Journal of Histochemistry and Cytochemistry, Vol. 45, 403-412, Copyright © 1997 by The Histochemical Society, Inc.


ARTICLE

Intracellular Accumulation of Rhodamine 110 in Single Living Cells

Valérie Jeannota, Jean-Marie Salmona, Michel Deumiéa, and Pierre Vialleta
a Microfluorimétrie Quantitative et Pharmacocinétique Cellulaire, Laboratoire de Chimie-Physique, Université de Perpignan, Perpignan, France

Correspondence to: Jean-Marie Salmon, Microfluorimétrie Quantitative et Pharmacocinétique Cellulaire, Lab. de Chimie-Physique, Univ. de Perpignan, 52 ave de Villeneuve, 66860 Perpignan cedex, France.


  Summary
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Summary
Introduction
Materials and Methods
Results
Discussion and Conclusion
Literature Cited

To gain a better understanding of the internalization of rhodamines, vital staining of living cells in situ by two different rhodamines, R110 and R123, was studied by microfluorometry. These dyes differ strongly in their lipophilic properties because of differences in charge distribution. Microspectrofluorometry was used to study the fluorescence emission spectra of R110-loaded cells to determine reliable loading conditions. Cell uptake and cell efflux studies of R110 were performed by numerical microfluorescence imaging. A slower uptake was observed for R110 (14 hr) vs R123 (2 hr), but the R110 efflux was much more rapid (30 min) than that of R123 (>24 hr). Although it appeared in the R110 and R123 co-localization study that R110 was able to accumulate in mitochondria, labeling with R110 was lower than with R123. Our results indicate that, rhodamine 110 in its acid cationic form is able to cross the plasma and mitochondrial membrane and to accumulate in cell compartments as does the cationic rhodamine 123. However, because of its acido-basic properties, R110 should be able to decrease the pH of cell compartments, depending on their ability to regulate pH. In such a model, mitochondrial pH should be more greatly decreased than cytosolic pH, leading to a lower mitochondrial accumulation of R110 than of R123. Surprisingly, these effects, which should affect the energetic state of mitochondria, do not influence cell growth, because no cytotoxic effect was observed. (J Histochem Cytochem 45:403-412, 1997)

Key Words: microspectrofluorometry, numerical image analysis, rhodamine 110, single living cells, intracellular localization


  Introduction
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Summary
Introduction
Materials and Methods
Results
Discussion and Conclusion
Literature Cited

Mitochondria play a vital role in the energy metabolism of virtually all living cells (Mehlman and Hanson 1972 ; Wainio 1970 ). These organelles undergo morphological changes in shape and distribution, which can be affected by the metabolic state, proliferation, differentiation, aging, and pathological events (Weakley 1976 ; Rosenbaum et al. 1969 ; Hakenbrouck 1968 , Hakenbrouck 1966 ; Packer 1963 ; Rouiller 1960 ). There have been many attempts to study the localization of these organelles with indicator dyes. Although isolated mitochondria and mitochondria of fixed cells have been extensively investigated, much less attention was paid to mitochondria of living cells until some vital stains that allowed specific staining of mitochondria became available, e.g., styryl probes such as DASPMI (Bereiter-Hahn 1976 ), amphipathic rhodamines such as R123 (Johnson et al. 1980 ), and carbocyanines such as DiOC5 (Johnson et al. 1981 ).

A study by Johnson et al. 1980 demonstrated that, at physiological pH, positively charged rhodamines, i.e., 123, 6G, and 3B, could specifically stain mitochondria. Conversely, zwitterionic rhodamines simultaneously bearing a positive charge on the xanthene ring and a negative charge on the carboxyl group on the isolated benzene ring, such as rhodamines 110, 116, and B, were reported to be unable to stain mitochondria. However, nothing was reported concerning the labeling of other cell compartments by these compounds. These results and others (Ehrenberg et al. 1988 ; Wagoner 1979 ) suggested that an attraction of the cationic rhodamine molecules by relatively high negative electric potentials, first across the plasma membrane and then across the mitochondrial membrane, may be the basis for selective staining of mitochondria by rhodamine 123 (R123) in living cells.

