Interrelationships of endoplasmic reticulum, mitochondria, intermediate filaments, and microtubules--a quadruple fluorescence labeling study.

Interrelationships of endoplasmic reticulum, mitochondria, intermediate filaments, and microtubules-a quadruple fluorescence labeling study BOHDANJ. SOLTYS AND RADHEYS . GUPTA' Department of Biochemistry, McMaster University, Hamilton, Ont., Canada L8N 32.5 Received March 17, 1992

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SOLTYS,B. J., and GUPTA,R. S. 1992. Interrelationships of endoplasmic reticulum, mitochondria, intermediate filaments, and microtubules-a quadruple fluorescence labeling study. Biochem. Cell Biol. 70: 1174-1 186. To study the interrelationships of endoplasmic reticulum, mitochondria, intermediate filaments, and microtubules, we have developed a quadruple fluorescence labeling procedure to visualize all four structures in the same cell. We applied this approach to study cellular organization in control cells and in cells treated with the microtubule drugs vinblastine or taxol. Endoplasmic reticulum was visualized by staining glutaraldehyde-fixed cells with the dye 3,3'-dihexyloxacarbocyanineiodide. After detergent permeabilization, triple immunofluorescence was carried out to specifically visualize mitochondria, vimentin intermediate filaments, and microtubules. Mitochondria in human fibroblasts were found to be highly elongated tubular structures (lengths up to greater than 50 pm), which in many cases were apparently fused to each other. Mitochondria were always observed to be associated with endoplasmic reticulum, although endoplasmic reticulum also existed independently. Intermediate filament distribution could not completely account for endoplasmic reticulum or mitochondria1 distributions. Microtubules, however, always codistributed with these organelles. Microtubule depolymerization in vinblastine treated cells resulted in coaggregation of endoplasmic reticulum and mitochondria, and in the collapse of intermediate filaments. The spatial distributions of organelles compared with intermediate filaments were not identical, indicating that attachment of organelles to intermediate filaments was not responsible for organelle aggregation. Mitochondrial associations with endoplasmic reticulum, on the other hand, were retained, indicating this association was stable regardless of endoplasmic reticulum form or microtubules. In taxoltreated cells, endoplasmic reticulum, mitochondria, and intermediate filaments were all associated with taxol- stabilized microtubule bundles. Key words: endoplasmic reticulum, mitochondria, intermediate filaments, microtubules. SOLTYS,B. J., et GUPTA, R. S. 1992. Interrelationships of endoplasmic reticulum, mitochondria, intermediate filaments, and microtubules-a quadruple fluorescence labeling study. Biochem. Cell Biol. 70 : 1174-1186. Pour Ctudier les relations entre le reticulum endoplasmique, les mitochondries, les filaments intermediaires et les microtubules, nous avons ddveloppe une technique de marquage quadruple par fluorescence pour visualier les quatre structures dans la m&mecellule. Nous utilisons cette approche pour Ctudier l'organisation cellulaire dans les cellules contrdles et les cellules traitkes avec la vinblastine ou le taxol, drogues qui agissent sur les microtubules. Le reticulum endoplasmique est rendu visible par coloration des cellules fixCes au glutaraldihyde avec l'iodure de 3,3 '-dihexyloxacarbocyanine. Aprbs permkabilisation avec un detergent, nous prockdons a une triple immunofluorescence pour visualier specifiquement les mitochondries, les filaments intermediaires de vimentine et les microtubules. Dans les fibroblastes humains, les mitochondries sont des structures tubulaires trks allongees (d'une longueur de plus de 50 pm) et dans plusieurs cas, elles semblent fusionntes les unes aux autres. Les mitochondries sont toujours vues associkes au reticulum endoplasmique bien que le reticulum endoplasmique puisse Cgalement exister de f a ~ o nindependante. La distribution des filaments intermtdiaires ne peut rendre compte complttement des distributions du reticulum endoplasmique ou des mitochondries. Cependant, les microtubules sont toujours distribuks avec ces organites. La dCpolymCrisation des microtubules dans les cellules traitkes avec la vinblastine entraine la co-agregation du rCticulum endoplasmique et des mitochondries et le collapsus des filaments intermediaires. Les distributions spatiales des organites comparks aux intermediaires ne sont pas identiques, preuve que l'attachement des organites aux filaments intermtdiaires n'est pas responsable de l'agregation des organites. En revanche, l'association des mitochondries au reticulum endoplasmique est retenue, preuve que cette association est stable quelle que soit la forme du reticulum endoplasmique ou des microtubules. Dans les cellules traitkes au taxol, le reticulum endoplasmique, les mitochondries et les filaments intermtdiaires sont tous associks aux faisceaux de microtubules stabilisks par le taxol. Mots elks : rrCticulum endoplasmique, mitochondries, filaments intermediaires, microtubules. [Traduit par la redaction]

