Electron Microscopy

        Electron microscopy is a type of microscopy that uses a beam of electrons to create an image of the specimen. It is capable of much higher magnifications and has a greater resolving power than light microscopy, allowing it to see much smaller objects in finer detail. Sample preparation prior to electron microscopy is important for obtaining good results. 

     A. Reagents

  • 0.2M PB (pH 7.4): 21.8 g Na2HPO4, 6.4 g NaH2PO4, 1000ml ddH20
  • 4F1G Fixative (4% Formaldehyde & 1% Glutaraldehyde in 0.1 M PB, pH 7.4): 88 ml 0.1M PB (pH 7.4), 10 ml 37-40% Formaldehyde, 2 ml 50% Glutaraldehyde
  • 0.2M (8%) Sucrose in 0.1M PB: 8 g Sucrose, 100ml 0.1 M PB
  • 1% Osmium in 0.1M PB: 5ml 2% Osmium, 5ml 0.2M PB (pH7.4)
  • 5% Uranyl Acetate Solution (50 ml):  2.5 g uranyl acetate in 50 ml ddH20. Cover with foil and stir overnight.  Add 10 drops of glacial acetic acid. Store solution at 4 oC. [Solution is stable for 6 months at 4 oC]
  • Reynold’s Lead Citrate Solution (50 ml) [Add components in the order stated]: 1.33 g Lead nitrate, 1.76 g Sodium citrate, dihydrate (solution becomes cloudy), 5 ml 1N NaOH (solution becomes clear), 30 ml ddH20. Stir for 10 minutes to dissolve and add 15 ml ddH20. Can be stored for 3-6 months at 4 oC.
  • EMBed 812 (20.91 ml): 10 ml EMBed-812, 4.5 ml DDSA, 6 ml NMA, 0.41 ml DMP-30 (2%). Mix components together in a plastic beaker and stir with wood stick. Can be scaled up accordingly.

     B. Staining protocol for tissue samples

         Fixation 

  1. Fix 1 mm tissue blocks in 4% formaldehyde and 1% glutaraldehyde in 0.1 M PB (pH 7.4) for at least 2 hours to overnight.
  2. Immerse in 8% (0.2 M) sucrose in 0.1 M PB 3x15 min or overnight.
  3. Post-fix in 1% osmium tetroxide in 0.1 M PB 1 hour.

         Dehydration and Embedding

         Dehydrate samples in the following reagents:

  1. 50% Ethanol  15 min
  2. 70% Ethanol  15 min
  3. 95% Ethanol  15 min
  4. 100% Ethanol  2x15 min
  5. 100% Propylene oxide  2x15 min
  6. 1:1 EMBed 812 and Propylene Oxide 1-2 hours.
  7. 2:1 EMBed 812:Propylene Oxide overnight in dessicator with top off.
  8. Embed in beam capsules.
  9. Bake in 60 oC oven for 48 hours.

         Sectioning

  1. Take Semithin/thick sections (0.5-1 um) and stain with toluidine blue for 2-5 min.
  2. Observe sections under microscope for precise location to cut for ultrathin sections.
  3. Make ultrathin sections at 60-90 nm thick (silver-yellow colour) and collect sections onto grids. Dry sections overnight before staining.

         Staining

  1. Stain grids with uranyl acetate for 15 minutes and lead citrate for 3-5 minutes. Rinse with ddH20.
  2. Observe under electron microscope.

     C. Staining protocol for cultured cells

         Fixation 

  1. Fix cell suspension or free cells in 4% formaldehyde and 1% glutaraldehyde in 0.1 M PB (pH 7.4) by mixing equal volumes of fixative and cell suspension.
  2. Transfer cells to centrifuge tube and spin for 10 minutes. Discard fixative (keep pellet of cells) and add fresh fixative for at least 2 hours or overnight.
  3. Remove fixative and replace with 8% (0.2 M) sucrose in 0.1 M PB 3x15min or overnight.
  4. Post-fix in 1% osmium tetroxide in 0.1 M PB 1 hour.
  5. Remove osmium tetroxide and rinse in 0.1 M 3x10 minutes.

         Dehydration and Embedding

         Dehydrate samples in the following reagents:

  1. 50% Ethanol  15 min
  2. 70% Ethanol  15 min
  3. 95% Ethanol  15 min
  4. 100% Ethanol  2x15 min
  5. 100% Propylene oxide  2x15 min
  6. 1:1 EMBed 812 and Propylene Oxide 1-2 hours.
  7. 2:1 EMBed 812:Propylene Oxide overnight in dessicator with top off.
  8. Embed in beam capsules.
  9. Bake in 60 oC oven for 48 hours.