In their study Rashid and Horobin 1991 have shown that anionic and non-ionic fluorescent probes can also stain mitochondria and have addressed the question of the internalization process that allows mitochondrial accumulation of these probes. This is also relevant to the work of Lampidis et al. 1989 , who studied the influence on cellular accumulation of the net charge borne by rhodamine molecules. These authors showed (a) that positively charged R123 accumulates in the cells to a tenfold greater extent than the zwitterionic rhodamine 110 (R110) that did not carry a net charge, and (b) that R110 was 1000-fold less toxic than R123.

It should be noted that in these studies the evaluations of intracellular R110 and R123 accumulation were performed on butanolic extracts from the cells. To gain a better understanding of rhodamine internalization, it would be interesting to supplement the large number of studies on intracellular fluorescence distribution of R123 (Canitrot et al. 1993 ; Rashid and Horobin 1991 ; Farkas et al. 1989 ; Wagoner 1979 ) with a comparative study of the intracellular accumulation of R110.

Although R110 has been reported to accumulate to a tenfold lesser extent than R123 in living cells, it should be detectable by appropriately sensitive apparatus and procedures. For this purpose, fluorescent techniques that allow high temporal and spatial resolution are powerful methods for investigation of rhodamine interactions in cells and cellular organelles. As a first step, we studied by microspectrofluorometry the (steady-state) fluorescence emission spectra of intracellular R110 to delineate the interactions of the dye with its intracellular environment and to detect changes in the probe environment. In a second step, low light-level microfluorometry associated with quantitative image analysis was used to investigate the dynamics of R110 and R123, i.e., the R110 intake and efflux kinetics in the CCRF-CEM cell line, and to study the intracellular distribution of rhodamine fluorescence in these cells and in 3T3 fibroblasts.


  Materials and Methods
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Summary
Introduction
Materials and Methods
Results
Discussion and Conclusion
Literature Cited

Chemicals
Rhodamine 110 chloride (R110) was a laser grade dye from Eastman (Rochester, NY) Rhodamine 123 (R123) was obtained from Sigma (St-Quentin Fallavier, France). Stock solutions of 1 mM were prepared in ethanol (R110) and in PBS (R123). Both solutions were kept at -20C.

Cell Culture and Staining Procedures
Experiments were performed on 3T3 mouse fibroblasts (Flow; Cergy Pontoise, France) and human lymphoblastoid cells (CCRF-CEM cell line). CCRF-CEM cells were a gift from Dr. W. T. Beck (St Jude Children's Research Hospital; Memphis, TN). 3T3 and CCRF-CEM cells were routinely cultured in DMEM (Gibco BRL; Orsay, France) or RPMI 1640 (ICN; Orsay, France) medium, respectively, both supplemented with 2 mM glutamine (ICN) and 10% heat-inactivated fetal calf serum (Gibco). Antibiotics were added to the RPMI medium. The 3T3 fibroblasts were grown in 25-cm2 flasks and were resuspended in fresh culture medium. CCRF-CEM cells were seeded at 2 x 105 cells/ml every 3 days to maintain continuing exponential growth.

For experiments, 3T3 cells were seeded in Sykes-More chambers at 40,000-60,000 cells per chamber. After 48 hr, the cells were stained with R110 10 µM for 24 hr, and then were incubated with R110 10 µM for various periods of time. All experiments with CCRF-CEM cells were carried out when the density of the cell culture was between 106 cells/ml and 1.5 x 106 cells/ml, so that the cells were in the exponential phase of growth. Both kinds of cells were washed three times with PBS and resuspended in Hank's balanced salts solution (HBSS) (ICN) before intracellular fluorescence studies.

For probe efflux assays, the CCRF-CEM cells were incubated in complete culture medium with R110 10 µM for 24 hr, then washed and resuspended in HBSS. The fluorescence intensity of intracellular R110 was recorded at various times on the same cell samples at room temperature.

Fluorescence Microscopy
Fluorescence microscopy investigation was performed using low light levels of exciting radiations and high-sensitivity detection systems. These conditions enabled us to avoid cell damage and photodestruction of the probes. Furthermore, the use of computer-controled shutters enabled us to irradiate the samples only for the duration necessary to record spectra (3.3 sec) and images (0.4 sec).