Introduction The

organization o f organelles is widely recognized t o arise from associations with the cytoskeleton (e-g.9 Hyams a n d Stebbings 1979). Rapid redistributions o f organelles along polymer tracks can function in effecting ABBREVIATIONS: MTs, microtubules; ER, endoplasmic reticulum; IFs, intermediate filaments; CHO, Chinese hamster ovary; (YMEM, alpha minimal essential medium; EM, electron microscopy; DiOC6(3), 3,3'-dihexyloxacarbocyanine iodide; PBS, phosphate-buffered saline; SDS, sodium dodecyl sulfate; I ~ G , immunoglobulin G; HSP60, 60-kilodalton heat shock protein; NP-40, Nonidet P-40. ' ~ u t h o rto whom all correspondence should be addressed. Printed in Canada / Imprime au Canada

o r responding t o local changes i n cell physiology. T h e association o f the organelle with t h e cytoskeleton, however, results in more than just organelle transportation. changes in actual organelle form caused by the physical linkage to the cytoskeleton p o t e n t i d y modify organelle activity. There may also b e t h e indirect effects, arising from bringing two organelles in apposition with each other, that result in new

properties or interrelationships among organelles. These concepts are well illustrated i n t h e case of the Golgi apparatus. The role of MTs in ~ o lorganization ~ i has been reviewed by Kreis (1990) a n d Kelly (1990). I n this study we were interested in evaluating interrelationships among E R , mitochondria, IFs, a n d MTs. There


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is evidence that mitochondria associate with ER (see Bereiter-Hahn 1990). The extension of ER networks along MTs has recently been demonstrated (Terasaki et al. 1986; Dabora and Sheetz 1988; Lee et al. 1989). There is good evidence that mitochondria associate with MTs and that MTs have a role in their cellular distribution (e.g., Smith et al. 1975; Heggeness et al. 1978; Wang and Goldman 1978; Ball and Singer 1982; Summerhayes et al. 1983; Price et al. 1991). In contrast, the relationship of IFs to organelle distribution is not clear. Studies with different cell types or in cells treated with different agents that alter the cytoskeleton have suggested that IFs may play an auxiliary, and sometimes exclusive, role in determining mitochondrial distributions (e.g., Wang and Goldman 1978; Mose-Larsen et al. 1982; Summerhayes et al. 1983; Chen et al. 1984; Eckert 1986; Cambray-Deakin et al. 1988; Chen 1988; Shyy et al. 1989; Dudani et al. 1990; Stromer and Bendayan 1990). We chose to reevaluate the cellular distribution of these structures by visualizing all four components in the same cell to gain new insights into their interrelationships. We report here the necessary methodology to do this with a standard fluorescence microscope. Included in this is a procedure for overcoming the common problem of glutaraldehyde-fixed antigens that antibodies often no longer recognize. We evaluate the form and distributions of these organelles in relation with the cytoskeleton in unperturbed cells and in cells in which the assembly state of MTs is altered with antimitotic drugs. We present evidence that in the absence of MTs it is ER-mitochondrial associations, and not IF-mitochondria1 associations, that determine mitochondrial distributions. Materials and methods Cell culture The origins of human embryonic lung fibroblasts (HSC172) and CHO-K, cell line (strain DR31) were described earlier (Buchwald and Ingles 1976; Gupta et al. 1978). Cells were grown in aMEM medium, supplemented with 5% fetal calf serum at 37°C in a humidified atmosphere of 5% CO, and 95% air. Human fibroblasts for correlative dye labeling and immunofluorescence microscopy were grown on cover slips, on which a carbon grid pattern had been laid to provide coordinates for relocating cells (Soltys and Borisy 1985). Briefly, these cover slips were prepared by carbon evaporation through Pinpointer or Coordinate EM grids, removal of the grids, and baking the cover slips at 400°C for 1 h. Cells for experimentation were grown for 2 days or more on cover slips and only sparse to subconfluent cultures were used. Vinblastine sulfate and tax01 were obtained from sources described in earlier studies (Gupta et al. 1982). Stock solutions of these drugs were at 10 and 2 mg/mL in ethanol and dimethyl sulfoxide, respectively. Drugs were diluted into aMEM supplemented with 5% fetal calf serum and then added to cells. The final concentrations of the organic solvents used during cell treatment were less than 0.7% dimethyl sulfoxide or 0.02% ethanol, neither of which on their own affected cellular organization. Drug treatment was carried out for the indicated period of time at 37OC in a humidified atmosphere of 5% CO, and 95% air. Dye staining ER and mitochondria were visualized with the cyanine dye, DiOC6(3) (Sigma Chemical Co., St. Louis, Mo.) using a variation of the procedure of Terasaki et al. (1984). The main change was the increased glutaraldehyde fixation of cells. Stock solutions of DiOC,(3) were 2.5 mg/mL in ethanol and were stored at -40°C.

