Negative Staining. Negative stain solutions. Staining samples on formvar-carbon coated grids

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1 Negative Staining Negative stain solutions Aqueous Uranyl Acetate A 1% to 3% solution of uranyl acetate dissolved in water can be used to negatively stain many samples. The stain has a low ph so this solution is not recommended for particles that are unstable in acid conditions. Neutral Phosphotungstic Acid A 1% to 3% solution of phosphotungstic acid is made up in water and the ph is adusted to 7 using sodium hydroxide. This is a useful stain for many samples but is especially good for viruses that dissociate at low ph. The stain produces less contrast than the uranyl acetate. Ammonium Molybdate Make up a 1% solution of ammonium molybdate in water. This solution has also been used to negatively stain thawed, thin cryosections of fixed cells. Methylamine Tungstate Make up a 4 % solution in water. It s difficult to dissolve so you need to vortex. Centrifuge before use to pellet insoluble particulates. Introduction Specimens that are to be examined in the transmission electron microscope (TEM) usually have to be thin, dry and contain contrast (usually from a heavy metal stain. One of the easiest ways of preparing biological samples, of small size, for the TEM is by negative staining. This preparation method is useful for visualizing suspensions of small particles, which includes viruses, purified proteins, liposomes and small vesicle fractions. Negative staining is a simple technique for routine examination of structure. It does not allow for high resolution examination of samples - more technically demanding methods, using sample vitrification (or rapid freezing) are used for this. As with most techniques, there are levels of complexity which yeild increasingly better results. The major differences between the techniques is in the choice of sample support. Staining samples on formvar-carbon coated grids The sample is suspended (or diluted) into water (if possible) and adsorbed onto a carbon-coated formvar film which is attached to a metal specimen grid. The carbon surface of the grid becomes contaminated when stored, and thus hydrophobic, so it is best to glow discharge the grid surface (making it hydrophilic) prior to use. This is usually done in a vacuum chamber of a vacuum evaporator.

2 Once the specimen has been adsorbed onto the film surface, the excess sample is blotted off and the grid is covered with a small drop (5 µl) of stain solution (see below). This is left on the grid for a few minutes and then blotted off. The sample is dried and examined in the TEM. If carbon-formvar films are prepared on specimen grids with small mesh size, the films, with adsorbed sample, can usually be immunolabeled without them breaking. This protocol is described in more detail in the section on immunolabeling. Briefly, the grids are floated onto drops of diluted antibody. washed by floating on drops of buffer, and then floated on drops of diluted visualization probe. We normally use colloidal gold coupled to staphylococcal protein A (protein A-gold) or other affinity markers such as antibodies. Methods using carbon films alone I. Carbon-coated grids The grids are prepared by first preparing formvar-carbon coated grids and then removing the formvar support. This is done by placing the grids in an atmosphere of solvent vapour, which dissolves the formvar (which is a plastic). The grids are placed on a wire mesh in a glass perti dish, the solvent (chloroform or carbon tetrachloride) is placed in the dish below the wire mesh and the dish is closed by replacing the lid. If the vapour alone does not remove the film (which should take a few hours), the process can be accelerated by dipping the coated grids into the solvent prior to closing the dish and placing them on the mesh to soak. Allow the solvent to completely evaporate before removing your grids, which will now have only carbon on them. Remember that the carbon must be thick enough to be self supporting. Advantages include high stability in the electron beam and high resolution examination of the adsorbed specimen. Disadvantages include a hydrophobic surface which is not of uniform thickness. II. Carbon films For this technique, carbon film is evaporated onto a freshly cleaved mica surface and the sample is applied to the surface of the carbon film attached to the mica. The carbon film and the sample is then picked up onto a specimen grid and examined in the TEM. The carbon films can be stored on the mica for long periods without them becoming contaminated, the films are thin but tough so can be easily examined in the TEM without any noticable specimen movement. However, they are fragile and must be manipulated with care. They are more stable if supported on specimen grids with small mesh size (e.g. 400 mesh). Hexagonal grids seem to offer a more stable support.