Microspectrofluorometry on Single Cells. The microspectrofluorometer previously described (Allegre et al. 1985 ; Salmon and Viallet 1981 ) was composed of an inverted microscope (Leitz) connected to an optical multichannel analyzer (OMA) from Princeton Applied Research equipped with a silicon-intensified target (SIT) detector. The excitation wavelength at 464 nm was selected by a monochromator from the emission of a high-pressure xenon lamp. Spectral data were transferred to a PDP 11/73 microcomputer (Plessey) for storage and calculations. All fluorescence spectra records were performed during the 5-min period after rinsing of the cells.

Numerical Image Analysis. The fluorescence image cytometry system has been previously described (Lahmy et al. 1991 ; Vigo et al. 1991 ). In short, it uses an inverted fluorescence microscope (Olympus IMT2) equipped with an epi-illuminating system (x 40 and x 100 objectives from Leitz and final magnification of x 60 or x 150, respectively) in association with an SIT camera from Lhesa (LH 4036) and a TITN SAMBA 2005 image processor. The mercury line at 435 nm of a high-pressure mercury lamp (100 W), selected by a dichroic mirror, was used as fluorescence excitation.

Quantitative measurements were performed with a specific program that allows subtraction of the camera background and correction of the noise linearity of the digitizer and of the heterogeneity (pixel to pixel) of the gain of the camera (Vigo et al. 1991 ). A procedure was developed to record the fluorescence of the cells using image segmentation by thresholding. This was followed by opening and closing sequences ( Salmon et al. 1992 ) that resulted in smoothing of the cell contour. In addition, some specific parameters related to the mask of the cell (surface and shape factors) allowed exclusion of data related to subcellular debris or cell clumps. The main parameters obtained for each cell were cell size, shape factor, fluorescence intensity, mean fluorescence intensity, and standard deviation from R110 or R123 fluorescence. The SAMBA 2005 image processor that enabled recording of five or six corrected images in less than 2 min was used for study of the intake and efflux kinetics of R110 and of the intracellular localization of R123 and R110.


  Results
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Summary
Introduction
Materials and Methods
Results
Discussion and Conclusion
Literature Cited

Spectrofluorometric Study of R110 in Solution and in Cells
To study and compare the R110 fluorescence changes depending on its environment, we recorded the R110 fluorescence spectra in water solution, in complete culture medium (without phenol red but complemented with 10% v/v fetal calf serum), and in cells. The similarity of the fluorescence spectra of R110 in water and in culture medium indicated that, as observed previously (Canitrot et al. 1993 ; Salmon et al. 1992 ), there was no interaction between R110 and any components of the culture medium. Conversely, comparison of the fluorescence emission spectra of R110 in culture medium and in cells (Figure 1) showed that the fluorescence spectrum of intracellular R110 is slightly red-shifted (8 nm for the maximum). This red shift results more from a modification in the shape of the fluorescence spectrum (a significant decrease in the blue edge) than from a shift of the entire fluorescence spectrum. Both the low thickness of the cell and the expected low intracellular concentration of R110 preclude the shift from being related to any reabsorption of the fluorescence. The red shift therefore indicates a perturbation of the electronic state of R110, resulting from specific interactions of this molecule with its intracellular environment. Canitrot and al. (1993,1994) similarly observed that the fluorescence spectrum of intracellular R123 was clearly red-shifted (7 nm) as compared with the R123 spectrum obtained in water, although modification of its shape was very slight. The less important modification of the fluorescence emission spectrum obtained for intracellular R110 suggests that the perturbations of the R110 electronic state induced by the intracellular environment were less for R110 than for R123. Consequently, the interactions of R110 with the local cellular environment should be weaker than those observed for R123.



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Figure 1. Fluorescence emission spectra of the R110 rhodamine in water ({circ}), in culture medium ({bullet}), and in CCRF-CEM cells ({square}). The concentration of R110 in aqueous solutions was 3 µM. Cells were incubated with R110 10 µM for 24 hr. Both spectra were recorded with a microspectrofluorometer under an excitation wavelength of 464 nm. All spectra were normalized.