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Working dye solutions were prepared immediately before use and made fresh for each cover slip of cells. Cells were rinsed briefly in 0.1 M sucrose - 0.1 M cacodylate (pH 7.4) (sucrose-cacodylate buffer) and then fixed in 0.5% glutaraldehyde in sucrosecacodylate buffer for 10 min. After a brief rinse with a large excess of sucrose-cacodylate buffer, cells were stained for 30 s with 5 pg/mL of DiOC6(3) in sucrose-cacodylate buffer, rinsed twice in sucrose-cacodylate buffer, and then mounted in the same buffer on a slide. Spacers were positioned between the cover slip and the slide to prevent cell damage, especially during cover-slip removal after photography. Cover slips were secured in position with nail polish. Cells were examined with an Olympus BHA microscope equipped with epifluorescence accessories by use of a 100 x UVFL objective (N.A. 1.3) and fluorescein optics. Cells were photographed on T-Max 400 (Eastman Kodak Co., Rochester, N.Y.) within 15 min after dye staining. Exposure times were 5-10 s and the film was developed at an exposure index of 400.

Immunoj7uorescent labeling Glutaraldehyde-fixed cells, prepared as above, were rinsed extensively with PBS (containing per litre, 8 g NaCI, 0.2 g KCl, 1.15 g Na2HP0,, and 0.085 g KH,PO,, pH 7.2) and then permeabilized with 0.5% SDS in PBS for 20 min, with mild movement on a platform rotator. This treatment also completely removed DiOC6(3) dye staining. Cells were then rinsed with PBS and treated with three changes of 10 mg/mL of sodium borohydride in H,O (Soltys and Borisy 1985) for a total of 5 min, to quench unreacted glutaraldehyde and to prevent glutaraldehyde-induced autofluorescence. Cells were then rinsed extensively with PBS and labeled with antibodies. For triple immunofluorescent labeling, the labeling sequence was as follows. All antibody binding reactions were at 37°C for 45 min. Further, since all secondary antibodies used here were goat antibodies, normal goat serum (20%) was used as a carrier for all antibodies to prevent nonspecific reactions. Between each reaction, cells were washed with two changes of PBS at room temperature. (i) First, cells were treated with 1:40 dilution of rabbit anti-vimentin antiserum (a generous gift of Dr. E.H. Ball of the University of Western Ontario, London, Ont.). The secondary antibody employed in this case was a 1:40 dilution of fluorescein conjugated goat anti-rabbit IgG (Jackson Immuno Research, West Grove, Pa.). (ii)Next, cells were treated with mouse monoclonal antibody to human HSP60 (PI protein) which provides a mitochondrial marker (Gupta and Austin 1987; Gupta and Dudani 1987). The characterization of this antibody which was used at 1:40 dilution is reported elsewhere (Singh and Gupta 1992). The secondary antibody used in this case was a 1:20 dilution of Cascade blue conjugated goat anti-mouse IgG (Molecular Probes, Junction City, Oreg.). (iii) Finally, the cells were labeled with a 1:40 dilution of monoclonal anti-tubulin (YL 1/2) antibody (Accurate Chemical and Scientific, Westbury, N.Y.), followed by treatment with 1:40 dilution of Texas red conjugated goat anti-rat IgG (Jackson Immuno Research, West Grove, Pa.). One or more of the above steps were omitted in certain montages of cells. When only immunofluorescence for mitochondria was done, HSP60 labeling was as above except that the secondary antibody was a 1:40 dilution of either fluorescein conjugated or Texas red conjugated antibody to mouse IgG (Jackson Immuno Research, West Grove, Pa.). Cells were mounted in medium containing 1 mg/mL of p-phenylenediamine (an anti-oxidant to minimize photobleaching), 90% glycerol, and 50 mM Tris-HCl (pH 8.0) and examined with an Olympus BHA microscope. Cascade blue mitochondria staining was observed in the UV channel (UG-1 exciter filter). Texas red MT staining was observed in the rhodamine channel. Cells were photographed as described above with the exception that in certain low magnification micrographs (see figure legends) a 40 x UVFL objective (N.A. 1.3) was used.

