Preparation Of Ciliated Protozoa For Scanning Electron Microscopy General notes: The same procedures are used to fix and stain cells for SEM and for TEM. Cells can be fixed using conventional glutaraldehyde-osmium fixation described for transmission electron microscopy. To preserve ciliary orientation, use the "instant fixation" protocol described here. With this method, the cortex and ciliary beat form is well preserved but the cytoplasm is poorly preserved and membrane breakage and blebbing on the cell surface is evident.
To suspend cells, use a wide bore pipette and gentle flow to avoid damaging cells. Polypropylene disposable Pasteur pipettes work very well for all procedures.
Carry out all fixation procedures in a properly vented fume hood.
Supplies and most EM chemicals can be obtained from Electron Microscope Sciences, Ernest F. Fullum, Ladd Research Industries, Polysciences, Ted Pella, and Bio-Rad.
- Polypropylene conical centrifuge tube (Falcon 35-2097 or equivalent)
Ice in ice bucket
- 200 mM stock solution of sodium cacodylate buffer, pH 7.2
2% aqueous uranyl acetate (allow a few hours to dissolve)
Deionized water Glutaraldehyde - 10% or 25% stocks in sealed ampoules
Osmium tetroxide - 4% solution in deionized water.
To prepare, score and break an ampoule containing crystalline OsO4 and drop the open ampoule into a small amber jar. Add appropriate amount of deionized water and let dissolve at room temperature for several days. Store in a second jar at 4 degrees C. OsO4 tends to evaporate and blacken surfaces, so the second jar is highly recommended. It is best to use a separate refrigerator for fixatives. OsO4 solutions also can be frozen in small aliquots and stored at -20 degrees C.
Fix: 2.5% glutaraldehyde, 2% OsO4, 100 mM sodium cacodylate, pH 7.2
Mix glutaraldehyde and cacodylate. Add OsO4 and mix immediately before adding to cells.
- Harvest cells in a clinical or appropriate low speed centrifuge. Centrifuge to obtain a loose pellet of cells (setting #3, 5 min with a clinical centrifuge).
- (Optional, but produces cleaner cell surface) Gently pellet cells from the culture medium and suspend in room temperature HNMK ( 50 mM HEPES, pH 6.9, 36 mM NaCl, 0.1 mM Mg acetate, 1 mM KCl) at a concentration of 106-108 cells/ml. Put in a shallow container and gently agitate for 10-30 min at room temperature to aerate cells.
- Decant most of the medium/HNMK and suspend cells in a slurry (<1 ml of remaining medium or HNMK). Transfer to a 16 ml polypropylene conical centrifuge tube.
- Rapidly add the freshly prepared glutaraldehyde-OsO4, mixture, mix well, and immediately put the tube in ice. Fix for 30-60 min. If the solution turns black, pellet the cells, remove the fix, and add a fresh aliquot of a freshly prepared mixture of glutaraldehyde-OsO4.
- Wash cells at least 3 times with deionized water at room temperature. If necessary, gently pellet cells during each wash.
- Suspend cells in 0.5-1% aqueous uranyl acetate and leave at room temperature for at least 1 hr. Cells can be stored for weeks in uranyl acetate.
- Proceed to ethanol dehydration and critical point drying (SEM) or acetone dehydration and embedding (TEM), below.
Critical point drying We use one of two methods.
Method 1: This is preferred but requires microporous specimen capsules (# 4620; Ted Pella, Inc., Redding, CA), which cost ~$2.00 each.
- Dehydrate uranyl acetate-treated cells in a series of ethanol-water washes (25%, 50%, 75%, two 95%, and three 100% ethanol). If possible, use ethanol dried over molecular sieve pellets (Sigma Chemical Corp. #M2260). Dehydrate in polypropylene conical centrifuge tubes and let cells settle between washes.
- Place specimen capsule in a shallow dish (ex: glass petri dish) containing 100% dry ethanol and transfer dehydrated cells into the capsule. Do not fill the capsule more than about one third full with cells or they will cake during the drying procedure. Firmly fit the lid to the capsule and transfer to the critical point dryer.
If more than one sample capsule will be dried at the same time, label the outside of each capsule with pencil and/or insert a tiny piece of paper (marked with pencil) into the capsule before capping.
- Critical point dry cells using a critical point drying apparatus and CO2.
- Dust the dried cells on a SEM specimen stub covered with a piece of double stick tape. Spreading is easier if cells are brushed on the tape with a fine camel hair brush while observing spreading with a dissecting microscope.
- Coat specimens on the stubs with gold/palladium using a sputter coater.
- Store specimens in a dry container.
Method 2: This method is simple but requires that cells or organelles stick to a coated glass coverslip. Many cells are lost during the dehydration procedure but, usually, more than enough cells will stick for SEM observation.
- Place a 12 mm circular coverslip on a piece of Parafilm or in a small plastic or glass petri dish.
- Apply a drop of an 0.5% aqueous solution of polyethylenimine (Sigma P-3143) to the coverslip. Let sit for 10-20 min.
- Remove the drop of polyethylenimine and rinse with one or two drops of deionized water.
- Put a drop of fixed cells in water or uranyl acetate on the coverslip and let settle for 5-10 min.
- Dehydrate samples with a series of ethanol-water washes (25%, 50%, 75%, two 95%, and three 100% ethanol). For the last, washes, use ethanol dried by molecular sieve pellets (Sigma Chemical Corporation, Cat #M 2260). Many cells will detach from the coverslips during the dehydration procedure. This can be minimized if solutions are removed with a pipette held vertically over the specimens.
- Dry cells using a critical point drying apparatus and CO2. Use a cover-slip holder designed for the dryer or prop the coverslips on a plastic separator removed from a typical microscope slide box.
- Remove the coverslips from the dryer and attach them to SEM specimen stubs using double stick tape or adhesive tabs (#76760 from Electron Microscopy Sciences).
- Coat specimens with gold/palladium (60-40) using a sputter coater.
- Store specimens in a dry container.