The influence of the intracellular R110 concentration on the fluorescence spectra has also been investigated on cells loaded with R110 25 µM. The fluorescence spectra of the R110-treated cells shown in Figure 2 illustrate the large red shift (8 nm) of the maximum and the widening of the fluorescence emission observed after increasing the R110 concentration. However, the intracellular fluorescence spectra obtained from cells loaded at 5 µM (data not shown) are similar to those observed with cells loaded at 10 µM. These results suggest that at high intracellular concentration the fluorescence arises from R110 aggregates (Deumié et al. 1995 ; Bunting et al. 1989 ; Valdez-Aguilera and Neckers 1989). To avoid the formation of intracellular aggregates, the cells were therefore incubated with R110 extracellular concentrations of less than 10 µM.



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Figure 2. Effect of R110 concentration in the incubation medium on the shape of the fluorescence spectrum of R110 in CCRF-CEM cells. Cells were loaded with R110 for 24 hr with 10 µM ({circ}) and 25 µM ({bullet}) dye concentrations in the culture medium. Both spectra were recorded with a microspectrofluorometer under a fixed excitation wavelength of 464 nm and were normalized.

Control of R110 Cytotoxicity
Experiments with R110 were performed on the two cell lines to assess its toxicity. The cytotoxicity of R110 at the doses used in the assay was checked on CCRF-CEM cells grown in complete culture medium containing R110 10 µM at 37 C with 5% CO2. At 72-hr later, the cells were Coulter-counted and the data were compared to those obtained with control cells grown under the same conditions but without R110. Cell growth and morphology were unchanged in the presence of R110. When the same observations were performed on 3T3 cells, neither morphology nor cell growth was affected by R110. It therefore appears that R110 is not cytotoxic for the cell strains studied in our experimental conditions. This may be related to the results reported by Lampidis et al. 1989 , who found R110 to be cytotoxic only at concentrations greater than 100 µM. Our assays on cells confirm the lack of R110 toxicity at concentrations up to 10 µM.

Kinetics of R110 Uptake by CCRF-CEM Cells
To assess the optimal incubation time necessary for the R110 to be taken up by cells, using digitized video fluorescence microscopy we recorded the time dependence of intracellular accumulations of R110 for different extracellular concentrations of dye. The main purpose of this study was to establish a reliable loading protocol and, according to the above results, we followed the uptake process for extracellular concentrations of 10 µM and 25 µM R110.

As shown in Figure 3, the cell uptake of R110 was exponential and then reached a plateau level, a phenomenon commonly observed for intracellular accumulation of xenobiotics (Lahmy et al. 1995 ) and for lipophilic cationic dyes (Canitrot 1994 ; Irion et al. 1993 ). For an R110 extracellular concentration of 10 µM, a plateau was reached after 14 hr, which remained stable after 28 hr of further incubation.



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Figure 3. Uptake kinetics and intracellular concentrations of R110 in CCRF-CEM cells for different concentrations of R110 in the incubation medium. An aliquot (400 ml) of cells in exponential growth (1 x 106 cells/ml) was incubated with R110 10 µM ({diamond}) and 25 µM ({diams}) for the time indicated. The respective fluorescence intensities were evaluated as described in Materials and Methods.

These results indicate that R110 is able to cross the plasma membrane and to accumulate inside the cells. Furthermore, this plateau might be caused by interaction of R110 with a limited number of specific sites, as observed for labeling of DNA by Hoescht 33342 (Lahmy et al. 1995 ) or, alternatively, might correspond to an intracellular accumulation of R110 until equilibrium between the chemical potentials of intra- and extracellular R110 is reached. As shown by the fluorescence emission spectra, R110 interacts more strongly with intracellular chemicals than with the extracellular medium. Therefore, we should observe an intracellular accumulation.