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F I G . 1 . Staining of ER and mitochondria in glutaraldehyde-fixed cells using the cyanine dye DiOC,(3). (a) Human fibroblasts. Large arrow, right side, points to a strongly fluorescent tubular mitochondrion. Small arrow, left side, indicates a region of interconnected tubular ER, which stains less strongly. ( b ) CHO-K, cells (strain DR31). Arrow points to an ER cisterna or membrane island. In the more general region of the cell process, mitochondria are found associated with ER cisternae. The bar in b represents 10 pm.

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FIG. 2. Immun~flu~rescent visualization of mitochondria and the cytoskeleton in human fibroblasts. Cells were fixed with glutaraldehyde and then permeabilized using SDS.A 40 x objective was used. (a) HSP60 antibody labeling of mitochondria. The bar represents 50 pm. (b) Labeling of IFs using vimetin antibody (fluorescein channel). The bar represents 30 pm. (c) Labeling of MTs in the same cells as in b using tubulin antibody (rhodamine channel). Results Dye labeling of endoplasmic reticulum and mitochondria Simultaneous visualization of both mitochondria and ER can be obtained by dye labeling in either living or fixed cells using the fluorescent cyanine dye DiOC6(3) (Terasaki et al. 1984). Figures l a and 1b show comparative DiOC6(3)

staining of glutaraldehyde-fixed human fibroblasts and CHO (DR31) cells, respectively. The strongly fluorescent tubular structures are mitochondria, one of which is indicated in Fig. l a (large arrow, right side). Their identity will be uniquivocally established by correlative immunofluorescent labeling in results described below. The ER stains


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FIG. 3. Correlative dye staining and HSP60 antibody labeling of mitochondria. (a) DiOC6(3) staining of glutaraldehyde-fixed human fibroblasts (fluorescein channel). (b) Immunofluorescent labeling of mitochondria in the same cell using HSP60 antibody (rhodamine channel). Arrows indicate the two ends of a single mitochondrion, identifiable in both channels. The bar in b represents 10 pm. less strongly. Radiating outward from the cell centre towards the cell periphery, the ER is composed of (i) interconnected tubules and (ii) sheet-like cisternae or membrane islands. ER tubules are particularly well observed in human fibroblasts (Fig. la, e.g., small arrow), the cell type used for most of the work described in this study. For comparative purposes, the ER in the CHO cell shown in Fig. l b exhibits both large accumulations of cisternae, containing the majority of mitochondria (e.g., region indicated by arrow), as well as tubular ER. Thus, different cells or cell types could differ in relative proportions of ER cisternae versus tubules. From the results shown in Fig. 1, it is evident that while mitochondria are regularly associated with ER, ER also exists independently. Based on our observations and photography of these structures in > 600 cells, a qualitative observation that could be made is that subcellular distribution of mitochondrial mass generally shows a good correlation with the subcellular distribution of ER mass. Mitochondria1 and cytoskeletal immunofluorescence Except in thinly spread peripheral regions of cells, superimposition of DiOC6(3)-labeled structures impairs or prevents identification of individual organelles. For this reason, in previous studies of ER network formation using DiOC6(3) staining (Terasaki et al. 1986; Lee and Chen 1988; Lee et al. 1989) observations were not made on

mitochondrial distributions. Mitochondria can be better identified by immunofluorescence labeling using a monoclonal antibody specific for HSP60 (PI protein), a protein localized in the mitochondrial matrix (Gupta and Austin 1987; Gupta and Dudani 1987). We encountered difficulty at first in labeling mitochondria in cells that had been glutaraldehyde fixed. We tried a variety of detergents as permeabilizing agents (e.g., Triton X-100, NP-40, deoxycholate, saponin) and none were effective in rendering the mitochondrial HSP60 epitope accessible to antibody (results not shown). We solved this problem by using the ionic detergent SDS to permeabilize cells. All immunofluorescence in this study was with cells treated in this way. The use of glutaraldehyde as a fixative was sufficient to stabilize the structures studied (see also Discussion). Figure 2a shows anti-HSP 60 staining of human fibroblasts. Immunofluorescent labeling allows individual mitochondria to be clearly identified throughout the cytoplasm, except where they are too numerous to be resolved. The cell shown is an example of senescent human fibroblasts which exhibit increased cell mass and a higher number of mitochondria. As seen, mitochondria can assume strikingly ordered arrays and appear cytoskeletal-like, reflective of underlying associations. Further, mitochondria are well preserved as tubular structures. Figures 2b and 2c show double-label immunofluorescence of IFs and MTs, respectively, in human fibroblasts of more normal dimensions and