3 Making the carbon film Take a piece of mica and cut off a square or rectangular piece that is approximately 2 x 3 cm. Cleave the mica with a razor blade or scalpel that has been cleaned with acetone. Once the mica has begun to separate along a cleavage plane, forceps may be used to pull the mica completely apart. It is better to do this than to scratch the surface of the mica with the razor blade. Mica, as purchased from most EM supply companies, is rather thick and a new piece may be split around 4 times. Attach the mica to a filter paper, with sticky tape, with the freshly cleaved plane facing upward. Place the filter paper in a vacuum evaporator for carbon coating and deposit a film of carbon onto the mica surface. This mica should be cleaved immediately before coating as the freshly cleaved surface is clean and hydrophilic, but it becomes contaminated (and thus hydrophobic) over time time. After coating, the filter paper should be light gray in color (compare the filter paper behind the mica with that which was exposed, and therefore carbon coated). This is a good indicator of the film thickness. Negative staining Prepare the sample (the staining works best if the sample is in water) by making several serial dilutions of the preparation. The best negative stain preparations are those that have a single layer of individual, separated particles adsorbed onto the film. This is achieved by dilution of the sample. With a pair of scissors, cut off a small square of mica, about 4 x 5 mm, about twice the size of an EM grid. Put the square on a piece of parafilm, carbon side up. With a pipetman, gently squirt 5-10 ml of the sample under the carbon by placing the pipette tip to the side of the mica square. The sample will do one of two things: 1) It will flow between the carbon and mica, in which case you will need only a small amount of sample. 2) It will flow between the mica and the parafilm, in which case just keep applying sample to the side of the square until it goes under the carbon. Place a 400 mesh specimen grid onto the carbon. (Wash the grid in 0.5% acetic acid and then acetone prior to use) Break the carbon film to free the specimen grid, lift the grid and place it on a drop of stain solution for about 30 seconds (sample side down). Blot dry, and examine in the TEM when completely dry. Staining and washing times can be varied. A better support for the carbon film can be obtained by using specimen grids coated with a holey film of formvar. This is a film containing lots of small holes which are covered by the carbon film. There are many ways to produce these films but one quick, simple way is to breath onto a film of formvar before it dries onto the glass slide. The moisture droplets will displace the film, leaving small holes. Althernatively, the wet film on the glass slide can be placed in a cloud of steam.

4 Adenovirus particles An example of negative staining. These adenovirus particles have been adsorbed onto a carbon film that was deposited onto a freshly cleaved mica surface. The film was picked up onto a clean, 200 mesh specimen grid coated with a holey formvar film. The preparation was stained for 1 minute with neutral 1% aqueous phosphotungstic acid and photographed in a transmission electron micrograph. Other Negative Staining procedures Techniques for the preparation of negative stain specimens are simple and direct. The essential aim of the procedure is to embed the specimen in a uniformly thin deposit of stain. Resolution of molecular features is only accomplished at the stain-specimen boundary where there is maximum contrast. This result is only achieved if the deposition of buffer salts or other materials with densities less than the stain at that boundary are severely limited; otherwise the specimen molecules will be imaged at low resolution and will appear as nondescript blobs. The specimen sample is usually applied directly to the surface of the support film where a population of specimen particles becomes adsorbed. Attachment of the molecules is usually secure enough that they are not removed by subsequent rinsing and staining operations which do remove most of the buffer salts. Since different specimens often have different affinities for the particular support film being used, some measure of control over how much specimen attaches to the film may be effected by adjustment of the specimen and buffer concentrations, adsorption time, etc. Appropriate conditions must be established by experiment for each new specimen. For protein solutions, typical concentrations range between µg/ml and adsorption times from as little as 1-5