When the extracellular R110 concentration was set at 25 µM, we observed a very rapid (<10-hr) internalization of R110, associated with a higher plateau level. This indicates that binding of R110 to a limited number of specific sites was not the process used, because in that case we should reach an identical plateau level no matter what the concentration of extracellular dye (Lahmy et al. 1995 ). Furthermore, the absence of proportionality between the plateau level and the R110 extracellular concentration (concentration x 2.5 vs intensity x 1.8) could be related to the presence of some aggregated R110 forms leading to high intracellular accumulations. It is well known that, in addition to the red shift of their fluorescence emission spectra, these aggregated dye species exhibit lower quantum fluorescence yields (Deumié et al. 1995 ; Deumié and El Baraka 1993 ) than do monomeric species. This explains why the level of the plateau was not directly proportional to the extracellular R110 concentration. In consequence, our incubation protocol will routinely use an incubation time of 14 hr in culture medium at 10 µM.

Kinetics of R110 Elimination from CCRF-CEM Cells
Once the level of the intracellular R110 had reached the plateau, we studied the R110 efflux to determine whether this plateau is due to strong binding to intracellular macromolecules or to an equilibrium between internalization and elimination. R110 elimination kinetics were investigated in CCRF-CEM cells at room temperature. After rinsing of the cells, the fluorescence intensity of the R110 intracellular fluorescence was recorded vs time in the same cell population (Figure 4). After 30 min, the fluorescence intensity returned to the level of the intrinsic cell fluorescence, presumably due, under the excitation conditions used, to flavin ring emissions (flavoproteins, flavin mononucleotides, flavin dinucleotides) (Salmon et al. 1982 ). The rate of clearance of the R110 zwitterionic species is short, because the experiment demonstrated that R110 elimination from the CCRF-CEM cells was complete within 30 min. Therefore, the speed of the efflux kinetics justified our use of equipment that enabled rapid recording of the fluorescence spectra and of the cellular fluorescence images.



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Figure 4. Efflux kinetics of R110 from CCRF-CEM ({blacksquare}) cells. An aliquot (400 ml) of cells in exponential growth (1 x 106 cells/ml) was loaded with R110 10 µM for 14 hr. Cells were then washed and resuspended at 1 x 106 cells/ml in HBSS. ({circ}) represents the evolution with time of the intrinsic fluorescence [NAD(P)H] of control cells under the same experimental conditions. The fluorescence intensity of R110 and NAD(P)H was quantified as described in Materials and Methods.

Study of the Intracellular Localization of R110
Before we discuss the intracellular localization of R110 and R123, two points should be made. First, at the doses used, no toxic effects were detected (this article) or reported (Canitrot et al. 1993 ; Lampidis et al. 1989 ). Second, the conditions used for image recording (low light excitation levels) allow elimination of the probe redistribution usually observed after the mitochondrial explosion caused by intense illumination. Therefore, the study of the intracellular localization of R110 is performed only in healthy cells.

Comparative Localization of Rhodamines in CCRF-CEM Cells. Drug distribution in cells depends on a variety of parameters, including the chemical properties of the exogenous agent, i.e., its hydrophobic or hydrophilic nature and its state of ionization. The distribution of intracellular fluorescence in CCRF-CEM cells incubated with R110 10 µM for 14 hr was studied by videomicrofluorimetry. The distribution pattern is shown in Figure 5A. Depending on the direction from which the cell was observed, we obtained more or less heterogeneous distributions of intracellular fluorescence, usually with a granular appearance. These observations are more clearly shown on the fluorescence profiles in Figure 6. The R110 profile shows an abrupt rise in intensity of the cellular fluorescence, which corresponds to the edge of the cell. The surface of an R110-incubated cell is viewed as an undulating plateau, with distortions and short spikes that might correspond to the granular aspect of the intracellular fluorescence. This profile was very different from that obtained with uniformly stained spherical objects (fluorescent beads), giving a Gaussian-like distribution (data not shown). This result indicates a rather heterogeneous distribution of the R110 intracellular fluorescence.



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Figure 5. (A) CCRF-CEM cells stained with R123 (10 µM) for 1 hr. The excitation wavelength was 435 nm. Neutral filters were used so that the excitation intensity was equal to 5% of the lamp intensity. Distance C to D = 26 µm. (B) CCRF-CEM cells stained with R110 (10 µM) for 14 hr. The excitation wavelength was 435 nm. Neutral filters were used so that the excitation intensity was equal to 12.5% of the lamp intensity. Distance C to D = 31 µm.