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proliferative state. The cytoskeletal arrays display their


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In evaluating interrelationships of ER and mitochondria, we used correlative DiOC6(3) staining and anti-HSP60 mitochondrial labeling. Figure 3a shows DiOC6(3) staining of a slender process at a cell periphery. Although mitochondria can be identified, ER staining is intense and in several focal planes, preventing observation of ER substructure. This type of ER staining will be referred to as ER membrane accumulations or ER aggregates, not ER cisternae. The term ER cisternae will be reserved for where the ER clearly forms flat membrane island@)or peninsulas. Experimentally, after photographing the DiOC6(3) image, cells were detergent treated to completely remove the DiOC6(3) stain and to permeabilize the cell for antibody labeling. Mitochondria were always visualized in a different fluorescence channel from the original DiOC6(3) stain. Cells were relocated after antibody labeling using cover-slip coordinate patterns (see Materials and methods) and rephotographed. The correlative mitochondrial labeling in Fig. 3b demonstrates that (i) HSP60 labeling identifies mitochondria that correspond with mitochondria identified by dye labeling in Fig. 3a, (ii) the detailed structure of mitochondria is preserved along their entire length following manipulations involving detergent treatment and antibody labeling, and (iii) there is a better signal-to-noise ratio for detecting mitochondria by antibody labeling as compared with dye labeling. Correlative labeling thus allows us to unequivocally identify mitochondria, even when they are present as small vesicular, rather than tubular, structures. Dye staining can also be compared with irnrnunolabeling of the cytoskeleton. Figure 4 shows a low magnification example of the results obtained. Dye labeling alone for identifying organelles (Fig. 4a) is usually useful only in flat regions of the cell periphery where endomembrane mass is minimal. The distributions of IFs (Fin. 46) and MTs (Fig. 4c) can be identified throughout the celi. However, where these polymers overlap, the resolution limit of light microscopy (0.2 pm) may lead to the interpretation that the organelles associate with both cytoskeletal components. It is more useful to examine interrelationships in control cells in regions of the cell periphery at higher magnification. Quadruple labeling of organelles and the cytoskeleton ER, mitochondria, IFs, and MTs can all be visualized in the same cell by multiple labeling procedures, a useful approach if the objective is to directly evaluate the interrelationships of these structures. DiOC6(3) staining and photography in the fluorescein channel was done first (Fig. 5a), followed by detergent treatment and triple immunofluoresence. Because no reorganization of mitochondrial distribution was observed, it is likely that the cytoskeleton was similarly not affected by the procedure. Cascade blue mitochondrial HSP60 labeling is shown in the UV channel (Fig. 5b), fluorescein vimentin IF labeling is in the second channel (Fig. 5 4 , and Texas red MT labeling is in the rhodamine channel (Fig. 5d). The results obtained are as follows. First, the ER (Fig. 5a) for the most part is reticular, though more of a fibrous than of a tubular appearance, and there are numerous small ER aggregates. A large ER island or cisterna is present towards the upper right edge of the cell and there is a second cisterna in the

FIG. 4. Triple labeling of organelles and the cytoskeleton. (a) DiOC,(3) staining of glutaraldehyde-fixed human fibroblasts. (b) Immunofluorescent labeling of IFs in the same cell using vimentin antibody (fluorescein channel). (c) Immunofluorescent labeling of MTs using tubulin antibody (rhodamine channel). The bar in c represents 20 pm.

lower right quadrant. The upper ER cisterna is, interestingly, confined on all sides by tubular mitochondria (Figs. 5a and 5b). These bordering mitochondria appear to be fused one to another, forming a mitochondrial reticulum. This mitochondrial reticulum is best observed in Fig. 5b. A second example can be seen in Fig. 3. One striking observation that is apparent from Figs. 3 and 5 is the length of mitochondria, which often exceed 50 pm. For the most part,



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FIG. 5. Quadruple labeling of organelles and the cytoskeleton in human fibroblasts. (a) DiOC,(3) staining of ER and mitochondria. (b) HSP6O antibody labeling of mitochondria (UV channel-Cascade Blue fluorescence). (c) Labeling of IFs using vimentin antibody (fluorescein channel). (d) Labeling of MTs using tubulin antibody (rhodamine channel). An arrow identifies a mitochondrion in (a) and (b) that aligns with the I F bundle indicated in (c) and the M T bundle in (d). The bar in (c) represents 10 pm.