5 seconds to several minutes. Stains are usually applied in a range of concentration from %. Adjustment of stain concentration provides some control over the thickness of the deposit. It does NOT follow that a procedure that is successful with one type of specimen is also suitable for another, so various modifications should always be tried until good contrast and spreading conditions are achieved. There are two common procedures for preparing negatively-stained specimens on EM grids: Adhesion (drop) method (Figs. II.42 and II.43) A droplet of specimen is placed on the surface of the grid support film, making sure it sufficiently wets the surface. After an appropriate time interval, excess specimen is wicked away by touching a piece of filter paper to the edge of the grid surface. Without letting the grid dry, a droplet of rinse or stain solution is applied to the grid. Rinsing is necessary if the specimen preparation contains high concentrations of buffer salts or other solutes which may interfere with deposition of stain. The nature of an appropriate rinse depends on the conditions that the specimen can tolerate. Many viruses, for example, can withstand rinsing with distilled water. In some instances the stain solution can itself act as a suitable rinse. After rinsing and staining, excess fluid is wicked from the grid, leaving a thin aqueous film on the surface which is left to dry, usually in air. Fig. II.42. Preparation of a specimen from particles in aqueous suspension. (From Hall, p.290) Fig. II.43. Washing a specimen. (From Hall, p.290) The specimen can also be applied to the support film by floating the grid on top of a droplet of the specimen solution. The grid is then transferred to droplets of rinse and stain solutions and then dried as before. An additional variation of the usual adhesion method is to apply the sample to a holey support film in the same way as is done on regular films. The sample dries in a thin layer of stain stretched out over the holes, thereby giving maximum contrast since there is no

6 plastic and/or carbon support. Also, the stain tends to be more evenly distributed around the particle although the particle often undergoes distortions (shrinkage and flattening) due to the surface tension forces created as the layer of stain dries. The stain layer also has a tendency to break either before or after it is exposed to the electron beam. The layer of stain can be stabilized with a thin layer of evaporated carbon. Another advantage of this technique over the usual method is that, if small enough, the specimen particles will be randomly oriented in the stain layer. On regular support films, particles often settle on the surface of the film in one or a few preferred orientations, thus limiting the possible views of the specimen. Spray droplet technique(fig. II.44) The normal adhesion method of preparing a particulate suspension may lead to erroneous conclusions about the relative proportion of particles since different particles are likely to have different affinities for the substrate. Also the microscopist may select fields attractive to the eye but which are not representative. The only reliable way of preparing specimens without introducing a bias is to dry a drop of the original sample in its entirety. Non-volatile salts and buffers must be removed by centrifugation and washing or by dialysis so they don't obscure the particles under study or alter the structure when the salt concentrates in the last stages of drying. The entire residue from the drop must be examined, thus it is necessary to obtain very small drops. The suspension is atomized as a fine mist and the droplets are allowed to impinge on the substrate. Fig. II.44 A simple hand-held nebulizer from which the sample (s) is sprayed onto mica (m). (From Willison and Rowe, p.69) The spray droplet technique is particularly useful for examining specimens that adsorb so poorly to the support film that application and removal of rinsing and stain solutions also removes the specimen. Appropriate volumes of the specimen and stain solutions are mixed and sprayed in small droplets onto a wetable support surface. If the solution itself has the propensity to wet and spread over the surface, uniformly thin deposits of negatively-stained specimen result. The resultant aqueous film will be of uniform depth, and the mass of stain deposited per unit area of support film tends to be constant. When the specimen appears to have dried, some water may still be present in the stain bed, and a rearrangement of the stain deposit could result from its rapid vaporization if the

7 specimen is suddenly placed in the vacuum of the microscope. Thus, the specimen is usually allowed to dry (sometimes over a desiccant) for at least 10 minutes. Drying of the aqueous film proceeds from the edges, the central area covered by the droplet being last to dry. Minor solutes tend to be held in solution until the last stages of drying and are deposited in highest concentrations in the central area. As a result, the specimen in this area ordinarily is of inferior quality. Fig. II.45. Drop pattern. The small particles are tomato bushy stunt virus, and the large spheres are polystyrene latex particles 2,600 Å in diameter. The wedge-shaped sector has been magnified and superimposed. (From Hall, p.359) Note that, using the adhesion drop method, there may be preferential adherence of particles so relative particle distribution counts cannot be made. A major advantage of the spray technique over the adhesion method is that preferential adherence to the grid of one type of particle over another type of particle in a mixture cannot occur. Thus, this is the method of choice in quantitative studies where relative concentrations of particles in the sample need to be determined. By knowing the volume of the original drop (from adding known concentrations of polystyrene spheres) a count of the number of particles in a drop pattern provides immediately the number of particles per unit volume (Fig. II.45: Note that this figure shows a metal-shadowed specimen). This, together with the mass per unit volume obtained by weighing the dried residue from a measured volume can be used to calculate a value for the mean molecular weight of the particles.