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Figure 6. Fluorescence profile of CCRF-CEM cells stained with R123 along the segment C-D = 26 µm (top) and R110 along the segment C-D = 31 µm (bottom). These fluorescence profiles were obtained as described in Materials and Methods.

When CCRF-CEM cells were stained for comparison with R123, which is known to accumulate specifically in the mitochondria of living cells, typical fluorescence patterns, with some heterogeneous intracellular distribution (Figure 5B), were observed. In the R123-treated cells some very bright spots appeared, and the cell contour was less clearly defined than in R110-treated cells. However, as shown in Figure 6, rather similar fluorescence profiles were observed for R110 and R123. These results suggest that R110 is at least partially localized to the same intracellular compartments as R123, i.e., to the mitochondria.

Successive Staining of 3T3 Fibroblasts by R110 and R123: Co-localization Studies. To confirm the above results obtained on spherical CCRF-CEM cells, we studied the intracellular localization of R110 on 3T3 adherent cells. 3T3 fibroblasts, which grow spread on the culture support, provide a suitable model for convenient observation of the intracellular compartments. Then, to study the intracellular localization, we have compared the respective co-localizations, of the R110 and R123 dyes on the same cells. After being seeded in a Sykes-More chamber, the cells were stained with R110 µM and their images recorded. The cells were then rinsed in HBSS without moving the Sykes-More chamber tightly bound to the microscope stage. After 30 min to allow the R110 efflux to be achieved, the cells were incubated for 15 min at room temperature in complete culture medium with R123 25 µM. Finally without moving the Petri dish, the cells were rinsed again with HBBS and images of the same cells were recorded.

The result of this double staining procedure is shown in Figure 7. Although the contrast was higher for R123-treated (Figure 7B) than for R110-treated (Figure 7A) cells, both cell types exhibited some heterogeneity in fluorescence distribution patterns, with some specific bright spots that were presumed to represent the same intracellular localization. Because of the reported properties of R123 (Shapiro 1991 ; Farkas et al. 1989 ; Johnson et al. 1981 ) and the granular fluorescence distribution spread over the entire cell, it is highly unlikely that this distribution can be correlated with the labeling pattern of endoplasmic reticulum or Golgi apparatus by xenobiotics (Kohen et al. 1989 ). Therefore, these distributions, particularly the distribution observed for R123, represent important perinuclear fluorescence that may originate from superimposition of perinuclear clouds of mitochondria in this thicker region of the cell, in which mitochondria are quite crowded. For comparison of the distributions and the localization of the bright spots shown in Figure 7A and Figure 7B, the fluorescence profiles of the cells stained with R110 and with R123 and obtained along the same line are shown in Figure 8. The R123 fluorescence profile is serrated, which means that there are wide variations in the distribution of intra-cellular fluorescence. This observation is in agreement with the mitochondrial localization of R123. To a lesser extent, R110 also presents a serrated profile. In addition, there is a large spike corresponding to that of the R123 profile. Other similarities can also be observed between the lower spikes of the two profiles. Although a marked spike in the R123 profile is absent from that of R110, these observations suggest that R110 is at least partially localized to the mitochondria.



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Figure 7. (A) 3T3 cells stained with R110 (10 µM) for 14 hr. The excitation wavelength was 435 nm. Neutral filters were used so that the excitation intensity was equal to 25% of the lamp intensity. Distance C to D = 88 µm. (B) The same cells as in A stained afterwards with R123 (25 µM) for 15 min. The excitation wavelength was 435 nm. Neutral filters were used so that the excitation intensity was equal to 5% of the lamp intensity. Distance C to D = 88 µm.



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Figure 8. Fluorescence profiles of the 3T3 cell, respectively stained with R110 and R123, along the same segment from C to D in Figure 7A and Figure 7B. These fluorescence profiles were obtained as described in Materials and Methods.

