Whole-Mount Electron Microscopy of the Cytoskeleton: Negative Staining Methods J. Victor Small and Antonio Sechi I. INTRODUCTION Various techniques may be used for observing the cytoskeleton of whole cultured cells in the electron microscope (EM). These include freeze drying followed by metal coating (Heuser and Kirschner, 1980) and chemical fixation followed by either critical point drying (Wolosewick and Porter, 1979; Schliwa et al., 1982; Svitkina et al., 1995) or negative staining (Small and Celis, 1978; Small, 1988). Each of these techniques has its advantages and drawbacks (see, e.g., Small, 1988), and a choice between one or the other will depend on the problem at hand. The negative staining method described here is the simplest and is well suited to studies of the thin, peripheral areas of cultured cells. II. MATERIALS AND INSTRUMENTATION Sodium chloride (Cat. No. 1.06404), magnesium chloride hexahydrated (Cat. No. 5833), and glucose (8337) are from Merck. Triton X-100 (Cat. No. T9284), MES (Cat. No. M8250), bacitracin (Cat. No. B-0125), and streptomycin sulfate (Cat. No. S6501) are from Sigma. Phalloidin was a generous gift from Professor H. Faulstich (Heidelberg), but can be obtained from Sigma (Cat. No. P2141). EGTA (Cat. No. 03780) is from Fluka. Falcon petri dishes are from Becton-Dickinson. Electron microscopy grids EMBRA 135 or 150 mesh hexagonal [code numbers: 8GG135/150 (gold), 8GZ135/150 (silver), and 8GN135/150 (nickel)] are from Graticules Ltd. Glutaraldehyde (EM grade Cat. No.R1020) is from Agar Scientific Ltd. Sodium silico tungstate (Cat. No. S019) and EM forceps (Nos. 4 and 5) are from TAAB. Filter paper (Whatman No. 1) and disposable syringe filters (0.20/0.45 mm cutoff) are from Sartorious. Glass coverslips (24 1 24 mm or 8 1 24 mm), glass slides, coplin jars, a glass trough (7 cm or more deep), tissue paper, lint-free paper, razor blades, Agepon, Pasteur pipettes, petri dishes, stoppered volumetric flasks, and multiwell dishes are from local suppliers. III. PROCEDURES A. Preparation of Collodion Film For preparation of formvar films, see Small and Herzog (1994). Solutions 1. Collodion stock solution: 2 g collodion dissolved in 100 ml amyl acetate. Collodion needs about 3 4 days to dissolve at room tempereture with constant stirring. We usually CELL BIOLOGY: A LABORATORY HANDBOOK, Second Edition. Vol. 3. Copyright 1998 by Academic Press. All rights of reproduction in any form reserved. 285
filter the collodion solution through glass wool to remove unwanted particles and store it in a coplin jar at room temperature. 2. Collodion working solution: 1.3% collodion in amyl acetate. To make 100 ml, mix 65 ml 2% collodion with 35 ml amyl acetate. Store in a coplin jar at room temperature. 3. Agepon solution: 5% in water (v/v), store in a coplin jar at room temperature. 4. Polybutene solution: 1% in toluene. Store in a stoppered flask at room temperature. (Polybutene can normally be obtained as a free sample from an oil company.) See Fig. 1. 1. Dip a glass slide in chromic acid in a coplin jar for at least 30 min (Fig. 1a, 1). 2. Wash the slide extensively with tap water and then rinse it in distilled water (Fig. 1a, 2). 3. Place the coplin jar in a oven at 60 C in order to dry the top part of the slide (Fig. 1a, 3). This allows the slide to be readily handled and prevents the Agepon (next step) from forming a drainage line down the center of the slide. 4. Dip the slide in a 5% Agepon solution (or equivalent photodetergent) for 5 min and then transfer it to a dust-free place for 12 24 hr (we normally use an exicator, Fig. 1a, 4 and 5). Agepon facilitates separation of the film from the slide in Procedure B, step 2. 5. Dip the dried, Agepon-coated slide in 1.3% collodion solution for 10 30 sec and then transfer it to another exicator to allow the collodion to drain and dry (the drying time depends on the collodion concentration; in our hands, a 1.3% collodion solution needs a few hours to dry completely) (Fig. 1a, 6 and 7). B. Preparation of Filmed Grids 1. Remove the slide from the exicator and score the film on the glass with a razor blade into pieces of about 8 1 24 or 24 1 24 mm in size. To improve the detachment of the film, it can also be scored around the edges of the slide (Fig. 1b, 1). 