mV more negative than the extracellular space. From Eq. (1), such large ÁÉ's correspond to cation concentration ratios between mitochondria and the cell exterior that can exceed 10,000:1. Such large gradients cannot be captured in digital images using a conventional linear scale of 256 gray levels per pixel (8-bit pixel memory). Instead, 12- to 16-bit memory, sequential imaging at diVerent laser powers, or a nonlinear logarithmic (gamma) scale must be used (Chacon et al., 1994). Gamma scales, commonly used in scanning electron microscopy, compress input signals logarithmically into the available 256 gray levels of pixel memory. Early confocal microscopes, such as the Bio-Rad MRC600, included gamma imaging circuits, but gamma circuitry is generally not available in current commercial laser scanning confocal/multiphoton microscope systems. Accordingly, this chapter addresses measurement of ÁÉ's using high bit memory and sequential imaging.

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B. Cellular Loading of Potential-Indicating Fluorophores Cells plated on glass coverslips are loaded by simple incubation with 50­500 nM of TMRM, rhodamine 123, or other indicator, for 15­30 min at 37 C. No special loading buVer is required, and cells can be loaded with equal success in complete culture medium or in a simple Ringer's or saline solution. However, when the loading buVer is replaced, the new medium should contain a third to a fourth of the initial loading concentration of the potential-indicating probe in order to maintain its equilibrium distribution. Otherwise, with each buVer change, some fluorophore will leach from the cells irrespective of changes of intracellular É. Loading concentrations above 1 mM should be avoided. Even with loading below 1 mM, fluorophore accumulation can reach millimolar concentrations inside the mitochondrial matrix space. Such high concentrations can cause metabolic inhibition. For example, rhodamine 123 at high matrix concentrations inhibits the mitochondrial ATP synthase of oxidative phosphorylation (Emaus et al., 1986), and DiOC (6), a cationic fluorophore still used in flow cytometry, strongly inhibits mitochondrial respiration even when loaded at nanomolar concentrations (Rottenberg and Wu, 1997). Of commonly used fluorophores, TMRM exhibits the least mitochondrial toxicity (Scaduto and Grotyohann, 1999). In general, fluorophore concentrations exceeding 500 nM should be avoided unless an adequate fluorescent signal cannot otherwise be produced.

C. Image Acquisition and Processing After loading, confocal and multiphoton images may then be collected. At least two images must be collected. The first is an optical section through the specimen of interest. The second is an image obtained after refocusing inside the glass coverslip. The latter image serves as a background in an area devoid of fluorophore. Both images should be collected using identical instrumental settings of laser power, gain, and brightness. Image oversaturation (pixels at highest gray level) and undersaturation (pixels with a zero gray level) should be negligible. A low laser power setting ( 1%) should be used to minimize photodamage, especially if serial imaging over time of the same portion of the specimen will be performed. If the microscope can collect images with pixel depth of 12 or 16 bits (4096 and 65,536 gray levels, respectively), then additional images may not be needed. However, if pixel depth is only 8 bits (256 gray levels) or if the ratio of fluorescence intensity between mitochondria and the extracellular space exceeds the gray level range, then additional images at $10 times greater laser power ($3.3 times laser power for two-photon microscopy since two-photon excitation varies with the square of laser intensity) should be collected both in the cell and within the glass coverslip. With multitracking systems, lower and higher laser power images can be collected simultaneously one line at a time. Each line (row of pixels) in the images is collected in sequence. A first line scan is performed at lower laser power and is

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followed immediately by a second line scan at higher laser power, a process that is repeated with each succeeding line in the image. Multitracking permits the two images to be acquired simultaneously and eliminates problems associated with specimen movement and focus drift. When images at higher laser power are collected, oversaturation of mitochondria will be marked, but oversaturation in higher power images does not pose a problem, since these images will only be used to quantify areas of weak fluorescence intensity in the extracellular space. By processing images collected through the sample and within the glass coverslip, maps of the intracellular distribution of É may be generated by a three-step procedure of: (1) background subtraction, (2) quantitation of extracellular fluorescence, and (3) calculation of É for each pixel gray level to create a pseudocolor map of É.