Figure 7A and Figure 7B show a large difference in the labeling of the nuclear area. Whereas for R123 it appeared as a dark hole, for R110 the fluorescence intensity of this area is about two thirds the maximum of the intensity profile. This finding suggests that R110 is distributed throughout the entire intracellular space, including the nucleus. In addition, the picture of R123-treated 3T3 cells was recorded with an excitation intensity one fifth of that used for R110. Therefore, it is clear that with a fivefold increased excitation intensity that should enhance the fluorescence by a factor of 5, the nuclear area of R123-treated cells should reach a level close to that of the R110-treated cells. The main difference in the two stainings may therefore arise from the much greater accumulation of R123 in mitochondria than in the cytosol or nucleus. With an experimental excitation light reduced by a factor of 5 compared to that for R110, detection of the low level of R123 cytosolic fluorescence became impossible; the intracellular fluorescence that mainly originated from mitochondria appeared as spots.

Although slight concentrations of R110 appear to be able to accumulate in mitochondria, the gradient of that accumulation between the intracellular space and the mitochondria is much greater for R123 than for R110.


  Discussion and Conclusion
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Summary
Introduction
Materials and Methods
Results
Discussion and Conclusion
Literature Cited

Although it has previously been stated that rhodamines and other similar compounds simultaneously bearing a positive and a negative charge are unable to accumulate in cells (owing to the absence of membrane potential driving forces) (Ehrenberg et al. 1988 ; Wagoner 1979 ), an intracellular accumulation of R110 that supports some previous studies (Rashid and Horobin 1991 ; Lampidis et al. 1989 ) was observed. The fluorescence emission spectrum of intracellular R110 loaded into cells and recorded by microspectrofluorometry shows a slight red shift and shape modification compared to the spectrum observed in culture medium. These changes, related to environmental differences, indicate slightly stronger interactions of intracellular R110 in situ compared to the interactions in culture medium. In addition identical fluorescence spectra were obtained for R110 loaded into cells for concentrations up to 10 µM, indicating a similar R110 intracellular environment for these dye concentrations. However, this was not the case when the cells were treated with R110 25 µM culture medium. The staining resulting from this greatest concentration revealed an important red shift, which might be interpreted as arising from the formation of R110 aggregates. Because cytotoxic effects were not observed at extracellular R110 concentrations of 10 µM, and to avoid the possibility of aggregate formation, that dose appeared appropriate for a convenient cell loading.

Our investigation of R110 uptake and efflux has shown that the R110 uptake is exponential and reaches a plateau after 14 hr of incubation. That plateau remains stable for the following 28 hr at least. Increasing the extracellular R110 concentration to 25 µM also leads to a plateau, but its level is not proportional to the extracellular concentration used. This presumably can be attributed to the lower quantum yields of R110 aggregates (Deumié and El Baraka 1993 ; Kelkar et al. 1990 ; Bunting et al. 1989 ) and indicates that, as for R123 (Canitrot 1994 ), there is no binding of R110 to a limited number of identical binding sites. In such a case we should have obtained the same plateau level regardless of the extracellular R110 concentration.

The existence of a driving force, resulting from the negative intracellular and intramitochondrial electric potentials, and the ability of the R110 molecule to cross the plasma and mitochondrial membranes are obviously parameters that may control the uptake of R110. However, it remains to be explained how a negatively charged molecule, presumably unable to accumulate inside the cells (Ehrenberg et al. 1988 ; Wagoner 1979 ), can cross the plasma membrane. As did Rashid and Horobin 1991 to explain the mitochondrial accumulation of analogous compounds, let us consider the acido-basic equilibrium of R110 at the carboxylic site:

With a pKa of around 4.3, at the physiological pH of 7.2 we have only 1/1000 of R110 molecules in the protonated acidic form bearing a positive charge in the extracellular medium. In this form, the R110 should be able, as is R123, to cross the plasma and mitochondrial membranes. In the aqueous compartments inside the cells, the acido-basic equilibrium of R110 is reestablished. Such multiple equilibria are dependent on the pH of the compartments. The higher the pH of the compartment, the more R- zwitterionic form of R110 is present. Therefore, we should expect, for the R110 acid form RH, a distribution in the compartments according to the driving forces, similar to that for R123. Alternatively, the amount of the R110 basic form R- should depend on the amount of RH and on the pH of each compartment. However, the pH of these two compartments, cytosol and mitochondria, should also reflect the capacities of each compartment to regulate the buffer effect of the acido-basic equilibrium of R110. In preliminary studies using the pH probe C-SNARF-1, a pH around 6.3 was found for the mean intracellular pH of R110-treated CCRF-CEM cells (data not shown), whereas control cells presented a pH of around 7.2 (Vigo et al. 1995 ). This mean value of pH indicates that compartments can be at pH values higher or lower than 6.3, depending on their capacity to regulate the pH. Therefore, owing to the poor pH regulation capacities, which are necessary to their function (Stryer 1988 ), mitochondria should exhibit an intramitochondrial pH of less than 6.3, and the cytosolic pH should be maintained at higher values by the pH regulation mechanism. Because the mitochondrial membrane potential results from a pH gradient across the mitochondrial membrane (Stryer 1988 ) (higher pH inside than in the cytosol) a decrease in mitochondrial pH relative to cytosolic pH should cause a decrease in the mitochondrial membrane potential and consequently a decrease in the driving force at the origin of R110 mitochondrial accumulation.

Although labeling the same compartments, R123 and R110 may therefore exhibit differences in their relative accumulation levels in cytosol and mitochondria, i.e., lower mitochondrial accumulation of R110 (RH) than R123 owing to a decrease in the driving force related to lowering of mitochondrial pH. This model is in agreement with the observed reduced mitochondrial accumulation of R110 compared to that of R123.

According to this model, the slowness of the intake process of R110 could be explained by the very low amount of R110 (1/1000 of total R110) in its acid form, RH, in the extracellular medium. In a similar way, the efflux results from lowering of the R110 extracellular chemical potential that follows rinsing of the cells. The efflux tends to equilibrate intracellular and extracellular R110 chemical potentials. Because of the high dilution factor (103) resulting from the differences between the total cell volume and the volume of the extracellular medium, the intracellular level of R110 was too low to be detected at the end of the efflux when the new equilibrium was reached. However, the fast efflux observed--30 min to reach the new equilibrium--compared to the intake suggests that the trans-membrane diffusion of RH was more efficient on the way out than on the way in. This swiftness may be due to a relative increase in the intracellular R110 concentration under its acid form, RH. This hypothesis is in agreement with the mean pH value, approximately 6.3, for R110-treated cells. At this pH we should have 1% of the intracellular R110 in its acidic form, RH, which is 10 times greater than what we have in the extracellular medium at pH 7.2.

Finally, this slow intake and rapid efflux might also explain the usual failure to observe the intracellular accumulation of R110. Consequently, the R110 loading protocol should use a time long enough to reach a plateau for the intracellular R110 fluorescence; 14 hours was the time required to achieve easy and reliable loading of the cells. In addition, the rapid efflux of R110 from cells induces experimental constraints of very fast rinsing and data recording. It is also essential to use the same schedule and procedure to study the cell populations in the same state of R110 loading.

In conclusion, these results indicate that, in its acid cationic form, rhodamine 110 can cross the plasma and mitochondrial membrane and can accumulate in cell compartments in the sameway as cationic rhodamine 123. However, because of its acido-basic properties, R110 should be able to decrease the pH of cell compartments depending on their capability to regulate pH. In such a model, mitochondrial pH should be decreased to a greater extent than cytosolic pH, thus leading to lower mitochondrial accumulation of R110 than of R123. Surprisingly, these effects, which should affect the energetic state of mitochondria, do not influence cell growth, because no cytotoxicity was observed. This suggests an efficient ATP production at the cytosolic level through the glycolytic pathway. To confirm these conclusions, we plan to investigate more precisely the effect of R110 in two directions: first, by studying the pH of different cell compartments with the use of fluorescent probes and videomicrofluorometry (Stryer 1988 ), and second, by investigating the production of NADH by the glycolytic pathway to evaluate the capacity of cells to compensate for a decrease in the mitochondrial energetic state resulting from a decrease in intramitochondrial pH.


  Acknowledgments

This work was supported by the Comité Départemental des Pyrénées Orientales de la Ligue Contre le Cancer.

We thank Ms Elsa Matzner for help in writing this manuscript.

Received for publication October 28, 1996; accepted October 30, 1996.


  Literature Cited
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Summary
Introduction
Materials and Methods
Results
Discussion and Conclusion
Literature Cited

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