2. Fill the glass trough with distilled water and sweep surface clean with one piece of lint-free paper. Holding the frosted strip of the slide, dip it slowly into the water at an angle of around 45 to release film from the surface. If film separates poorly, breath gently on slide surface. Film should be silver-gold in color in reflected light. (Fig. 1b, 2 and 3). 3. Place grids gently onto film, taking care not to touch the film with forceps. To pick up the grid film combination, hold one end of a glass coverslip with forceps and position the other end at one end of the film with the coverslip tilted upward. Then push the coverslip downward into the water to force the film against the coverslip surface and rotate the combination under water before removing so that the film finally faces upwards, as shown in Fig. 1b, 4. Place the grid sets on filter paper and allow them to dry (Fig. 1b, 5). 4. Sterilize grid set under UV light in an open petri dish before plating the cells. C. Cells Cells are plated onto the grids the same as for coverslips, and the density is chosen to give one to two cells per grid square after attachment and spreading. In general, cells spread more slowly on plastic films than on glass; to encourage spreading, the filmed grids may be incubated with a drop of serum (or extracellular matrix molecules) overnight, prior to plating. Check for adequate cell spreading in an inverted microscope prior to fixation. 286 Electron Microscopy
FIGURE 1 Schematic diagram indicating steps for preparing collodion-filmed slides (a) and the filmed grid sets on coverslips (b) used for cell culture. For a further explanation, see text. Whole-Mount Electron Microscopy of the Cytoskeleton 287
D. Fixation Solutions 1. Cytoskeleton buffer (CB): 10 mm MES, 150 mm NaCl, 5 mm EGTA, 5 mm MgCl 2 r6h 2 O, and 5 mm glucose, ph 6.1, at room temperature. For 1 liter, weigh out 1.95 g MES, 8.76 g NaCl, 1.90 g EGTA, 1.02 g MgClr6H 2 O, and 0.90 g glucose. Dissolve in 900 ml H 2 O, adjust ph to 6.1 with 1 N NaOH, and fill up to 1 liter. Store at 4 C. For extended storage, add 100 mg/liter streptomycin sulfate. 2. Glutaraldehyde stock: 2.5% solution of glutaraldehyde made up in CB and stored at 4 C. Dilute 25% EM grade glutaraldehyde 1:9 with cytoskeleton buffer. Recheck that ph is 6.1. Aliquot and store at 020 C. 3. Glutaraldehyde Triton mixture: This fixative consists of a mixture of glutaraldehyde and Triton X-100 made up by diluting appropriate volumes of the glutaraldehyde stock and Triton X-100 (stored as a 10% aqueous stock) into CB. We commonly use a mixture of 0.25% glutaraldehyde and 0.5% Triton in CB. 1. The washing and fixation solutions are all used at room temperature and the 24- multiwell dish serves as a useful reservoir for holding the different solutions. Fill the wells of the dish as follows: wells 1 and 2, CB; 3, the glutaraldehyde Triton mixture ; 4, CB; 5, 2.5% glutaraldehyde; 6 and 7, CB. A small petri dish (3.5 or 5 cm diameter) containing CB should also be prepared. 2. After removing the cells from the incubator, transfer the coverslips carrying the grids through the wells of the dish as follows: 1 and 2, 2 sec each; 3, 60 sec.; 4, 2 sec.; 5, 10 min.; 6 and 7, 2 sec. each. Store in petri dish in CB. E. Actin Stabilization Solution 1. Phalloidin: 10 mg/ml phalloidin in CB. Store phalloidin as a 1-mg/ml stock in methanol at 020 C. To stabilize actin so that filaments are better visualized after negative staining, treatment with phalloidin is advisable. At this stage the back side of the grids may also be dried in preparation for negative staining. 1. Remove a grid from coverslip with forceps, after scoring around the edge to break the surrounding film, and rinse twice briefly in water (conveniently placed in two 250-ml beakers). Then blot the back side of the grid on a piece of filter paper. At this stage it is best to briefly release the grid onto the filter paper, drain excess liquid from the forceps, retrieve the grid, and then invert it on a drop of phalloidin on a sheet of Parafilm. It is important that the cell side of the grid does not dry or come into contact with filter paper during these steps. If liquid drains rapidly through the grid onto the filter paper during blotting, the film is broken and the grid may quickly dry if appropriate care is not taken. 2. Leave the grid on phalloidin for 30 60 min at room temperature. Alternatively, the grids can be incubated in phalloidin at 4 C overnight or longer at the end of fixation, step 2 (above). For extra stabilization of microtubules, taxol could be used, if desired. 288 Electron Microscopy