8 High resolution negative staining (From Valentine et al, Biochemistry 7: ) Rationale: For the highest resolution with negative staining, there should be little or no support film, but some support is necessary to hold the protein. In this method the proteins are supported by a carbon film and "embedded" in a film of uranyl acetate. The film is cast on mica, which provides the cleanest possible surface for the carbon. 1. Freshly cleave a piece of mica, coat with a carbon film using the vacuum evaporator. 2. Put ~30 mg/ml protein solution in a small vessel. 3. Cut the mica to 3-4 mm 2 pieces. Hold a piece with forceps and push into the solution of protein at a degree angle. Do not let the film detach completely from the mica. 4. Let the film sit on the protein solution for seconds. 5. Pull the mica back and allow the film to sit on the mica.

9 6. Then slide the mica and film onto a solution of 1-2% uranyl acetate or uranyl sulfate. Pick up the film with a copper grid, dry, and examine in the electron microscope. Variations: A more stable film can be obtained if you pick up the film on a grid coated with a holey formvar film. A wrinkled carbon film may be better than a smooth one, as it seems to keep pools of uranyl acetate around the protein better than smooth films. Try varying the protein concentration for optimal staining. The proteins should be closely spaced to collect the stain but if they are too crowded proteins will be overlapped and will not be easily resolved. The carbon support film should be as thin as possible (3-10 nm) and should be prepared on freshly cleaved mica. Once the carbon coated mica has been prepared it can be stored in a dessicator almost indefinitely. Thicker support films will lower image contrast. Stain solution should be removed from below the surface of a stock stain solution to avoid contaminants from precipitates that may exist at the surface or the bottom of the stock solution. The precipitates do not affect the overall quality of the staining procedure but will result in local heating of the support film when exposed to the electron beam, which in its most benign form will result in drift, and at its worst will result in a ripping and curling of the support film. The carbon support film should only be exposed to proteins that exist in the bulk solution and must not be exposed to denatured material that is usually present in the meniscus. The greatest potential source of contamination and the one most often ignored is the contamination that arises from the tweezers and the support grid. At no time should the tweezers or the support grid be dipped into either the protein solution or the negative staining solution.

10 Procedures for the Preparation of Carbon Support Films Norm Olson Last Updated March 2000 A. Preparation of normal carbon support films 1. Copper grids should be pre-cleaned by sonicating for 10 sec. in acetone followed by 10 sec. of sonication in ethyl alcohol. Allow grids to dry on filter paper in a dust-free environment before use. 2. Add 0.12g of Formvar powder to 50 ml of ethylene dichloride and mix well on a magnetic stirrer until dissolved. Pour the solution into a clean coplin jar. 3. Clean a glass slide with water and detergent. Rinse well to make sure that all of the detergent is removed and finally rinse in de-ionized water before drying with a paper towel. Blow off any lint on the slide with compressed air. Place the slide in a dry, dustfree environment such as on filter paper under an upturned beaker. If there are problems in getting the plastic film to be released from the slide (Step 5), using a slide that has not been as thoroughly cleaned might help. 4. Dip the cleaned slide into the Formvar solution (Fig. 1-1) and touch edge to filter paper to drain off the excess fluid (Fig 1-2). Dry upright in a dust-free environment (this requires 5 to 10 min.). 5. Score the edges of the Formvar film with an acetone-cleaned razor blade (Fig. 1-3). Breathe on the slide to loosen the film, and slowly slide off onto a clean water surface by immersing the slide into the water at a ~15 angle (Fig. 1-4). Place grids, dull/rough surface down, onto good (uniform, gray color, un-wrinkled, etc.) areas of the film. Place a small piece of clean, white office paper onto the surface of the grids and film and allow the paper to soak up water. Pick up the paper, grids and film and place in a covered petri dish to dry. 6. Carbon coat film according to directions (see Sec.C) to desired thickness (A lightbrown color indicates a thickness of <100Å.).