1. Background Subtraction

Unless photon counting or an equivalent procedure is performed, light detectors such as photomultiplier tubes generate signals even in the absence of light. This background signal must be subtracted from the signal collected in the presence of light to obtain an output truly proportional to fluorescence intensity. In confocal/ multiphoton microscopy, images collected in the plane of the coverslip represent this background, since added fluorophore cannot penetrate into the glass. A mean pixel value is then calculated for all the pixels of each background image using utilities in ImageJ, Adobe Photoshop, MetaMorph, or other image analysis software. This average value is then subtracted from every pixel of the corresponding specimen image. Such background-corrected images represent the true relative distribution of fluorescence intensity within the images.

2. Fluorescence of the Extracellular Space

The next step is to estimate extracellular fluorescence. Since fluorescence should be the same everywhere in the extracellular space, all areas in the cell-free extracellular space in background-subtracted specimen images are selected, and average pixel intensity is determined. If extracellular fluorescence is too weak, then extracellular fluorescence can be measured in the same way in a higher laser power image. Division (after background subtraction) of mean extracellular fluorescence from the higher laser power image by the power ratio (or its square for twophoton excitation) between the higher and lower power images then yields an estimate of extracellular fluorescence for the lower power image. The latter estimate is valid even if it is less than one.

3. Pixel-by-Pixel Calculation of É

Using Eq. (1), a value for É can be calculated for every pixel of backgroundcorrected images on the assumption that fluorescence intensity is proportional to monovalent cationic fluorophore concentration. To display the intracellular

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distribution of É, colors can be assigned to specific millivolt ranges of É. To determine the pixel value corresponding to a specific millivolt value of É, Eq. (1) is rearranged: Pi ¼ antilog ðlog Pout À É=59Þ ð2Þ

where Pout is average background-subtracted pixel intensity in the extracellular space, and Pi is the pixel value representing a particular millivolt value of É. Using Eq. (2), ranges of gray level values corresponding to a specific range of É can be calculated, to which individual colors are assigned. Figures 1 and 2 illustrate the pseudocolored images that result for a TMRMloaded mouse hepatocyte and adult feline cardiac myocyte imaged by confocal microscopy. The diVerence of É between the extracellular space (where É is zero) and the cytosol and nucleus represents plasmalemmal ÁÉ, whereas the diVerence between the cytosol/nucleus and mitochondria represents mitochondrial ÁÉ. In the hepatocyte, plasmalemmal ÁÉ was about À30 mV, and mitochondrial ÁÉ was as

(mV): -150 -120 -90 -60 -30 0

25 mm

Fig. 1 Distribution of electrical potential in a mouse hepatocyte. A mouse hepatocyte plated on a

Type 1 collagen-coated coverslip for 5 h was loaded with 200 nM TMRM for 20 min in KRH at 37 C and imaged in KRH containing 50 nM TMRM. The distribution of É was determined by laser scanning confocal microscopy using 543-nm excitation from a helium­neon laser and a 565- to 615-nm emission barrier filter.

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(mV): -200 -160 -120 -80 -40 0

20 mm

Fig. 2 Electrical potential of an adult feline cardiac myocyte. An adult feline cardiac myocyte plated on a Matrigel-coated coverslip for 5 h was loaded with 200 nM TMRM for 20 min in Medium 199 at 37 C and imaged in Medium 199 containing 50 nM TMRM, as described in Fig. 1.

great as À120 mV. In the cardiac myocyte, the cytoplasm was so densely packed with mitochondria that cytosolic É needs to be estimated from the nucleus, which is equipotential with the cytosol. From nuclear É, plasmalemmal (sarcolemmal) ÁÉ was estimated to be about À100 mV. The greater plasmalemmal ÁÉ of the cardiac myocyte is consistent with the greater plasmalemmal polarization of excitable cells such as myocytes. However despite the greater plasmalemmal ÁÉ, mitochondrial ÁÉ of the myocyte was similar to that of the hepatocytes. In hepatocytes and cardiac myocytes, mitochondrial diameter is about 1 mm. Since the thickness of confocal and multiphoton optical sections imaged with a high numerical aperture lens is slightly less than 1 mm, some mitochondria occupy the entire thickness of the optical slices, but other mitochondria will occupy only part of the thickness of the section. The presence of mitochondria that are only partially included in confocal/multiphoton optical sections causes an apparent heterogeneity and underestimation of mitochondrial ÁÉ. Consequently, the highest values of mitochondrial É most likely reflect true mitochondrial ÁÉ.