mitochondria are similarly oriented and associated with ER along their entire length (Fig. 5a). Vimentin labeling (Fig. 5c) indicates that IFs have a restricted distribution in this region, being present only in one bundle that does not extend to the cell margin. Alignment studies show that the IF bundle

runs along a MT bundle (Fig. 5d) and both of these are associated with one long mitochondrion. This mitochondrion continues further towards the cell margin in the absence of detectable IFs in its vicinity. The other mitochondria in the region do not have any detectable IFs in close

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F I G . 6. Effect of low concentrations of vinblastine on organelles and the cytoskeleton. Human fibroblasts were treated with 10 nM vinblastine for 6 h, glutaraldehyde-fixed and then quadruple labeled as in Fig. 5. (a) Dye staining of ER and mitochondria. (b)Mitochondria1 labeling with HSP60 antibody. (c) Labeling of IFs with vimentin antibody. (d) MT labeling using tubulin antibody. The bar in d represents 20 pm.



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FIG. 7. Effect of high concentrations of vinblastine on organelles and the cytoskeleton. Human fibroblasts were treated with 2 pM vinblastine for 6 h. (a) Dye staining of ER and mitochondria. (b) Mitochondria1 labeling with HSP60 antibody. (c) Labeling of IFs with vimentin antibody. ( d )MT labeling using tubulin antibody. Only tubulin paracrystals are present, two examples indicated by arrows. The bar in c represents 10 pm.

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proximity. These observations indicate that the mitochondrial distribution in the cell cannot be fully accounted for by IF distribution. MTs, on the other hand (Fig. 5d), are present throughout the cell region and thus are capable of interacting with all mitochondria as well as with IFs. Vinblastine effects on interrelationships To understand how changes in tubulin assembly affect organelle interrelationshipsand the status of IFs, the cellular effect of vinblastine was examined. Vinblastine is known to cause net depolymerization of preexisting MTs at low concentrations (nanomolar levels) and to induce the formation of tubulin paracrystals at much higher concentrations (micromolar levels) (see Jordan et al. 1991 and references therein). At vinblastine concentrations of < 5 nM, we were unable to observe changes in cells by our fluorescence approach (not presented). Figure 6 shows a cell treated with 10 nM vinblastine for 6 h. MTs have depolymerized (Fig. 6d; compare with Figs. 2c and 4 4 , although several single MTs still remain in the perinuclear region (these may not be visible in photographic reproductions). ER (Fig. 6a) and mitochondria (Figs. 6a and 6b) have aggregated together near the nucleus, leaving a lamellipodium nearly devoid of detectable subcellular membrane systems. The organelles are found along the anterior side of the nucleus (i.e., towards the lamellipodium) and surround the nucleus. IFs (Fig. 6c) are observed to have collapsed near the nucleus. In contrast to the location of the organelles, IFs are found primarily on the posterior side of the nucleus. The lack of identical distributions suggests that organelle aggregation and IF collapse are not directly related to each other. The result also indicates that the normal distributions of ER, mitochondria, and IFs are all dependent on the state of MTs. Figure 7 shows a cell treated with 2 yM vinblastine for 6 h to drive the formation of tubulin paracrystals. The cell shows significant rounding and retraction of lamellipodia. ER (Fig. 7a) is in the form of membrane aggregates, and both ER and mitochondria (Figs. 7a and 7b) are distributed together in several locations. In addition to their colocalization in the perinuclear region, they are found together in remnant cell processes, notably the two protruding from the upper part of the cell. IFs (Fig. 7c) have collapsed, as observed previously, and are located immediately on the left and top sides of the nucleus. The distributions of organelles and IFs are different. The distribution of tubulin paracrystals, visualized in Fig. 7d, does not suggest that these structures influence the distributions of organelles or IFs. Tubulin paracrystals are, however, localized in organellecontaining regions. Taxol effects Would an increase in MT stability affect interrelationships among organelles and the cytoskeleton? Taxol causes MT reorganization in cells and the formation in interphase cells of stable MT bundles. At an early time point ( c 4 h) taxoltreated cells are only distinguished from control cells by a gradual loss of an apparent centrosomal focus of MTs (results not shown; for a review, see DeBrabander et al. (1986)). Figure 8 shows a cell that has been treated with 15 p M taxol for 5 h. The observed MT reorganization into bundles (Fig. 8d) is typical of taxol-induced effects. Taxol does cause some cell rounding, making detailed investiga-

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tion of organelles difficult. Nevertheless, ER and mitochondria (Figs. 8a and 8b) are both organized along these MT arrays (Fig. 8d) and are distributed up to the cell periphery. IFs (Fig. 8c) appear to be following the MT distribution and forming a subset array within it. Thus, in taxol stabilized cells, the distributions of all four components appear interrelated.