11 7. Place the paper and coated grids onto a piece of filter paper that is soaked with ethylene dichloride in a covered petri dish. One half hour should be sufficient time to dissolve the Formvar film and not damage the carbon support. Remove the grids and paper and allow them to dry in a dust-free area. B. Preparation of perforated carbon support films 1. Copper grids should be pre-cleaned by sonicating for 10 sec. in acetone followed by 10 sec. of sonication in ethyl alcohol. Allow grids to dry on filter paper in a dust-free environment before use. 2. Add 0.17 g of Formvar powder to 50 ml of chloroform and mix well on a magnetic stirrer until dissolved. Pour the solution into a clean coplin jar. 3. Clean a glass slide with water and detergent. Rinse well to make sure that all of the detergent is removed and finally rinse in de-ionized water before drying with a paper towel. Blow off any lint on the slide with compressed air. Place the slide in a dry, dustfree environment such as on filter paper under an upturned beaker. If there are problems in getting the plastic film to be released from the slide (Step 6), using a slide that has not been as thoroughly cleaned might help. 4. Add about 50 drops of a 50% glycerol/water solution to the surface of the Formvar solution. Place the tip of a probe sonicator onto the surface of the solution and sonicate until mixed. Sonication intensity should be great enough to "violently" cause the solution to bubble. This often requires not much more than about 5 seconds. This should produce numerous holes that are 1-2 µm in diameter and suitable for use with frozen-hydrated samples. Sonicating for longer periods of time produces smaller holes in the film. 5. Immediately after sonicating, dip the cleaned slide into the Formvar solution (Fig. 1-1) and touch edge to filter paper to drain off the excess fluid (Fig 1-2.). Dry upright in a dust-free environment for about 5 to 10 min. 6. Score the edges of the Formvar film with an acetone-cleaned razor blade (Fig. 1-3). Breathe on the slide to loosen the film, and slowly slide off onto a clean water surface by immersing the slide into the water at a ~15 angle (Fig. 1-4). Place grids, dull/rough surface down, onto good (uniform, gray color, unwrinkled, etc.) areas of the film. Place a small piece of clean, white office paper onto the surface of the grids and film and allow the paper to soak up water. Pick up the paper, grids and film and place in a covered petri dish to dry. 7. Place the paper with the film and grids onto a methanol-soaked piece of filter paper in a covered petri dish for about 30 minutes. This should perforate any pseudo-holes (these occur when a small drop of glycerol was present but it was not enough to perforate the

12 film) that may be in the films. After allowing the paper and film to dry, the grids may be examined in a light microscope under phase contrast to determine the quality of the films. 8. Carbon coat film according to directions (see Sec.C) to desired thickness (A lightbrown color indicates a thickness of < 100Å.). 9. Place the paper and coated grids onto a piece of filter paper that is soaked with ethylene dichloride in a covered petri dish. One half hour should be sufficient time to dissolve the Formvar film and not damage the carbon support. Remove the grids and paper and allow them to dry in a dust-free area. C. Use of the Ladd shadow evaporator for carbon coating plastic films 1. Turn shadow evaporator on: Turn both the main and mechanical pump switches on. Move the black-knobbed, manifold valve handle downwards to "backing" position. Open the air inlet valve and CAREFULLY remove the implosion shield and bell jar. Set the bell jar upside down on the rest on the adjacent cabinet 2. Set up carbon coating apparatus: Plug one lead to ground ("E") and the other to "1" (See lower diagram). Remove the cylindrical glass shield. Release the tension spring that holds the right carbon rod in place and remove the rod. File the edge of the left carbon rod flat with a piece of emery cloth. Replace the right rod with a fresh one or sharpen it by the procedure described below. 3. Carbon rod sharpening procedure: Place the carbon rod in the chuck of the sharpener. Pull the rod out until its edge is aligned with the edge of the aligning arm and then tighten the chuck. Turn on the sharpener and run the first sharpener tool against the rod until a conical point is formed. Then run the other sharpener tool against the rod until a narrow point is formed. Turn off the sharpener and clean off all carbon dust. Put the newly sharpened rod in the chuck of the carbon coater and tighten. Replace the tension spring and then the glass shield. 4. Set up grids: Place the grids and paper support on a piece of filter paper on top of the base of the carbon coating apparatus (See diagram below). Place a thumbtack along side the slide. This provides a "shadow" on the filter paper and helps you determine the relative thickness of the carbon coating. 5. Diffusion pump warm up: Replace the bell jar and the implosion shield. Close the air inlet, and move the manifold valve handle slowly upwards to the roughing position. Allow the vacuum to reach 0.04 Torr on the bell jar gauge and then move the handle downwards to backing. IMPORTANT: Turn on the water supply. The water supply-line valve is located on the wall behind the shadow evaporator. Turn on the diffusion pump switch and allow the pump to warm up for 15 minutes before continuing. 6. Obtaining a high vacuum: Move the manifold valve handle slowly upwards to the roughing position and allow the vacuum to reach 0.04 Torr on the bell jar gauge. While