         Sectioning

  1. Take Semithin/thick sections (0.5-1 um) and stain with toluidine blue for 2-5 min.
  2. Observe sections under microscope for precise location to cut for ultrathin sections.
  3. Make ultrathin sections at 60-90 nm thick (silver-yellow colour) and collect sections onto grids. Dry sections overnight before staining.

         Staining

  1. Stain grids with uranyl acetate for 15 minutes and lead citrate for 3-5 minutes. Rinse with ddH20.
  2. Observe under electron microscope.

    

     D. Staining protocol for paraffin sections mounted on slides

         Preparing sections 

  1. Mount Paraffin sections on microscope slides, air dry overnight, and then dry in 50 oC oven for 1 hour. [The thickness of section should at least 5 µm]
  2. Deparaffinise sections by placing slides in three changes of xylene (or substitute), 10 minutes for each.
  3. Re-hydrate sections through a series of alcohols (100% - 3 changes 5 min each, 95% - 1 min and 70% 1 min) to water 

         Re-fixation 

  1. Re-fix sections with 4% formaldehyde and 1% glutaraldehyde in 0.1 M PB (pH 7.2) for 1-2 hours.
  2. Rinse twice in 0.1 M PB and incubate in 8% (0.2M) sucrose in 0.1 M PB for 15 minutes (each rinse) or overnight.
  3. Post-fix in 1% osmium tetroxide in 0.1 M PB for 1 hour. Rinse with 0.1 M PB briefly.

         Dehydration

         Dehydrate samples in the following reagents:

  1. 50% Ethanol  15 min
  2. 70% Ethanol  15 min
  3. 95% Ethanol  15 min
  4. 100% Ethanol  2x15 min
  5. 100% Propylene oxide  2x15 min
  6. 1:1 EMBed 812 and Propylene Oxide 1-2 hours.
  7. 2:1 EMBed 812:Propylene Oxide overnight in dessicator with top off.
  8. Drain off 2:1 EMbed-812:P.O. and place slides on a flat surface. Put a few drops of EMbed-812 on the sections and incubate for 1 hour.
  9. Drain and change fresh EMbed-812 and incubate for 1 hour. 

         Embedding

         Use step 1 or 2, and then 3 to perform embedding. 

  1. Use a flat pre-hardened block that you have previously made in a flat end beam capsule. Place the flat end of the block on top of the section, being careful not to introduce air bubbles.
  2. Place slide on top of an Aclar Embedding Film, make sure the size of the film is larger than the slide. Then cut off another piece of film a little bigger than the slide, place it on top of the slide from one end to the other, being careful not to introduce air bubbles.
  3. Bake in 60 oC oven for 16 hours or overnight. Be sure to place the slide on a flat surface.

         Detaching EMbed block from slide

         Use step 1 or 2, and then 3 to detach the EMbed block. 

  1. Use this step if you used steps 1 & 3 for embedding. Take the slide and Epon block out of oven and let it cool down. Cut a rectangle or circle shape surrounding the EMbed block using a razor blade. Then immerse slide in liquid nitrogen for about 30 seconds to 1 minute, and this will detach the EMbed block together with section from the slide.
  2. Use this step if you used steps 2 & 3 for embedding. Take the slide out of oven and let it cool down. Peel off the Aclar Embedding Film from both sides of the slide. Cut a rectangle or circle shape surrounding the section using razor blade.
  3. Immerse slide in liquid nitrogen for about 30 seconds to 1 minute to detach the section from the slide. [If using the Aclar Embedding Film (step 2), then select a region of interest (ROI) on the “EMbed section”, cut a piece of ROI and glue it using tissue adhesive onto a flat end EMbed block. Air dry for at least 30 minutes].

         Sectioning

  1. Trim the block and cut semithin/thick sections (0.5-1.0 µm), be careful to line up the block surface perfectly and cut only one or two thick sections and no more, or you will lose the entire section if you used 5µm sections.
  2. Transfer the sections to a drop of water on a slide and dry it on slide warmer or lamp until the water drop has gone. Stain with Toluidine blue stain for 2 min and rinse in ddH20. [Adding water drop will flatten sections to avoid wrinkles].
  3. Observe sections under microscope for precise location to cut for ultrathin sections
  4. Make ultrathin sections at 60-90 nm thick and collect sections onto grids. Dry sections overnight before staining.

         Staining

  1. Stain grids with uranyl acetate for 15 minutes and lead citrate for 5 minutes. Rinse with ddH20.
  2. Observe under electron microscope.