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D. Nonideal Characteristics of Fluorophores The Nernst equation [Eq. (1)] describes ideal electrophoretic distribution of membrane-permeant monovalent cations, but probes may not behave ideally. Rhodamine 123, for example, can accumulate in mitochondria to levels that exceed those predicted by Eq. (1) (Emaus et al., 1986). Excess accumulation may in part be due to concentration-dependent fluorophore stacking and formation of so-called J-aggregates. For fluorophores like rhodamine 123, TMRM, and others, J-aggregate formation causes fluorescence quenching and a redshift of absorbance. In cuvette and multiwell assays, absorbance changes and fluorescence quenching can be used to monitor mitochondrial fluorophore accumulation and hence mitochondrial ÁÉ (Blattner et al., 2001; Emaus et al., 1986; Scaduto and Grotyohann, 1999). J-aggregate formation and quenching are dependent on fluorophore concentration and may be minimized by using smaller loading concentrations. With some monovalent cationic fluorescent probes, such as JC-1, J-aggregate formation leads to a shift from green to red fluorescence emission rather than to fluorescence quenching (Smiley et al., 1991). As more JC-1 accumulates in mitochondria, more and larger J-aggregates form. Thus, increased red fluorescence of JC-1 signifies increased mitochondrial polarization. Confocal images of JC-1-loaded hepatocytes visualize these aggregates of JC-1 within the matrix of individual green-fluorescing mitochondria (Fig. 3). These J-aggregates are literally microprecipitates and characteristically do not fill the mitochondrial matrix. In Fig. 3, J-aggregates are seen to localize to the lateral margins of mitochondria, presumably in association with mitochondrial membranes. Heterogeneity of red and green fluorescence in single JC-1-loaded mitochondria represents the physical distribution of J-aggregate microprecipitates and does not signify variation of ÁÉ within the single organelles. Formation of physical aggregates of JC-1 inside mitochondria limits the probe's usefulness for high-resolution imaging of cellular and mitochondrial ÁÉ.

III. Visualization of Depolarized Mitochondria

A. Covalent Adduct Formation by MitoTracker Probes When mitochondria depolarize, they release their potential-indicating fluorophores. As a result, the organelles disappear from fluorescence images. To circumvent this problem, membrane-permeant cationic fluorophores have been developed that incorporate reactive chloromethyl groups that form covalent bonds with protein sulfhydryls (Haugland, 1999). These MitoTracker probes, such as MitoTracker Green and MitoTracker Red, accumulate electrophoretically into mitochondria in the same fashion as do other potential-indicating cationic fluorophores (Fig. 4, left panel). After intramitochondrial accumulation, adducts form between MitoTracker probes and mitochondrial matrix proteins, although adduct formation may require many minutes to go to completion. After adduct formation, MitoTracker fluorescence

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Nuc

2 mm

Fig. 3 Red and green mitochondrial fluorescence after hepatocyte loading with JC-1. A mouse hepatocyte plated as in Fig. 1 was loaded with 100 nM JC-1 for 30 min in KRH at 37 C and imaged by multitrack confocal microscopy. Shown is an overlay of green fluorescence excited with 488-nm light and red fluorescence excited with 543-nm light. Note that many green-fluorescing mitochondria contain peripheral red J-aggregate inclusions. Nuc, nucleus.