13 waiting for the vacuum to recover, fill the baffle with liquid nitrogen. When the bell jar vacuum has reached 0.04 Torr, move manifold valve handle down to the backing position. Depress the metal guard beneath the red mains valve knob and move the knob handle upwards to the open position Allow the vacuum to reach a minimum of 2x10-5 Torr or better. 7. Carbon coating: Turn the electrode selector to #1. Turn the electrode switch on. Slowly turn the electrode current control knob until there is a slight glow at the point where the two carbon rods meet. Slowly increase the current until the rods become white hot. The proper current setting should be just before the point where the carbon starts to sputter. Frequently monitor the thickness of the carbon by turning down the current, checking the darkening of the filter paper and then turning the current back up again. 8. Diffusion pump cool down: Turn down the electrode current control knob and turn off the electrode switch. Make sure the manifold valve is set to the backing position and close the mains valve. Open the air inlet, remove the implosion shield and bell jar and remove the grids. Then replace the bell jar and implosion shield, close the air inlet and move the manifold valve handle to the roughing position. Allow the vacuum to reach 0.04 Torr on the bell jar gauge, move the manifold valve handle to the backing position, turn off the diffusion pump, and allow the pump to cool for 20 minutes. 9. Turn Shadow Evaporator off: Close the manifold and turn off the mechanical pump and main power switches. Turn off the cooling water. D. Glow discharging carbon films 1. Note: Place the very edge of your carbon coated grids along the edge of a piece of double-sided tape on a glass slide. This will help to prevent your grids from flying around inside the shadow evaporator when the air release switch is opened. 2. Turn shadow evaporator on: Turn the main power switch on, turn on the mechanical pump and move the manifold valve handle (black knob) downwards to the backing position. Open the air inlet. CAREFULLY remove the implosion shield and bell jar. 3. Set up glow discharge unit: Plug the lead into the proper receptacle (bnc connector). Place the glass slide with your grids on the unit and replace the bell jar and implosion shield. Close the air inlet, turn the butterfly switch by the current gauges to glow discharge and move the manifold valve handle slowly upwards to the roughing position. Allow the vacuum to reach Torr on the bell jar gauge. The manifold valve may be turned to the closed position if the vacuum rises above 0.10 Torr. 4. Glow discharging: Turn the electrode selector to position #1 and turn the electrode switch on. Slowly turn up the electrode current until there is a bright purple glow surrounding the glow discharge unit. Maintain this setting for approximately 10 seconds

14 while monitoring vacuum. Turn off the electrode current control knob and the electrode switch. Move the manifold valve handle to the backing position. Turn the butterfly switch back to the evaporator setting. 5. Turn shadow evaporator off: Slowly open the air inlet to prevent your grids from being blown around the bell jar. Remove the grids, replace the shields and then close the air inlet. Move the manifold valve to the roughing position. Allow the vacuum to reach 0.04 Torr on the bell jar gauge before moving the manifold valve handle to the horizontal (closed) position. Turn off the mechanical pump and the main switch. Prepared by Ricardo A. Bernal