Discussion We have developed a quadruple fluorescence labeling approach which enables visualization in the same cell of ER, mitochondria, IFs, and MTs. We sought, by this means, to gain insight into interrelationships between these organelles and cytoskeletal components, and how these are affected by MT drugs. The methodology Several aspects concerning our methodology are novel. Procedures were necessary to overcome problems associated with the use of glutaraldehyde as a fixative. Glutaraldehyde fixation is required for the preservation of ER for dye staining (Terasaki et al. 1984), optimally preserves MTs, and prevents the fragmentation and vesicularization of mitochondria caused by formaldehyde or cold methanol fixation (data not presented). One problem often faced is that when intact cells are fixed with glutaraldehyde, it is difficult to permeabilize them with detergents in order to label with antibodies. Prepermeabilizing cells before glutaraldehyde fixation, a standard procedure for visualizing cytoskeletal components (e.g., Soltys and Borisy 1985), is not an option for membranous organelles. Furthermore, it is well known that many antibodies cannot recognize antigens on glutaraldehyde- or formaldehyde-fixed proteins (e.g., Riederer 1989). This could be due to other cross-linked proteins blocking access of the antibody or it could result from fixing the protein in a conformation where the epitope is not accessible. We encountered this problem in the case of anti-HSP60 immunofluorescenceof mitochondria where, despite having several monoclonal and polyclonal antibodies, none reacted with glutaraldehyde-fixed cells permeabilized using several detergents (e.g., Triton X-100, deoxycholate, NP-40, etc.). We solved this problem by using the ionic detergent SDS for permeabilizing the cells. The detergent was effective at permeabilizing and rendering antigens irnrnunoreactive even when glutaraldehyde reactions were prolonged and at concentrations meant for electron microscopic preservation. The use of this detergent in immunocytochemistry is new. One concern in the use of SDS might be that protein denaturation or redistribution may occur. However, unfolding of proteins in the presence of SDS is temperature dependent and there is evidence that it is negligible at or below room temperature. Much less effect would be expected with glutaraldehyde cross-linked proteins. For example, in immunoprecipitation of native proteins, reactions with antibody are usually done in the presence of SDS (plus NP-40 and deoxycholate; so-called RIPA buffer), often overnight (e.g., Harlow and Lane 1988). This indicates no effect on native protein structure and antibody function. A second example is that immunofluorescence specimens, fixed with glutaraldehyde and labeled with antibodies, can be washed with SDS at room temperature without altering the distribution of bound antibody, indicating no effect on



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FIG. 8. Cellular distribution of organelles and cytoskeleton in tax01 treated cells. Human fibroblasts were treated with 15 pM tax01 for 5 h. (a) Dye staining of ER and mitochondria. (b) Mitochondria1 labeling with HSP60 antibody. (c)Labeling of IFs with vimentin antibody. (d) MT labeling using tubulin antibody. The bar in c represents 10 pm.


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the bound native antibodies (data not presented). We have similarly concluded that the distribution of properly fixed (cross-linked) antigens in our preparations are not affected by SDS treatments. The use of SDS is further advocated by the unprecedented preservation of mitochondria we observed by antibody labeling, corresponding exactly with mitochondrial structure seen by dye staining of intact cells, and by the ability to label IFs and MTs. We have additionally observed that in living cells labeled with fluorescent transferrin, the endocytic marker is retained following fixation and permeabilization, indicating preservation of ligand receptor complexes contained within endosomes (results not shown). The quadruple-labeling approach that we have used has general applicability. In practice, it consists of two steps. The first involves dye labeling of a specific organelle and this label must be extractable during detergent permeabilization of cells. The second step is triple immunofluorescence for other cellular components. Triple immunofluorescence with conventional optics has been reported (e.g., Ball and Singer 1982). We used the cyanine dye DiOC6(3)as a stain for ER (Terasaki et al. 1984). Although the dye reacts additionally with mitochondria, the latter could only be observed well at flat cell peripheries where ER density was minimal. This required us to incorporate immunofluorescent visualization of mitochondria into our specific application. DiOC6(3) stains ER and mitochondria in both living and glutaraldehyde-fixed cells (Terasaki et al. 1984), and this allows flexibility in experimental design. Comparable labels are available for other organelles. A fluorescent ceramide analogue stains the Golgi apparatus (Lipsky and Pagano 1985) and can be used in both living and glutaraldehydefixed cells (Pagano et al. 1989). Being a fluorescent lipid, it is extractable with detergent treatments. Finally, endosomes and lysosomes can be selectively labeled by uptake in living cells of fluorescent ligands or fluid phase markers (see Willingham and Pastan 1985).