becomes independent of mitochondrial polarization and is retained even if mitochondria subsequently depolarize (Fig. 4, right panel). MitoTracker fluorescence may even survive chemical fixation with aldehydes and other chemical preservatives. Thus, MitoTracker labeling of mitochondria has been valuable for use in conjunction with immunocytochemistry and related procedures that require some type of fixation. B. FRET Between Cationic Fluorophores When fluorophores with diVerent spectral characteristics come into proximity, the phenomenon of FRET can occur (Stryer, 1978). In FRET, photons excite a donor fluorophore to an excited state. If an acceptor fluorophore is nearby whose excitation spectrum overlaps the donor emission spectrum, then nonradiative energy transfer can occur from the donor to the acceptor, leading to fluorescence

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MTG

MTG+TMRM

De-energized

20 mm

Fig. 4 FRET-dependent quenching of MTG by TMRM in cultured rat hepatocytes. Overnight cultured rat hepatocytes were loaded with 0.5 mM MTG for 60 min in Waymouth's growth medium, and green and red fluorescence was imaged by confocal microscopy using blue (488-nm) excitation from an argon­ krypton laser (left panel). Subsequently, 1 mM TMRM was added (middle panel), followed by de-energization with 2.5 mM KCN and 10 mM CCCP in the presence of 20 mM fructose and 1 mg/ml oligomycin (right panel). After MTG loading, mitochondria fluoresced green but not red (left panel). After subsequent TMRM loading, green fluorescence was quenched, and mitochondria fluoresced red after blue excitation (middle panel). De-energization restored green MTG fluorescence and simultaneously caused red TMRM fluorescence to disappear.

emission at wavelengths characteristic of the acceptor rather than the donor. When FRET occurs, ordinary donor fluorescence becomes quenched. Instead, longer wavelength photons are emitted with wavelength characteristic of the acceptor. FRET varies with the sixth power of the distance between donor and acceptor. For FRET to occur to any appreciable extent, donor and acceptor molecules must be within 10 nm of one another.

C. FRET Between MitoTracker Green FM and TMRM MitoTracker Green (MTG) FM is an example of a cationic MitoTracker probe that accumulates into mitochondria and binds covalently to mitochondrial proteins. Green-fluorescing MTG can serve as a donor for FRET with TMRM as the acceptor, a phenomenon likely promoted by stacking interactions between the two fluorophores. When cultured rat hepatocytes are loaded with 0.5 mM MTG for 1 h in culture medium, green MTG fluorescence excited with blue (488 nm) light shows a characteristic mitochondrial pattern in confocal images (Fig. 4, left panel). Red fluorescence is absent. After incubation with 1 mM TMRM, green fluorescence excited with blue light becomes quenched and red fluorescence occurs instead (Fig. 4, middle panel). Since blue light does not excite red TMRM fluorescence, the observed shift from green to red emission represents FRET. This shift of emission is reversible, since de-energization of mitochondria with KCN (a respiratory inhibitor), CCCP (an uncoupler), and oligomycin (a mitochondrial ATPase inhibitor) in the