F. Negative Staining Solutions 1. Negative staining solutions: 1% aqueous uranyl acetate and 3% aqueous sodium silicotungstate, both filtered through a 0.20-mm filter and stored at 4 C in stoppered flasks. 2. Spreading solution: 100 mg/ml bacitracin (Sigma) in H 2 O, made fresh from a 2001 concentrated stock solution stored at 4 C and supplemented with 0.1% amyl alcohol. FIGURE 2 Examples of 3T3 fibroblast cytoskeletons prepared as described and stained with uranyl acetate. The figure shows the lamellipodium at the cell edge that is rich in actin filaments (a) and a medial region (b) in the cell in which actin filaments (a), intermediate filaments (if), and microtubules (mt) are visible. Bars: 0.2 mm. Whole-Mount Electron Microscopy of the Cytoskeleton 289
1. Clamp the grid in a pair of forceps using a large paper clip over the shaft to hold the tips together. 2. Keeping liquid away from the back side, rinse the cell side of the grid with several drops of bacitracin (for this step the grid can be tilted vertically and drops added from the side with a Pasteur pipette). 3. Place forceps on the lid of a petri dish so that the grid is held cell side up and drain away excess liquid with the torn edge of a filter paper held where the forceps grip the grid. FIGURE 3 As in Fig. 2 but stained with sodium silicotungstate. Bars: 0.2 mm. 290 Electron Microscopy
4. Immediately add one drop of the negative stain solution, leave a few seconds, and drain again with filter paper to leave a thin film. 5. Allow to air dry and then observe in the microscope. IV. COMMENTS AND PITFALLS We routinely use EMBRA grids from Graticules. The advantage of these grids is that they are thick enough to prevent the support film from damage caused from sagging onto the glass surface of the carrier coverslip used for cell culture. Moreover, EMBRA grids are slightly convex (toward their shiny face); this also helps keep the film away from the glass surface of the coverslip. In our hands collodion films give a slightly clearer background after negative staining than formvar, but with whole cells they can break more readily under the electron beam. For this reason we use quite thick support films. Some thinning of the film occurs during irradiation, leading to improved contrast. Uranyl acetate produces higher contrast than sodium silicotungstate and better visualization of microtubules and intermediate filaments (Fig. 2). With sodium silicotungstate, the actin filament order, especially within the lamellipodia, is better preserved (Fig. 3). As is normal with negative staining, variability in stain intensity across the grid is not uncommon, so cells must be sought that exhibit the appropriate contrast. Experiment with different accelerating voltages if problems with stain contrast consistently arise. References Heuser, J. E., and Kirschner, M. W. (1980). Filament organization revealed in platinum replices of freeze-dried cytoskeletons. J. Cell Biol. 86, 212 234. Schliwa, M., van Blerkom, J., and Pryzwansky, K. B. (1982). Structural organization of cytoplasm. Cold Spring Harb. Symp. Quant. Biol. 46, 51 67. Small, J. V. (1988). The actin cytoskeleton. Electr. Microsc. Rev. 1, 155 174. Small, J. V., and Celis, J. E. (1978). Filament arrangements in negatively stained cultured cells: The organization of actin. Cytobiologie 16, 308 325. Small, J. V., and Herzog, M. (1994). Whole-mount electron microscopy of the cytoskeleton: Negative staining methods. In Cell Biology: A Laboratory Handbook (J. E. Celis, ed.), Vol. 2, pp. 135 139. Academic Press, San Diego. Svitkina, T. M., Verkhovsky, A. B., and Borisy, G. G. (1995). Improved procedures for electron microscopic visualization of the cytoskeleton of cultured cells. J. Struct. Biol. 115, 290 303. Wolosewick, J. J., and Porter, K. R. (1979). Microtrabecular lattice of the cytoplasmic ground substance. Artifact or reality. J. Cell Biol. 82, 114 139. Whole-Mount Electron Microscopy of the Cytoskeleton 291