Interrelationships among organelles and the cytoskeleton What insight has our approach provided concerning the interrelationships of ER, mitochondria, IFs, and MTs? We have observed that mitochondria are always associated with ER, although ER can also exist independently. That the codistribution is maintained following aggregation owing to MT disassembly indicates this association is stable. This suggests that similar cytoskeletal requirements are operative in both cases for the maintenance of an extended state. Following MT depolymerization, we observed that the aggregation of mitochondria and the collapse of IFs are spatially separate. This indicates that these are independent phenomena, yet both owing to lack of MTs. Previous studies have observed concurrent mitochondrial aggregation and IF collapse in response to various treatments, including MT depolymerizing agents (see Chen et al. 1984; Chen 1988), acrylamide (Eckert 1986), and heat shock treatment (Shyy et al. 1989). These phenomena, however, were interpreted to be causally related and resulting from mitochondria being bound to IFs. Mitochondria-ER associations were not considered. Our observations on mitochondria-ER associations instead indicate that mitochondrial aggregation is due to retained association with ER that can no longer maintain an extended state. Thus, our result may serve to correct these previous interpretations, although differences in cell treatments or cell types may need

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to be reevaluated. Unfortunately, there are no drugs available which specifically alter IF assembly. Our prediction is that mitochondrial aggregation (disorganization)is a general indicator of an aggregated state of ER membranes. Other studies have implicated MTs as being solely responsible for mitochondrial distributions. Notably, Ball and Singer (1982) used cycloheximide treatment of fibroblasts to cause IF collapse and observed that mitochondria remained associated with MTs, which were unaffected. Further, microinjection of monoclonal antibody to vimentin causes IF collapse without affecting motility and mitochondrial distributions (Lin and Feramisco 1981; Summerhayes et al. 1983). Thus, IFs may at best have only an auxiliary role to that of MTs in the distribution of mitochondria (Klymkowsky et al. 1989). Studies using microinjection of fluorescently labeIed tubulin (e.g., Soltys and Borisy 1985) have shown that MTs in living cells are highly dynamic in terms of subunit exchange. In the case of vinblastine-treated cells, we find that tubulin polymerized into stable paracrystals does not influence organelle distributions. This could result if MTassociated proteins or MT motors that mediate interactions with organelles are not able to bind or exert effect on an altered polymer lattice. In taxol-treated cells both organelles and IFs appeared distributed along taxol-stabilized MTs, indicating polymer stability per se does not interfere with the operating mechanism(s). We observe that mitochondria can form networks that have some morphologic similarities with ER. Both can be present as elongated tubules forming an interconnected reticulum. This would be consistent with mitochondrial form and distribution resulting from mechanisms analogous with those postulated for ER formation along MTs (Terasaki et al. 1986; Dabora and Sheetz 1988; Lee et al. 1989). Mitochondria are additionally present in vesicular form. The cellular consequences of tubular versus vesicular mitochondria and their possible interconversion are areas that remain to be explored. Acknowledgements This work was supported by a research grant from the Medical Research Council of Canada to R.S.G. We thank Barbara Sweet for her secretarial assistance. Ball, E.H., and Singer, S.J. 1982. Mitochondria are associated with microtubules and not with intermediate filaments in cultured fibroblasts. Proc. Natl. Acad. Sci. U.S.A. 79: 123-126. Bereiter-Hahn, J. 1990. Behavior of mitochondria in the living cell. Int. Rev. Cytol. 122: 1-63. Buchwald, M., and Ingles, C.J. 1976. Human diploid fibroblast mutants with altered RNA polymerase 11. Somatic Cell Genet. 2: 225-233. Cambray-Deakin, M.A., Robson, S.J., and Burgoyne, R.D. 1988. Colocalization of acetylated microtubules, glial filaments, and mitochondria in astrocytes in vitro. Cell Motil. Cytoskeleton, 10: 438-449. Chen, L.B. 1988. Mitochondrial membrane potential in living cells. Annu. Rev. Cell Biol. 4: 155-181. Chen, L.B., Summerhayes. I.C., Nadakavukaren, K.K., Lampidis, T. J., Bernal, S.D., and Shepherd, E.L. 1984. Mitochondria in tumor cells: effects of cytoskeleton on distribution and as targets for selective killing. Cancer Cells, 1: 75-86. Dabora, S.L., and Sheetz, M.P. 1988. The microtubule-dependent


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