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presence of fructose (a glycolytic substrate to maintain ATP) restores green fluorescence as red fluorescence is lost (Fig. 4, right panel). TMRM alone in the absence of MTG produces little red fluorescence after blue excitation until MTG is loaded (data not shown) (Elmore et al., 2004). Radiative colorimetric quenching in which green fluorescence emitted by MTG is reabsorbed by TMRM, leading to red fluorescence, cannot account for these observations, since photon absorbance over the optical path length involved (i.e., the diameter of a mitochondrion or 1 mm) is negligible (Elmore et al., 2001, 2004). D. Use of FRET to Distinguish Depolarized from Polarized Mitochondria Mitochondrial depolarization frequently contributes to necrotic and apoptotic killing of cells from liver, heart, and other organs. The cellular fate of depolarized mitochondria is diYcult to study because after depolarization, labeling of mitochondria with potential-indicating probes like rhodamine 123 and TMRM simply disappears. Because MitoTracker probes are retained after depolarization, MitoTracker dyes do not distinguish mitochondria that depolarize from those that remain polarized. A strategy to visualize depolarized mitochondria selectively is to coload cells with MTG and TMRM. After coloading, TMRM quenches MTG fluorescence by FRET when the two fluorophores reside together inside polarized mitochondria. When an individual mitochondrion depolarizes, it releases TMRM, and MTG becomes unquenched. As a result, the fluorescence of the depolarized mitochondrion changes from red to green. Such green fluorescence allows discrimination of a single depolarized mitochondrion against a background of hundreds of polarized mitochondria. For coloading, cells cultured on coverslips or glass-bottomed Petri dishes are incubated at 37 C with 200 to 500 nM MTG FM for 1 h in either complete culture medium or a saline solution such as Krebs-Ringer-HEPES solution (KRH containing: 25 mM HEPES, 115 mM NaCl, 5 mM KCl, 1 mM KH2PO4, 1.2 mM MgSO4, and 2 mM CaCl2, pH 7.4). After mounting the cells on the microscope, TMRM (0.5­1 mM) is added for 30 min followed by baseline imaging, as shown for cultured rat hepatocytes in Fig. 5. For eVective MTG quenching, TMRM must be used at a higher concentration than when TMRM is used alone, e.g. 1 mM. After coloading with TMRM and MTG, green MTG fluorescence excited with blue light was quenched in nearly all mitochondria, but due to FRET, nearly all mitochondria fluoresced red (Fig. 5). A few mitochondria, however, did fluoresce green (circles), but none were yellow in the overlay of the red and green fluorescence images, signifying that no individual structures within the cells were simultaneously emitting red and green fluorescence (Fig. 5, right). Since cationic TMRM and MTG are both initially taken up only by polarized mitochondria, recovery of the green fluorescence of covalently bound MTG and loss of red fluorescence indicates spontaneous depolarization of these mitochondria. When the hepatocytes were then incubated in nutrient-free KRH plus the hormone glucagon, individual

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Serum

KRH+G

Baseline

60 min

20 mm

Fig. 5 Spontaneous mitochondrial depolarization in rat hepatocytes during nutrient deprivation plus glucagon. Cultured hepatocytes were loaded sequentially with MTG (0.5 mM) and TMRM (1 mM) in serum containing complete growth medium and mounted on the stage in growth medium. After collection of a baseline image (left panel), the cells were incubated in KRH plus 1 mM glucagon (KRH þ G) for 60 min (right panel). Shown are overlay confocal images of green and red fluorescence after blue (488 nm) excitation. Green-fluorescing structures increased when the medium was changed to KRH plus glucagon (circles).

mitochondria reverted from red to green fluorescence (Fig. 5, right panel, circles). Most of these mitochondria were undergoing lysosomal digestion by the process of mitochondrial autophagy or mitophagy (Elmore et al., 2004; Rodriguez-Enriquez et al., 2006).

IV. Conclusion

Quantitative confocal and multiphoton microscopy of the intracellular distribution of membrane-permeant cationic fluorophores provides a minimally perturbing means to measure dynamically both mitochondrial and plasmalemmal ÁÉ in cultured cells. A limiting factor is time resolution, since several seconds are required for cationic fluorophores to reestablish a new steady-state equilibrium after a change of ÁÉ. A further shortcoming is that mitochondria lose their potential-indicating fluorophores and become invisible in fluorescence images after depolarization. The latter problem can be overcome by coloading with TMRM and MTG. With coloading, ÁÉ can still be monitored by red TMRM

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fluorescence excited with green light, but additionally TMRM quenches green MTG fluorescence excited with blue light via a FRET mechanism. Consequently, when a mitochondrion depolarizes, TMRM is released, and the green fluorescence of covalently bound MTR recovers, which reveals depolarized mitochondria as individual green structures against a background of red-fluorescing polarized mitochondria. Unlike virtually any other technique, confocal and multiphoton microscopy permits nondestructive serial observation of the ÁÉ's of populations of cells and mitochondria. Live cell, three-dimensionally resolved confocal and multiphoton microscopy is now an indispensable tool for studying the mitochondrial function and pathophysiology.

Acknowledgments

We thank Insil Kim for mouse hepatocytes and Dr. Donald R. Menick for adult feline cardiac myocytes. This work was supported, in part, by Grants DK37034 and DK070195 from the National Institutes of Health.

References

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